University of Veterinary Medicine Hannover Institute of Virology Department of Infectious Diseases

Biological characterization of porcine

THESIS Submitted in partial fulfilment of the requirements for the degree

DOCTOR OF PHILOSOPHY (PhD)

awarded by the University of Veterinary Medicine Hannover

by Johanna Kennedy Koblenz

Hannover, Germany 2020

Supervisor Prof. Dr. Paul Becher

Supervision Group Prof. Dr. Paul Becher

Prof. Dr. Karl-Heinz Waldmann

Prof. Dr. Eike Steinmann

1st Evaluation Prof. Dr. Paul Becher

Institute of Virology, University of Veterinary Medicine Hannover, Germany

Dr. Imke Steffen

Institute for Physiological Chemistry, University of Veterinary Medicine Hannover, Germany

Prof. Dr. Eike Steinmann

Faculty of Medicine, Department of Molecular and Medical Virology, Ruhr-University Bochum, Germany

2nd Evaluation Prof. Benedikt Kaufer, PhD

Institute of Virology, Free University of Berlin, Germany

Date of final exam 30th March 2020

Once again… for science, obviously.

Instant classic.

Parts of the thesis have been published previously in:

Research article

Kennedy J, Pfankuche VM, Hoeltig D, Postel A, Keuling O, Ciurkiewicz M, Baumgärtner W, Becher P, Baechlein C. Genetic variability of porcine pegivirus in pigs from Europe and China and insights into tissue tropism. Sci Rep. 2019 Jun 3;9(1):8174. doi: 10.1038/s41598-019-44642-0

Poster and oral presentations

Kennedy J, Baechlein C, Hoeltig D, Becher P. Presence of porcine pegivirus in domestic pigs and phylogenetic analysis of pegivirus strains from different parts of the world. 10th Graduate school days, 2017, Bad Salzdetfurth, Germany.

Kennedy J, Baechlein C, Hoeltig D, Becher P. Presence of porcine pegivirus in domestic pigs and phylogenetic analysis of pegivirus strains from different countries in Europe and Asia. 28th Annual Meeting of the Society for Virology (GfV), 2018, Würzburg, Germany.

Kennedy J, Pfankuche VM, Hoeltig D, Postel A, Keuling O, Ciurkiewicz M, Baumgärtner W, Becher P, Baechlein C. Porcine pegivirus: genetic variability in pigs from Europe and China, insights into tissue tropism and establishment of antibody ELISA. 11th Graduate school days, 2018, Hannover, Germany.

Kennedy J, Pfankuche VM, Hoeltig D, Postel A, Ciurkiewicz M, Baumgärtner W, Becher P, Baechlein C. Characterization of persistent pegivirus : serology, and replication in PBMCs. 29th Annual Meeting of the Society for Virology (GfV), 2019, Düsseldorf, Germany.

Kennedy J, Pfankuche VM, Hoeltig D, Postel A, Ciurkiewicz M, Baumgärtner W, Becher P, Baechlein C. Insights into porcine pegivirus infection: global distribution, tissue tropism, and transmission. Keystone Symposia Conference on Positive-Strand RNA , 2019, Killarney, Ireland.

Contents

Table of contents ______I

List of abbreviations ______III

List of figures ______VI

List of tables ______VII

Table of contents

1 Introduction ______1

1.1 Genus Pegivirus ______1

1.1.1 Discovery of ______1

1.1.2 Taxonomy ______2

1.1.3 Morphology and organization ______4

1.1.4 Pegivirus protein functions ______5

1.2 Biology of pegivirus infection in pigs and other hosts ______6

1.2.1 Prevalence and seroprevalence ______6

1.2.2 Transmission ______10

1.2.3 Persistence ______10

1.2.4 Tissue tropism ______11

1.2.5 Co-infection with other pathogens and clinical relevance ______12

1.3 Aims of the study ______13

2 Genetic variability of porcine pegivirus in pigs from Europe and China and insights into tissue tropism ______15

I

Contents

3 Dissecting antibody reactivity and possible transmission routes in porcine pegivirus infection ______35

4 Overall Discussion ______59

4.1 PPgV RNA detection in domestic pig serum samples from Europe and Asia _

______59

4.2 Phylogenetic analyses of PPgV ______60

4.3 No detection of PPgV RNA in wild boar ______61

4.4 Persistent and transient PPgV ______62

4.5 Investigation of PPgV tissue tropism ______65

4.6 Insights into PPgV transmission routes ______66

4.7 Antibody reactivity in Western blot and ELISA ______67

5 Summary ______71

6 Zusammenfassung ______73

7 References ______75

II

Contents

List of abbreviations

°C degrees Celcius × g gravitational acceleration µg microgram µl microliter µm micrometer aa amino acid Ab antibody BPgV pegivirus bp base pair cDNA complementary deoxyribonucleic acid Cq cycle quantification CSFV classical swine fever C-terminally carboxyl-terminally E envelope protein E2t carboxyl-terminally truncated envelope protein 2 E. coli Escherichia coli ELISA enzyme-linked immunosorbent assay

ELISA100 ELISA coated with 100 ng protein per well

ELISA250 ELISA coated with 250 ng protein per well EPgV equine pegivirus FISH fluorescence in situ hybridization FPLC fast protein liquid chromatography GBV GB virus h hour HCV C virus HGV hepatitis G virus HHPgV hepegivirus HIV human immunodeficiency virus HPgV human pegivirus HRP horseradish peroxidase IgA immunoglobulin A

III

Contents

IgG immunoglobulin G IgM immunoglobulin M IMAC immobilized metal ion chromatography IPTG Isopropyl β-d-1-thiogalactopyranoside IRES internal ribosome entry site kb kilo base kDa kilo Dalton LB lysogeny broth M molar min minute ml milliliter mM millimolar nm nanometer no. number NS non-structural protein NS3h non-structural protein 3 helicase domain nt nucleotide N-terminal amino-terminal NW New World OD optical density OW Old World ORF open reading frame PAGE polyacrylamide gel electrophoresis PBS phosphate buffered saline PBS-Tw phosphate buffered saline containing 0.05% Tween20 PCR polymerase chain reaction PPgV porcine pegivirus PVDF polyvinylidene difluoride px protein x qRT-PCR quantitative reverse transcription polymerase chain reaction RdRp RNA-dependent RNA polymerase RNA ribonucleic acid RPgV rodent pegivirus

IV

Contents

rpm revolutions per minute RT room temperature RT-PCR reverse transcription polymerase chain reaction SDS sodium dodecyl sulfate SPgV simian pegivirus

SPgVcpz simian pegivirus (chimpanzee) ssRNA single-stranded RNA TBS Tris-buffered saline TDAV Theiler’s disease-associated virus TierSchVersV Tierschutz-Versuchstierverordnung TM TaqMan TMB tetramethylbenzidine UTR untranslated region WB Western blot

V

Contents

List of figures

Chapter 1

Figure 1-1. Phylogenetic relationship of pegivirus species A-K. ______3

Figure 1-2. Predicted genome organization of porcine pegivirus. ______5

Chapter 2

Figure 2-1. Phylogenetic analysis of porcine pegiviruses from different countries and other mammalian pegiviruses. ______21

Figure 2-2. Fluorescence in situ hybridization of porcine pegivirus (PPgV) positive and negative pigs using a PPgV specific probe; overlay phase contrast and immunofluorescence; bar = 100 µm. ______24

Chapter 3

Figure 3-1. Coomassie gel of NS3h protein before and after purification by IMAC. _ 45

Figure 3-2. Western blots of purified NS3h protein incubated with serum samples as first antibody (Ab). ______46

Figure 3-3. Western blot of crude E2t protein incubated with serum samples that showed NS3h-specific antibody (Ab) reactivity in Western blot and ELISA as first Ab.

______48

Figure 3-4. PPgV viral genome quantity in serum (RNA positive results only) during the course of infection in domestic pigs. ______50

VI

Contents

List of tables

Chapter 1

Table 1-1. Pegivirus species nomenclature and their respective hosts (Smith et al.,

2016). ______4

Chapter 2

Table 2-1. Porcine pegivirus genome detection rates and viral genome load in serum samples from individual animals and herds from different countries in Europe and

Asia. ______19

Table 2-2. Number of pegivirus positive pigs of different age groups from Europe and China. ______19

Table 2-3. Porcine pegivirus RNA quantities and fluorescence in situ hybridization results in blood and different tissues from two domestic pigs from Germany. _____ 22

Chapter 3

Table 3-1. Characterization of selected serum samples.______47

VII

Chapter 1

1 Introduction

1.1 Genus Pegivirus

1.1.1 Discovery of pegiviruses

The genus Pegivirus owes its name to the history of the discovery of its first members in 1995 (Simons et al., 1995b, Simons et al., 1995a, Linnen et al., 1996). The name was first proposed by Stapleton and colleagues in 2011, consisting of two parts: “pe” represents the characteristic of the viruses in frequently causing persistent infection in their hosts, and “g” acknowledges the previous names “GB virus” and “hepatitis G virus” (Stapleton et al., 2011). The first virus belonging to this group was discovered in 1995 in and was identified as a primate virus with a flavivirus-like genome closely related to the species virus (HCV) (Simons et al., 1995b). Following this, similar viruses were found in human sera by two independent working groups who tentatively named them GB virus C (GBV-C) and hepatitis G virus (HGV) (Simons et al., 1995a, Linnen et al., 1996). Though the newly discovered viruses were speculated to be the causative agent of hepatitis for some time, studies failed to show a clear association between virus infection and disease, rendering the name “hepatitis G virus” misleading (Alter et al., 1997, Alter, 1997, Simons et al., 1995a, Mohr and

Stapleton, 2009, Theodore and Lemon, 1997). Further pegivirus species were identified in the following years in a variety of mammalian hosts, including further primates, , horses and rodents (Epstein et al., 2010, Chandriani et al., 2013, Kapoor et al.,

2013a, Quan et al., 2013, Firth et al., 2014, Kapoor et al., 2013b). In 2016, pegivirus sequences were discovered in serum samples of domestic pigs from Germany, and the newly discovered virus species was designated as Porcine pegivirus (PPgV) (Baechlein et al., 2016).

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1.1.2 Taxonomy

The taxonomy within the family was recently updated based on phylogenetic relationships and virus characteristics to include newly discovered virus species, which lead to the addition of the genus Pegivirus (Smith et al., 2016, Simmonds et al., 2017, Adams et al., 2013). Highly significant human pathogens belong to this family, including HCV in the genus, and Yellow fever virus, dengue virus and West Nile virus in the Flavivirus genus. Economically important animal pathogens like bovine viral diarrhea virus, classical swine fever virus (CSFV) and border disease virus of sheep belong to the Pestivirus genus (Simmonds et al., 2017). In contrast to the major pathogens found within the first three genera of the Flaviviridae family, none of the pegivirus species have been clearly associated with causing disease in their hosts

(Smith et al., 2016).

The genus Pegivirus currently contains eleven species, Pegivirus A-K, that infect a variety of mammalian hosts, as shown in Figure 1-1 and Table 1-1 (Simons et al., 1995b,

Simons et al., 1995a, Linnen et al., 1996, Epstein et al., 2010, Quan et al., 2013,

Chandriani et al., 2013, Kapoor et al., 2013a, Kapoor et al., 2013b, Firth et al., 2014,

Baechlein et al., 2016). The type species of Pegivirus K is represented by “PPgV_903”, the full-length viral genome sequence of which was isolated from serum of a domestic pig from Germany (Baechlein et al., 2016, Smith et al., 2016).

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Figure 1-1. Phylogenetic relationship of pegivirus species A-K. The amino acid multiple sequence alignment of the complete coding region of the respective viruses was performed with ClustalW in BioEdit 7.0 (Hall, 1999). The Maximum Likelihood phylogenetic tree was calculated with MEGA X (Kumar et al., 2018) using the Le and Gascuel model (Le and Gascuel, 2008) with frequencies and a gamma distribution of variation with invariant sites. Analysis was performed with 100 bootstrap replicates (Felsenstein, 1985) and numbers along branches represent the percentage bootstrap values. The scale bar indicates substitutions per site. GenBank accession numbers are in parentheses.

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Table 1-1. Pegivirus species nomenclature and their respective hosts (Smith et al., 2016).

Species Hosts Previous / other names Abbreviation NW primate SPgV Pegivirus A GB virus A & OW bat BPgV Pegivirus B OW bat GB virus D BPgV human GB virus C HPgV Pegivirus C & OW primate hepatitis G virus SPgVcpz Theiler’s disease-associated Pegivirus D horse TDAV virus Pegivirus E horse equine pegivirus EPgV Pegivirus F NW bat bat pegivirus BPgV Pegivirus G OW bat bat pegivirus BPgV human hepegivirus Pegivirus H human HHPgV human pegivirus 2 Pegivirus I NW bat bat pegivirus BPgV Pegivirus J NW rodent rodent pegivirus RPgV Pegivirus K pig porcine pegivirus PPgV NW, New World; OW, Old World

1.1.3 Morphology and genome organization

Pegiviruses are enveloped viruses with a spherical virion of 60-70 nm in size (Xiang et al., 1999). In contrast to other members of the family Flaviviridae, pegiviruses appear to have a truncated core-coding region or absence thereof, though biochemical characterization and electron microscopy of human pegivirus (HPgV) suggest it has a of uncertain origin (Xiang et al., 1998, Xiang et al., 1999).

The positive-sense, single-stranded RNA genome of pegiviruses is non-segmented and ranges in size from 8.9-11.3 kilo bases (kb). It contains a single open reading frame

(ORF), which is flanked by 5’ and 3’ untranslated regions (UTRs) (Simmonds et al.,

2017). In contrast to the flavivirus 5’ UTR, which contains a type I cap, the 5’ UTRs of , pestiviruses and pegiviruses possess an internal ribosome entry site

(IRES) for translation initiation (Simmonds et al., 2017, Simons et al., 1996). Pegivirus

IRES elements are structurally similar to the type I IRES of picornaviruses, or to the 4

Chapter 1

type IV IRES elements (Pegivirus H, J and F) seen in hepaci- and pestiviruses, though in both cases sequence identity is limited (Quan et al., 2013, Kapoor et al., 2015). As is common in hepaciviruses, BPgVs belonging to Pegivirus F contain a micro RNA-122 binding site in their 5’ UTR, while such sites are lacking in other pegiviruses (Smith et al., 2016). Comparison of PPgV sequences evidences high amino acid (aa) identities indicative of conserved genome regions within the putative non-structural protein 3

(NS3) and non-structural protein (NS) 5B coding regions, which coincides with findings when comparing HPgV and HCV (Baechlein et al., 2016, Leary et al., 1996b).

1.1.4 Pegivirus protein functions

The ORF of members of the Flaviviridae family is translated into a large polyprotein that is co- and post-translationally cleaved by cellular- and viral proteases (Figure 1-2).

As mentioned above, the origin of the pegivirus capsid has not been determined, as a core-coding region like that seen in other Flaviviridae members appears to be lacking in most pegiviruses, including PPgV (Mohr and Stapleton, 2009, Simons et al., 1996,

Baechlein et al., 2016). Most pegivirus protein functions have not been studied in detail and predicted functions are mostly inferred from sequence comparison with homologous proteins within the hepaciviruses, mainly HCV (Simmonds et al., 2017,

Mohr and Stapleton, 2009).

Figure 1-2. Predicted genome organization of porcine pegivirus. Schema was newly constructed for this thesis modified from (Baechlein et al., 2016). Amino acid (aa) sizes of the individual predicted mature proteins are indicated below. The predicted cleavage sites are shown by grey (cellular signal peptidases) and black triangles (viral proteases).

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Chapter 1

Like hepaciviruses, pegiviruses have two predicted envelope glycoproteins (E), E1 and

E2, which appear to be inserted into the viral envelope in the form of heterodimers

(Mohr and Stapleton, 2009). E1 and E2 of PPgV have four N-X-S/T glycosylation sites each, while the number of such sites varies from three to eleven in other pegivirus species (Smith et al., 2016). Protein x (px), the homologue to p7 in HCV, differs in size within the pegivirus species (Mohr and Stapleton, 2009).

Of the predicted six non-structural proteins of pegiviruses, NS2 is thought to be involved in the cleavage of NS2-NS3 as an autoprotease, as it is seen in HCV. The predicted function of HPgV NS3 is that of a viral helicase and of a chymotrypsin-like serine protease, which is responsible for the cleavage of the remaining NS proteins and uses NS4A as a co-factor, a function that is also thought to occur in other pegivirus species (Epstein et al., 2010, Major and Feinstone, 1997, Mohr and Stapleton, 2009,

Moradpour et al., 2007, Robertson, 2001, Leary et al., 1996b, Stapleton, 2003). NS5A appears to be an interferone sensitivity-determining region, and the predicted function of NS5B is that of an RNA-dependent RNA polymerase (RdRp) (Leary et al., 1996b,

Stapleton, 2003, Linnen et al., 1996, Simons et al., 2000).

1.2 Biology of pegivirus infection in pigs and other hosts

1.2.1 Prevalence and seroprevalence

Porcine pegivirus RNA detection methods and rates

PPgV was first discovered in serum of domestic pigs from Germany using high- throughput sequencing methods. It was subsequently detected in 10 of 455 (2.2%) serum samples from 10 of 37 (27.0%) pig holdings by SYBR-Green-based quantitative reverse transcription polymerase chain reaction (qRT-PCR) targeting NS3 (Baechlein et al., 2016). Following this, Yang and colleagues used conventional RT-PCR, which targets a conserved region in NS5B, to screen 159 porcine serum samples from 15 US

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Chapter 1

states, of which 24 (15.1%, from 10 US states) were PPgV RNA positive, evidencing a much higher detection rate than that previously found in Germany (Yang et al., 2018).

A study from China reported the detection of PPgV genome in 34 of 469 (7.3%) porcine sera using nested RT-PCR targeting NS3, and observed an ascending detection rate from suckling piglets (1.6%) to nursing piglets (1.9%), finishing pigs (6.6%), and sows

(11.3%) (Lei et al., 2019).

Human pegivirus RNA and antibody prevalence

HPgV (Pegivirus C) is distributed globally and an estimated 750 million people are viremic, while another 750 million to 2 billion people have evidence of prior HPgV infection. It is thus possibly the most prevalent human RNA virus causing persistent infection, and a major contributor to the human virome (Stapleton et al., 2014,

Stapleton, 2003, Chivero and Stapleton, 2015).

Antibodies (Abs) against HPgV are usually detected after clearance of , thus exposure rates are calculated as the sum of RNA positive and Ab positive rates (Tacke et al., 1997, Gutierrez et al., 1997, Thomas et al., 1998). The prevalence of HPgV viremia in healthy blood donors from developed countries is 1-5%, and another 5-20% of individuals have anti-E2 Abs, leading to a total exposure rate between 6 and 25%

(Mohr and Stapleton, 2009, Stapleton et al., 2011, Blair et al., 1998, Gutierrez et al., 1997,

Pilot-Matias et al., 1996a, Tacke et al., 1997). Rates are higher in blood donors from developing countries, reaching close to 20% RNA detection rate in some regions (Mohr and Stapleton, 2009, Polgreen et al., 2003). However, prevalence of viremia is significantly higher in high-risk groups, namely individuals with coexistent blood- borne or sexually transmitted infections, and nearly universal exposure is demonstrated in some populations, such as intravenous drug users and human immunodeficiency virus (HIV)-positive men who have sex with men (Alter, 1997,

Theodore and Lemon, 1997, Stapleton, 2003, Scallan et al., 1998, Stapleton et al., 2011,

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Chapter 1

Williams et al., 2004, Dawson et al., 1996, Gutierrez et al., 1997, Schlauder et al., 1995,

Xiang et al., 2001).

Recently, Pegivirus H, a further pegivirus species infecting , was identified

(Berg et al., 2015, Kapoor et al., 2015). Prevalence of human hepegivirus (HHPgV, also previously named human pegivirus 2) infection is much lower than that of HPgV, with evidence of exposure to HHPgV (RNA and Ab) detected in 0.45-1.33% of cases (Berg et al., 2015, Coller et al., 2016, Kapoor et al., 2015). In contrast to HPgV, Abs against

HHPgV were frequently detected during viremia (Berg et al., 2015, Coller et al., 2016).

Characteristics and detection methods of HPgV-specific antibodies

While various HPgV proteins have been used for Ab detection by expression in

Escherichia coli (E. coli), mammalian expressed, C-terminally truncated E2 was identified as a useful antigen for studying HPgV exposure (Dawson et al., 1996, Pilot-

Matias et al., 1996b, Pilot-Matias et al., 1996a, Dille et al., 1997).

Anti-HPgV Ab development is usually restricted to conformation-dependent anti-E2

Abs that develop after clearance of viremia, as described above (Gutierrez et al., 1997,

Tacke et al., 1997, Tanaka et al., 1998, Thomas et al., 1998). Anti-E2 antibodies are long- lived and provide a certain degree of protection from reinfection, indicating neutralizing activity (Tillmann et al., 1998, Elkayam et al., 1999, Gutierrez et al., 1997).

Additionally, some studies have described the detection of anti-HPgV peptide reactivity during viremia; however, Ab development is restricted to E2 in most cases, suggesting that the E2 antigenic site is immunodominant in humans (McLinden et al.,

2006, Pilot-Matias et al., 1996b, Fernandez-Vidal et al., 2007, Gomara et al., 2010,

Schwarze-Zander et al., 2006, Tan et al., 1999, Van der Bij et al., 2005, Xiang et al., 1998).

Moreover, HHPgV-specific Abs have been detected using mammalian expressed E2, as well as bacterially expressed NS4A/B and additional peptides located to NS3,

NS4A/B and NS5B (Berg et al., 2015, Coller et al., 2016).

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Pegivirus RNA and antibody detection rates in other mammals

Equine pegivirus (EPgV, Pegivirus E) RNA has been detected in horse serum samples from the United States (9.5%), Brazil (0.8-14.2%), China (1.1%), Germany (13.4%) and

England, Scotland and France (3.6%) (Kapoor et al., 2013a, de Souza et al., 2015,

Figueiredo et al., 2019, Lu et al., 2018, Lyons et al., 2014, Postel et al., 2016). Theiler’s disease-associated virus (TDAV, Pegivirus D) was first detected in 16 horses from the

United States: one was the donor horse of TDAV-containing antitoxin serum and the other 15 horses had been exposed to this serum (Chandriani et al., 2013). Since then

TDAV has only been found in horses from Brazil in one study (1.6%), while two other studies failed to find TDAV RNA in 114 and 177 samples from Brazil and China, respectively (Figueiredo et al., 2019, de Souza et al., 2015, Lu et al., 2016).

For detection of anti-EPgV Abs, the NS3 helicase domain (NS3h) was expressed in E. coli and crude NS3h-containing bacterial lysates were used in indirect ELISA. Of 328 horses from Scotland, England and France, 218 (66.5%) were positive for Ab in ELISA and 88.5% of those were confirmed by Western blot with the same protein. Among these, of the 12 horses with active EPgV infection (3.7%), 10 were also Ab positive

(Lyons et al., 2014).

BPgV was initially detected in 5% (5 of 98) of free ranging bats from Bangladesh

(Epstein et al., 2010), and further BPgV species were identified in 4% of 1,615 animals belonging to 21 species of New World and Old World bats (Quan et al., 2013). Simian pegiviruses (SPgV) have been found in various Sanguinus, Callithrix, and Aotus species

(Bukh and Apgar, 1997, Leary et al., 1996a, Muerhoff et al., 1995, Simons et al., 1995b).

In addition, viruses belonging to Pegivirus C naturally infect chimpanzees (SPgVcpz), sequences of which form a separate phylogenetic group to HPgVs (see also Figure 1-1)

(Adams et al., 1998, Birkenmeyer et al., 1998).

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1.2.2 Transmission

HPgV has been shown to be transmitted by exposure to infected blood, sexually, and vertically from mother to child (Bhanich Supapol et al., 2009, Hino et al., 1998,

Kleinman, 2001, Lin et al., 1998, Ohto et al., 2000, Stapleton, 2003). These transmission routes are largely comparable with those found in HIV, explaining common HIV-

HPgV co-infection, and high-risk groups (i.e. intravenous drug users), as mentioned above (Stapleton, 2003). Sexual transmission of HPgV is much more efficient than that of HCV (Lauer and Walker, 2001, Sarrazin et al., 2000, Scallan et al., 1998, Nerurkar et al., 1998, Bourlet et al., 1999, Hollingsworth et al., 1998, Xiang et al., 2001).

TDAV was shown to be transmitted to healthy horses by experimental intravenous inoculation of antitoxin horse serum containing TDAV (Chandriani et al., 2013). EPgV and TDAV were detected in commercially available equine serum pools from various countries, indicating that transmission by products containing equine serum may be possible (Postel et al., 2016).

The transmission of bat pegiviruses has not been studied in detail, however, BPgV genome was detectable in saliva and rectal swabs, but not in urine of bats, indicating that horizontal transmission by shedding in excreta or during fighting, grooming, or sharing of food may be possible (Epstein et al., 2010, Quan et al., 2013). Studies on the transmission of PPgV between pigs have not been reported to date.

1.2.3 Persistence

As RNA viruses, HPgV and HCV are unusual in frequently causing persistent infections in immunocompetent hosts (Chivero and Stapleton, 2015). Though HPgV persistence is not as frequent as that seen in HCV, it is found in roughly 25% of cases, while the other 75% of infections are cleared within two years (Gutierrez et al., 1997,

Tanaka et al., 1998). HPgV infection can be long-lived – it was documented for 16 years in one individual (Masuko et al., 1996) – and during persistence the viral load usually

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remains constant (Lefrere et al., 1999). Many aspects of immune evasion by RNA viruses, including hepaciviruses and pegiviruses, remain poorly understood (Chivero and Stapleton, 2015). Hypervariable regions of the E2 protein, mutation of immunodominant T-cell epitopes, and chronic stimulation of T-cells leading to T-cell exhaustion are mechanisms that have been proposed for HCV immune evasion, but apparently do not apply to HPgV (Keck et al., 2009, Burke and Cox, 2010, Mohr and

Stapleton, 2009, Lemon, 2010, Stapleton et al., 2012). Persistence was also observed in

HHPgV infection (Kapoor et al., 2015, Berg et al., 2015), as was life-long SPgV infection

(Simons et al., 1995b). Persistent PPgV infection was observed in three pigs from

Germany, in which viral RNA was detected for 7, 16 and 22 months (Baechlein, 2016), but has not been studied in greater depth.

1.2.4 Tissue tropism

The tissue tropism of most pegivirus species, including PPgV, has not been assessed in detail. Though HPgV was speculated to be a causative agent of hepatitis for some years, clear evidence of a causal association with hepatitis and evidence of replication in liver tissue, such as viral negative-strand RNA as a replication intermediate, were lacking or inconclusive (Chivero and Stapleton, 2015, Fan et al., 1999, Pessoa et al.,

1998, Tucker et al., 2000, Berg et al., 1999, Laskus et al., 1997, Laras et al., 1999).

Compared with HCV, a hepatotropic virus in which viral RNA levels are higher in liver than serum, the opposite is the case for HPgV (Pessoa et al., 1998, Chivero and

Stapleton, 2015, Manns et al., 2017). Rather, HPgV negative-strand RNA has been detected in bone marrow and spleen of infected individuals (Radkowski et al., 2000,

Tucker et al., 2000) and further evidence suggested that HPgV is lymphotropic, which has also been demonstrated to be the case for SPgV (Fogeda et al., 1999, George et al.,

2006, Xiang et al., 2000, Kobayashi et al., 1999, Laskus et al., 1998, Stapleton et al., 2011).

HPgV replicates ex vivo in peripheral blood mononuclear cells (PBMCs) isolated from

HPgV RNA positive individuals (Fogeda et al., 1999, George et al., 2003, Rydze et al.,

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2012). PBMCs can also be infected in vitro using serum-derived HPgV (Chivero et al.,

2014, Xiang et al., 2000). Viral RNA was detected in natural killer cells, monocytes and diverse subsets of T- and B-lymphocytes, including naïve, central memory and effector memory T-cells, leading to the suggestion that HPgV may infect hematopoietic precursor cells, which maintain infection during differentiation (Chivero and

Stapleton, 2015, Chivero et al., 2014). However, the primary permissive cell type(s) for pegiviruses remain unknown (Chivero and Stapleton, 2015).

1.2.5 Co-infection with other pathogens and clinical relevance

Co-infection of HPgV and HIV in humans

As mentioned above, due to high similarity in transmission routes, HPgV and HIV co- infection is common (Stapleton, 2003). Several studies have shown beneficial effects on the outcome of HIV-infection attributed to HPgV, including longer survival in co- infected individuals (Heringlake et al., 1998, Tillmann et al., 2001, Nunnari et al., 2003,

Toyoda et al., 1998, Williams et al., 2004, Xiang et al., 2001, Zhang et al., 2006, Vahidnia et al., 2012). Such effects caused by HPgV can be explained by various mechanisms, including alterations of cytokine profile, modification of HIV co-receptor expression, direct inhibition of HIV entry through HPgV E2, and modulation of T-cell activation, among others (Schwarze-Zander et al., 2012, Nunnari et al., 2003, Capobianchi et al.,

2006, Chang et al., 2007, Haro et al., 2011, Herrera et al., 2009, Herrera et al., 2010, Jung et al., 2007, Koedel et al., 2011, Mohr et al., 2010).

Clinical relevance of pegivirus infection in humans

Several studies have suggested an association between HPgV infection and an increased risk of non- (NHL), attributing a possible etiologic role in the development of NHL to chronic immune stimulation or impaired immunosurveillance, to HPgV (Chang et al., 2014, Civardi et al., 1998, Collier et al.,

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1999, De Renzo et al., 2002, Giannoulis et al., 2004, Kaya et al., 2002, Minton et al., 1998,

Zignego et al., 1997, Ellenrieder et al., 1998, Keresztes et al., 2003, Michaelis et al., 2003,

Nakamura et al., 1997, Krajden et al., 2010). Furthermore, HPgV RNA has been detected in brain tissue and in cerebrospinal fluid of individuals with encephalitis of unknown cause, although no causal association has been shown (Kriesel et al., 2012,

Fridholm et al., 2016, Balcom et al., 2018, Tuddenham et al., 2019).

Disease association in animals

TDAV was suggested as the causative agent of a Theiler’s disease outbreak in horses in the United States, but recent studies indicate that a newly discovered member of the copiparvoviruses, namely equine parvovirus hepatitis, is responsible for acute serum hepatitis in horses (Chandriani et al., 2013, Divers et al., 2018).

To date, infection with PPgV has not been shown to cause any disease in pigs, and viral RNA can be detected in apparently healthy animals (Baechlein et al., 2016). One study detected PPgV in a serum sample from a farm with pigs exhibiting lameness and vesicles in the United States, but porcine parvovirus and were also detected. In the same study, nine vesicular swab samples that were additionally tested for PPgV presence were found negative (Yang et al., 2018).

Disease association is similarly unknown for other pegiviruses and they are considered as apathogenic to date. Further research is necessary to better understand clinical implications of pegivirus infection in animal hosts and humans.

1.3 Aims of the study

Since the first description of PPgV in domestic pigs from Germany in 2016, only a handful of studies have been published, which described mainly PPgV genome detection in pigs from the United States and China (Baechlein et al., 2016, Chen et al.,

2019, Lei et al., 2019, Li et al., 2019, Yang et al., 2018). Many aspects of pegivirus

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infection, even in human hosts, remain elusive, not least due to poor in vitro replication and the lack of an animal model in the case of HPgV (Chivero and Stapleton, 2015).

The overall objective of this project was to further biologically characterize porcine pegivirus, specifically including method development to allow insights into RNA and antibody prevalence, tissue tropism and sites of viral replication, host range, transmission routes, and antibody response. In addition to method development, the acquisition of blood, tissue and excretion samples from PPgV-infected pigs was an essential step in the furtherance of this project.

The first aim of the project was the establishment and validation of a TaqMan-based qRT-PCR and the development of an in vitro-transcribed RNA copy standard to allow accurate quantification of PPgV RNA isolated from serum, tissues and excretion samples. The second objective was the establishment of a nested PCR for the amplification of a genome region suitable for sequencing that permitted subsequent phylogenetic analyses. A further goal was the expression of PPgV proteins, namely of the NS3 helicase domain (NS3h), and of truncated E2 (E2t), to evaluate these as possible markers of past or present PPgV infection and, as such, as possible candidates in antibody detection assays.

The development of these methods allows the investigation of various aspects of PPgV biology, including virus distribution and spread, tissue tropism, transmission routes, and infection dynamics, among others. The understanding of such aspects is essential for examining not only how PPgV may influence porcine health, but also in determining whether PPgV infection in the porcine host may be a valuable asset in the study of HPgV infection in humans.

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2 Genetic variability of porcine pegivirus in pigs from Europe and

China and insights into tissue tropism

Kennedy J, Pfankuche VM, Hoeltig D, Postel A, Keuling O, Ciurkiewicz M,

Baumgärtner W, Becher P, Baechlein C.

This chapter was published in Scientific Reports journal

Kennedy J, Pfankuche VM, Hoeltig D, Postel A, Keuling O, Ciurkiewicz M,

Baumgärtner W, Becher P, Baechlein C. Genetic variability of porcine pegivirus in pigs from Europe and China and insights into tissue tropism. Sci Rep. 2019 Jun 3;9(1):8174. doi: 10.1038/s41598-019-44642-0

Contribution as first author:

Experimental work: Establishment and optimization of TM qRT-PCR for screening of the presence of PPgV RNA in serum and tissue samples, establishment and optimization of sequencing RT-PCR for phylogenetic analyses, sample preparation, genome amplification, submission for sequencing. Evaluation and scientific presentation of the results: Analyses and graphical presentation of qRT-PCR and sequencing data, performing phylogenetic analyses. Scientific writing: preparation of the manuscript, tables and figure (phylogenetic tree).

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Abstract

Pegiviruses belong to the family Flaviviridae and have been found in humans and other mammalian species. To date eleven different pegivirus species (Pegivirus A-K) have been described. However, little is known about the tissue tropism and replication of pegiviruses. In 2016, a so far unknown porcine pegivirus (PPgV, Pegivirus K) was described and persistent infection in the host, similar to human pegivirus, was reported. In this study, qRT-PCR, phylogenetic analyses and fluorescence in situ hybridization (FISH) were implemented to detect and quantify PPgV genome content in serum samples from domestic pigs from Europe and Asia, in tissue and peripheral blood mononuclear cell (PBMC) samples and wild boar serum samples from Germany.

PPgV was detectable in 2.7% of investigated domestic pigs from Europe and China

(viral genome load 2.4 × 102 to 2.0 × 106 PPgV copies/ml), while all wild boar samples were tested negative. Phylogenetic analyses revealed pairwise nucleotide identities

>90% among PPgVs. Finally, PPgV was detected in liver, thymus and PBMCs by qRT-

PCR and FISH, suggesting liver- and lymphotropism. Taken together, this study provides first insights into the tissue tropism of PPgV and shows its distribution and genetic variability in Europe and China.

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Introduction

Pegiviruses comprise a group of positive-sense, single-stranded RNA viruses, with a genome size of 9–13 kb, that were recently classified into eleven species (Pegivirus A-

K) within the genus Pegivirus in the Flaviviridae family1. They can infect humans as well as a range of mammalian species, including primates, bats, rodents, horses and pigs2–9.

While pegiviruses are known to cause persistent infections in humans and horses, their pathogenicity remains largely unknown1,4,10–12. Though a pegivirus was identified in horses with Theiler’s Disease in the USA5, recent studies imply that viruses of the copiparvovirus group are associated with serum hepatitis in horses13,14. Human pegiviruses (HPgV) are distributed globally and viral RNA is present in roughly 750 million people, making it one of the most prevalent human RNA viruses15.

Though HPgV was initially thought to be hepatotropic and a possible agent of Non-

A-E hepatitis, evidence of viral replication in the liver of infected patients is missing or inconclusive16–18. Rather, as HPgV replication has been shown in peripheral blood mononuclear cells (PBMCs) ex vivo for several weeks, the virus appears to be lymphotropic19–21. Additionally, HPgV RNA has been found in serum microvesicles, which have successfully delivered viral RNA to uninfected PBMCs that then supported HPgV replication ex vivo22. Interestingly, pegivirus infection in humans may have a beneficial effect on the outcome of human immunodeficiency virus type 1

(HIV-1) infections in individuals co-infected with both viruses, including reduced retroviral loads, slower progression to AIDS and improved survival rates. These benefits are attributed to immune-modulating effects as well as direct and indirect antagonistic mechanisms of HPgV on HIV-1 infection23.

Porcine pegiviruses (PPgV) were first described in domestic pigs from Germany in

20169. The study reported a PPgV detection rate of 2.2% (10 of 455) in porcine serum samples and described persistent infection for up to 22 months in three pigs that did

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not display any clinical signs of disease. Apart from Germany, presence of PPgV has been investigated in North America, where a recent study revealed a PPgV detection rate of 15.1% (24 of 159 samples) in the USA24. Additionally, a recent study investigated

469 porcine serum samples from China, 34 (7.25%) of which were found PPgV positive.

Samples originated from different age groups and proved an ascending trend in the

PPgV positive rate from suckling piglets (1.61%) and nursing piglets (1.85%) to

finishing pigs (6.56%) and sows (11.34%)25.

In this study we analyzed the presence of PPgV genome in pigs from Europe and Asia.

To clarify whether wild boar might play a role in the epidemiology of PPgV, as seen in infections with, for example, classical swine fever virus26,27, African swine fever virus28 and atypical porcine pestivirus (APPV)29, we also investigated the presence of

PPgV genome in wild boar serum samples from Germany. To date the primary permissive cell type(s) of HPgV and other pegiviruses remain unknown. For this reason, we analyzed the tissue and cell tropism of PPgV through detection and quantification of viral RNA in tissues and PBMCs from PPgV positive pigs using qRT-

PCR and fluorescence in situ hybridization (FISH).

Results

PPgV RNA in serum samples from Europe and Asia. The in vitro transcribed RNA copy standard evidenced a highly efficient qRT-PCR assay that was able to detect ten viral genome copies per reaction at Cq values around 36. PPgV genome was detectable in 47 of 1,736 (2.7%) serum samples from domestic pigs corresponding to 20 out of 132 herds (15.2%) (Table 2-1). Highest detection rates were found in individual animals from Great Britain (10.3%) and in herds from China (58.3%). In the different age groups investigated here, the PPgV positive rates were 1.9% in animals under 4 weeks of age,

1.2% in fattening pigs over 4 weeks of age, 3.4% in sows and boars, and 10.1% in pigs of unknown age and use (Table 2-2). Viral loads varied between 2.4 × 102 and 2.0 × 106

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PPgV RNA copies/ml serum, with an overall average of 3.8 × 105 copies/ml. For individual countries on average, lowest genome loads were detected in Poland (1.9 ×

104 copies/ml) and highest in Italy (7.1 × 105 copies/ml). All 800 wild boar samples were negative for PPgV RNA.

Table 2-1. Porcine pegivirus genome detection rates and viral genome load in serum samples from individual animals and herds from different countries in Europe and Asia 1. 1 PPgV, porcine pegivirus; pos., positive.

Table 2-2. Number of pegivirus positive pigs of different age groups from Europe and China.

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Phylogenetic analyses. Altogether 31 PPgV partial NS3 sequences were obtained from domestic pigs, of which nine were identical to one or more other sequences. In total, ten sequences from Germany, three sequences from Italy, four sequences from Poland, nine sequences from Great Britain and five sequences from China were acquired.

Sequences GER/SA/13, GER/SA/91, and PL/159 were identical to one additional sequence each, while IT/77, GB/16, and GB/23 were identical to two further sequences each. In Germany, Poland, and Italy, all identical sequences originated from samples from the same farms, while herd affiliation was unknown for samples from Great

Britain.

Twenty-two distinct sequences shown here (Figure 2-1) were submitted to GenBank.

They displayed nucleotide sequence identities of >90%. According to phylogenetic analysis, PPgV formed a separate branch in the tree of pegiviruses and viral sequences segregated into two main clusters, one of which contained only sequences from

Europe (Germany, Great Britain and Poland). Within the second main cluster, some branches contained sequences recovered from animals in Europe (GER/NDS/T72 and

IT/77) in close proximity to variants from China (i.e. CN/6/5) and USA (i.e.

33/ND/2017)24.

Overall, the most closely related pegivirus sequence found in other species when compared to PPgV was bat pegivirus sequence PDB-1715 (GenBank KC796088), which had a nucleotide sequence identity of 58.1% with PPgV/GB/30. A human pegivirus type 2 sequence, ABT0070P.US (GenBank KT427411), had the lowest nucleotide identity (47.1%) compared to PPgV sequences. When comparing PPgV sequences with pegivirus sequences originating from horses, nucleotide identities ranged from 53.7% to 55.7%. The sequence identities between PPgV and rodent pegivirus were around

54.0%, while the identities with simian pegiviruses ranged from 50.0% to 55.8%.

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Figure 2-1. Phylogenetic analysis of porcine pegiviruses from different countries and other mammalian pegiviruses. Numbers along branches represent percentage bootstrap values (bootstrap values < 80 % are not given). GenBank accession numbers are in parentheses. Scale bar indicates nucleotide substitutions per site. PPgV sequences are marked with a circle and the circle color indicates the country of origin. Pegivirus species A-K are indicated on the right.

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PPgV RNA in tissue samples. In tissue samples of PPgV positive pigs, PPgV RNA was most abundant in the liver (Table 2-3). Liver samples of animals A and B contained

343.9 and 142.5 viral RNA copies/mg tissue, respectively, while 119.3 copies/mg tissue were found in the liver of animal C using qRT-PCR. Serum samples of these animals contained 2,051.1 copies/µl (animal A), 388.6 copies/µl (animal B) and 157.0 copies/µl

(animal C). PBMCs were only available from animal A and contained 46 copies/µl whole blood used for isolation (Table 2-3).

Table 2-3. Porcine pegivirus RNA quantities and fluorescence in situ hybridization results in blood and different tissues from two domestic pigs from Germany 1. 1 PPgV, porcine pegivirus; FISH, fluorescence in situ hybridization; GE, genome equivalents; n.d., not determined; boldface indicates positive FISH results; *fresh blood was not available.

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FISH was used to investigate the liver, thymus, PBMCs and different lymph nodes of animal A, as well as the liver and thymus of animal B, and respective tissues of negative control pigs. PPgV specific signals were detected in the liver of both PPgV positive pigs (Table 2-3; Figure 2-2). Furthermore, several cells of the medullary and cortical region of the thymus of animals A and B were observed to be virus positive using the PPgV specific probe. Additionally, PBMCs of animal A were found to be virus positive in FISH, while lymph nodes, spleen, tonsils, bone marrow and pancreas of animal A tested virus negative. The non-probe incubation as well as the PPgV PCR- negative pigs showed no detectable positive area in the same tested organs, respectively. During necropsy of animal A, multifocal, mild, subendocardial hemorrhages were present. Histopathology showed a mild, portally accentuated, lymphohistiocytic hepatitis, a mild, diffuse infiltration of eosinophils within the thymus, tonsils and lymph nodes and single multinucleated giant cells within the medullary part of the thymus. Furthermore, lymph nodes revealed a mild, diffuse sinus histiocytosis. A moderate, focal, perivascular, lymphoplasmahistiocytic, partially eosinophilic dermatitis was present at the pinna. Additionally, a mild endocardiosis, a mild, lymphohistiocytic epicarditis and a mild to moderate, focal, follicular, lymphocytic conjunctivitis were observed.

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Figure 2-2. Fluorescence in situ hybridization of porcine pegivirus (PPgV) positive and negative pigs using a PPgV specific probe; overlay phase contrast and immunofluorescence; bar = 100 µm. (A) Single hepatocytes of the liver of a PPgV positive pig showed an intracytoplasmic positive signal for PPgV using a PPgV specific probe, also shown at higher magnification in the insert; arrows: nuclei of hepatocytes surrounded by intracytoplasmic, red, positive signals. (B) The liver of a PPgV negative pig lacked a PPgV specific signal. (C) Within the thymus of a PPgV positive pig, scattered cells showed an intracytoplasmic red positive signal for PPgV, also shown at higher magnification in the insert. (D) Within the thymus of a PPgV negative pig, all cells were negative for PPgV using a PPgV specific probe. (E) Several PBMCs from a PPgV positive pig showed a red positive signal using a PPgV specific probe, also shown at higher magnification in the insert. (F) PBMCs from a PPgV negative pig were negative for PPgV.

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Discussion

The genus Pegivirus has grown in recent years, as new viruses were identified in different hosts. Yet little is known about their pathogenicity and the impact on the host’s immune response. In this study, our aim was to gain detailed insights into the distribution of PPgV in different parts of the world and the genetic diversity of PPgV.

Viral RNA was detected in serum samples from domestic pigs from various European countries and China, with an overall individual detection rate of 2.7%.

Investigation of three different age groups from Europe and Asia showed a lower

PPgV positive rate in younger animals such as piglets (1.9%) and fattening pigs (1.2%) than in adult animals (3.4%). This observation is concordant with the results from a recently published study from China; however, the increase in PPgV positive rate was more prominent there (1.6–11.3%). Focusing on samples from China, we found similar results: 1.0% detection rate in piglets and 9.7% detection rate in sows and boar25.

The PPgV positive rates found in this study differ between countries. While no samples were PPgV positive from Switzerland, Serbia and Taiwan, samples from

Germany, Poland and Italy have a positive rate similar to the one described previously for German domestic pigs (2.2%)9. High detection rates in China (7.8%) and Great

Britain (10.3%) found here are nonetheless lower than the positive rate observed in the

USA (15.1%) in a previous study24. In humans, HPgV prevalence ranges from 0.5 to 5% in healthy blood donors from developed countries, but is higher in blood donors from developing countries (5–18.9%), and in individuals co-infected with blood borne or sexually transmitted diseases, like hepatits C virus or HIV-130–32. Equine pegivirus

(EPgV) has been found in 12 of 328 horses (3.7%) from Europe and 7 of 74 horses (9.5%) from USA4,12, thus showing similar detection rates as PPgV. The divergence in PPgV detection rates suggests uneven distribution of virus infection and local spread of

PPgV. This may be caused by the occurrence of other infectious diseases in pig

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populations, similar to observations in humans with co-infection, and needs to be studied further. The viral loads determined here (2.4 × 102 to 2 × 106 copies/ml) are similar to EPgV RNA loads described in one study, which ranged from 3.2 × 104 to 3.2

× 106 RNA copies/ml4. However, another study found higher EPgV viral loads (4.1 ×

105 to 2.0 × 109 RNA copies/ml)12, and the mean RNA load of HPgV in human plasma typically reaches >1 × 107 copies/ml. This may suggest lower replication of PPgV in vivo compared to HPgV and EPgV.

Although HPgV does not appear to be hepatotropic, high amounts of PPgV RNA in the porcine liver shown by qRT-PCR and in situ techniques suggest that viral RNA may accumulate in the liver or even that PPgV has the ability to replicate in hepatocytes. However, this hypothesis will have to be investigated in future studies, as well as whether PPgV infections might be the cause of histopathological changes in the liver, as seen in animal A. Moreover, presence of PPgV RNA in PBMCs and in the thymus supports lymphotropism analogical to HPgV22. Positive FISH results in primary but not secondary lymphoid organs, such as spleen or lymph nodes, imply that the virus might replicate in the thymus and spread to other tissues (e.g. the liver) via PBMCs, but successfully evades recognition by the , which could lead to a persistent infection in the host. Despite significant amounts of viral RNA detected in cells and tissues, highest viral loads were present in the serum of infected animals. With regard to this, low amounts of PPgV RNA in further organs and tissues can most probably be attributed to blood residues. Possible presence of viral RNA in serum microvesicles and associated virus uptake by PBMCs, as seen for HPgV, remain to be determined22.

Phylogenetic analyses showed close genetic relationships among PPgV sequences from different countries, like sequences GER/NDS/T72 and CN/6/5. This could suggest virus spread by international trade with pigs or pig products, such as feed. While all wild boar samples were tested negative for PPgV RNA in this study, other porcine

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viruses from the family Flaviviridae, such as APPV, have been shown to be present at a higher rate in wild boar (19%) than in domestic pigs from Germany (6.2%)29,33. For

APPV, virus transmission between wild boar and domestic pigs appears likely, as strains originating from wild and domestic animals show genetic distance of as little as 6.6%29. However, due to the comparatively low prevalence of PPgV in Germany, transmission of the virus from domestic pigs to wild boar and vice versa may be limited. Only samples from wild boar hunted in northern Germany entered the present study. Future studies with extended sampling will reveal whether PPgV is also absent in wild boar from other geographical regions. As genome detection alone may result in underestimation of virus dissemination, upcoming investigations of samples from domestic pigs, wild boar and other species will also address serological reactions upon infection with PPgV.

These results manifest that PPgV, like other pegiviruses, is distributed over several continents. It can be hypothesized that putative immune modulatory effects of PPgV infections are implicated in pig health worldwide. Detection of PPgV RNA in lymphoid cells suggests that the virus has the potential to affect the immune system of pigs. First insights into the cell- and organ tropism of PPgV suggest that the virus may be hepatotropic and/or lymphotropic. Future studies will clarify the pathogenic potential and immune modulatory effects of this newly discovered, widely distributed virus.

Methods

1,736 serum samples from domestic pigs from different countries in Europe (Germany,

Great Britain, Poland, Switzerland, Italy and Serbia) and Asia (mainland China and

Taiwan) originating from 132 different herds were analyzed in this study. For samples collected in Great Britain, the number of herds was unknown. Samples included 108 piglets up to four weeks old, 923 fattening pigs over four weeks old, 557 sows and

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boar, and 148 pigs of unknown age and use. Samples were taken between 2014 and

2018, originated from apparently healthy domestic pigs and were taken within the framework of national veterinary health management in concordance with national legal and ethical regulations. Residual volumes of these samples were provided for use in the current study, therefore no ethical approval was required for use of these samples. In addition, 800 serum samples from hunted wild boar from Lower Saxony,

Germany, were included. 456 of these wild boar samples were collected during the hunting seasons of 2015/2016 and 2016/2017 and were used in a previous study investigating APPV prevalence29. 344 additional wild boar samples were collected during the hunting season of 2017/2018. Furthermore, blood and post-mortem tissue samples originated from apparently healthy PPgV positive pigs (n = 3, animals A, B, and C) from the Clinic for Swine, Small Ruminants, Forensic Medicine and

Ambulatory Services (University of Veterinary Medicine, Hannover) and PPgV negative control pigs (n = 2). To rule out presence of co-infections with APPV and porcine reproductive and respiratory syndrome virus (PRRSV), PPgV positive animals were also tested using RT-PCR and found negative for both viruses (data not shown).

One pig (animal A) was submitted to the Department of Pathology, University of

Veterinary Medicine Hannover. A full necropsy was performed and samples of 40 different tissues were collected and stored at −80 °C or fixed in 10% neutral buffered formalin and embedded in paraffin wax. For histopathological examination, 3 µm thick sections were stained with hematoxylin and eosin. Different organ and tissue samples and a liver sample originated from two further PPgV positive pigs, animal B and animal C, respectively. Control samples for FISH were taken from PPgV negative pigs. PBMCs from animal A and one negative control animal were isolated from ~1 ml blood by density gradient centrifugation with Histopaque (Merck, Darmstadt,

Germany). Euthanasia and sampling were approved by Lower Saxony’s official authorities (LAVES AZ 15A602 and 17A195) and were carried out in accordance with

German legislation (TierSchVersV).

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RNA was isolated from 140 µl of serum using the QIAmp Viral RNA Mini Kit

(QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. Isolation of RNA from preweighed tissue samples was achieved using the NucleoSpin RNA kit

(Macherey-Nagel, Düren, Germany) or the RNeasy Mini Kit (QIAGEN) according to the manufacturer’s instructions and RNA samples were stored at −80 °C until testing.

For PPgV genome quantification, a TaqMan based qRT-PCR targeting the highly conserved NS3 encoding region with primers PPgV/ fwd/7

(5′-GTCTATGCTGGTCACGGA-3′), PPgV/rev/8

(5′-CACTCATCGCAAATGACCAC-3′) and probe PPgV/pro/11

(5′-[6FAM]-CCATTTCGCGAACCACTGATTCCA-[BHQ1]-3′) was developed and verified using samples that were PPgV positive in a SYBR Green qRT-PCR (QIAGEN) described in an earlier study9. For the new PCR assay, an in vitro transcribed RNA copy standard was developed using MEGAscript Kit (ThermoFisher Scientific, Germany) to allow for absolute quantification of genome copies. Real-time RT-PCR was performed using the Mx3005P QPCR System (Agilent Technologies, Santa Clara, USA) and the

QuantiTect Probe RT-PCR Kit (QIAGEN) according to the manufacturer’s instructions on samples and RNA standard dilutions. Briefly, 12.5 µl RT-PCR master mix, 0.25 µl reverse transcriptase, 0.8 pmol of each primer and 0.2 pmol of the probe, 5.25 µl water and 5 µl sample RNA were used in each reaction of 25 µl with the following temperature profile: 50 °C for 30 minutes, 95 °C for 15 minutes and 40 cycles of 94 °C for 15 seconds and 60 °C for 1 minute. Serum samples were initially screened in pools containing three to ten individual samples; subsequently samples from positive pools were tested individually.

For phylogenetic analysis, amplicons corresponding to a partial NS3 coding sequence were generated by one of the following two methods: a) RT-PCR with SuperScript III

One-Step RT-PCR System with Platinum TaqDNA Polymerase (Life technologies,

Germany) with primers PPgV/fwd/G1 (5′-CACCGGGCTGTTTCTGCTA-3′) and

PPgV/rev/G4 (5′-TTCCTTCCACACCAACCCAT-3′), or b) cDNA synthesis with 29

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SuperScript II Reverse Transcriptase (Invitrogen, Germany) using random hexamers followed by nested PCR with outer primers PPgV/fwd/G1 and PPgV/rev/G4, and inner primers PPgV/fwd/G3 (5′-CGGGCTGTTTCTGCTAGGT-3′) and PPgV/rev/G2

(5′-CACCAACCCATCGAGGATCA-3′) using Taq polymerase included in the

Maxima Hot Start Green PCR Master Mix (2X) (ThermoFisher Scientific) and the following cycling parameters: 95 °C for 4 min, 40 cycles of 95 °C for 30 s, 52 °C for 30 s, 72 °C for 75 s, and 72 °C for 10 min. PCR products with an expected length of 1,290

(method a) and 1,278 (method b) were purified using the GeneJET PCR Purification

Kit (ThermoFisher Scientific) according to the manufacturer’s instructions and submitted to Sanger sequencing (FlexiRun, LGC Genomics, Germany) with primers

PPgV/fwd/G3 and PPgV/rev/G2. Sequences were trimmed to a final length of 1041 base pairs and a multiple sequence alignment was performed with ClustalW implemented in BioEdit 7.034. Phylogenetic trees were calculated in MEGA7 using the

Maximum-likelihood method and the Kimura 2-parameter substitution model35 with

500 replicates for statistical evaluation.

FISH was performed on formalin-fixed, paraffin-embedded organ sections of two qRT-

PCR positive pigs (animal A and B) and on the PBMC pellet of one pig (animal A) using a PPgV specific RNA probe covering parts of the PPgV NS3. The probe set

(ViewRNA TYPE 1 Probe Set, ThermoFisher Scientific) covered positions 2–816 of a target sequence with 1,172 nucleotides that overlapped with the partial PPgV sequence of animal A (GenBank MH979651). The procedure was carried out according to the manufacturer´s protocol with minor variations as previously described (ViewRNA

TYPE 1 Probe Set; ViewRNA™ ISH Tissue Assay Kit (1-plex) and ViewRNA

Chromogenic Signal Amplification Kit; ThermoFisher Scientific;)36. Briefly, sections were deparaffinized, boiled in pretreatment solution® at 90 °C for 10 minutes, digested by a protease QF® at 40 °C for 10 minutes and afterwards fixed. Hybridization to the specific probe was performed for 6 hours. Following preamplification and amplification steps, sections were stained with Fast Red Substrate and counterstained 30

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with Mayer´s hemalum® (Carl Roth GmbH, Karlsruhe, Germany). Images were acquired with an inverted fluorescence microscope (Olympus IX-70; Olympus Life

Science Europe GmbH, Hamburg, Deutschland). The specificity of the probe was confirmed by including a non-probe incubation which served as system negative control and organ sections and cells of PPgV RT-PCR-negative pigs, respectively.

Accession codes. The obtained DNA sequences were deposited in GenBank (accession numbers: MH979651-MH979672).

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17. Fan, X. et al. Is hepatitis G/GB virus-C virus hepatotropic? Detection of hepatitis G/GB virus-C viral RNA in liver and serum. J Med Virol 58, 160–164 (1999).

18. Laskus, T., Radkowski, M., Wang, L. F., Vargas, H. & Rakela, J. Lack of evidence for hepatitis G virus replication in the livers of patients coinfected with hepatitis C and G viruses. J Virol 71, 7804–7806 (1997).

19. Fogeda, M. et al. In vitro infection of human peripheral blood mononuclear cells by GB virus C/Hepatitis G virus. J Virol 73, 4052–4061 (1999).

20. George, S. L., Xiang, J. & Stapleton, J. T. Clinical isolates of GB virus type C vary in their ability to persist and replicate in peripheral blood mononuclear cell cultures. Virology 316, 191–201 (2003).

21. Rydze, R. T., Bhattarai, N. & Stapleton, J. T. GB virus C infection is associated with a reduced rate of reactivation of latent HIV and protection against activation-induced T-cell death. Antivir Ther 17, 1271–1279, https://doi.org/10.3851/IMP2309 (2012).

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22. Chivero, E. T. et al. Human pegivirus RNA is found in multiple blood mononuclear cells in vivo and serum-derived viral RNAcontaining particles are infectious in vitro. J Gen Virol 95, 1307–1319, https://doi.org/10.1099/vir.0.063016-0 (2014).

23. Schwarze-Zander, C., Blackard, J. T. & Rockstroh, J. K. Role of GB virus C in modulating HIV disease. Expert Rev Anti Infect Ther 10, 563–572, https://doi.org/10.1586/eri.12.37 (2012).

24. Yang, C. et al. Detection and genetic characterization of porcine pegivirus in pigs in the United States. Transbound Emerg Dis, https://doi.org/10.1111/tbed.12844 (2018).

25. Lei, D. et al. Detection and genetic characterization of porcine pegivirus from pigs in China. Virus Genes 55, 248–252, https://doi. org/10.1007/s11262-018-1624-6 (2019).

26. Moennig, V. The control of classical swine fever in wild boar. Front Microbiol 6, 1211, https://doi.org/10.3389/fmicb.2015.01211 (2015).

27. Postel, A., Austermann-Busch, S., Petrov, A., Moennig, V. & Becher, P. Epidemiology, diagnosis and control of classical swine fever: Recent developments and future challenges. Transbound Emerg Dis 65(Suppl 1), 248–261, https://doi.org/10.1111/tbed.12676 (2018).

28. Guinat, C. et al. Transmission routes of African swine fever virus to domestic pigs: current knowledge and future research directions. Vet Rec 178, 262–267, https://doi.org/10.1136/vr.103593 (2016).

29. Cagatay, G. N. et al. Frequent infection of wild boar with atypical porcine pestivirus (APPV). Transbound Emerg Dis, https://doi. org/10.1111/tbed.12854 (2018).

30. Mohr, E. L. & Stapleton, J. T. GB virus type C interactions with HIV: the role of envelope glycoproteins. J Viral Hepat 16, 757–768, https://doi.org/10.1111/j.1365-2893.2009.01194.x (2009).

31. Alter, H. J. G-pers creepers, where’d you get those papers? A reassessment of the literature on the hepatitis G virus. Transfusion 37, 569–572 (1997).

32. Stapleton, J. T. GB virus type C/Hepatitis G virus. Semin Liver Dis 23, 137–148, https://doi.org/10.1055/s-2003-39943 (2003).

33. Postel, A. et al. High Abundance and Genetic Variability of Atypical Porcine Pestivirus in Pigs from Europe and Asia. Emerg Infect Dis 23, 2104–2107, https://doi.org/10.3201/eid2312.170951 (2017).

34. Hall, T. A. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp Ser (Oxf), 95–98 (1999).

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36. Postel, A. et al. Presence of atypical porcine pestivirus (APPV) genomes in newborn piglets correlates with congenital tremor. Sci Rep 6, 27735, https://doi.org/10.1038/srep27735 (2016).

Acknowledgements

We thank Vesna Milićević (Belgrade, Serbia), Ming-Chung Deng and Chia-Yi Chang

(New Taipei City, Taiwan), Serum Bank of the Federal Food Safety and Veterinary

Office (Bern, Switzerland), Animal and Plant Health Agency (Weybridge, UK),

Francesco Feliziani and Gian Mario De Mia (Perugia, Italy), Katarzyna Podgórska and

Katarzyna Stępniewska (Puławy, Poland), Hua-Ji Qiu and Yuan Sun (Harbin, China),

Sarah Derking and Jörg Tenhündfeld (Vreden, Germany) and Thomas Große Beilage

(Essen/Oldenburg, Germany) for providing porcine serum samples. We highly appreciate the help of Inga Grotha concerning sample database set-up and RNA preparation. This study was supported by the German Center for Infection Research

(DZIF), Thematic Translational Unit “Emerging Infections”, grant number 8002801801 as well as by the European Union’s Horizon 2020 research and innovation program under grant agreement No. 643476 (COMPARE) and the Deutsche

Forschungsgemeinschaft (DFG, German Research Foundation) – 398066876/GRK

2485/1. This publication was further supported by the Deutsche

Forschungsgemeinschaft and University of Veterinary Medicine Hannover,

Foundation within the funding programme “Open Access Publishing”.

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3 Dissecting antibody reactivity and possible transmission routes in

porcine pegivirus infection

Kennedy J, Hoeltig D, Becher P, Baechlein C.

This chapter is a manuscript in preparation.

Contribution as first author:

Experimental work: Sample preparation of serum and excretion samples, RNA isolation, screening for PPgV RNA presence by TM qRT-PCR, RT-PCR and sequencing of PPgV in serum samples, cloning of plasmids and bacterial expression of PPgV proteins, establishment of purification protocol for PPgV NS3h protein, establishment of SDS-PAGE and Western blot protocols for testing serum sample reactivity with

PPgV proteins, screening of serum samples for PPgV-specific antibodies by ELISA.

Evaluation and scientific presentation of the results: Analyses and graphical presentation of qRT-PCR results, presentation of Western blot and antibody-ELISA results. Scientific writing: preparation of the manuscript, tables and figures.

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Abstract

Porcine pegivirus (PPgV) is a positive-sense ssRNA virus belonging to the Pegivirus genus within the Flaviviridae family that can cause persistent infections in its host. The association of PPgV infection with disease remains unknown, but viral genome has been detected in up to 15.1% of pigs from the United States, China and Europe. Thus far, there are no reports on the virus transmission routes and assays for the detection of PPgV-specific antibodies (Abs) are not available. Therefore, this study investigated serum and excretion samples from PPgV viremic pigs to determine possible virus shedding. Additionally, we expressed PPgV non-structural protein 3 helicase domain

(NS3h) and C-terminally truncated E2 (E2t) intracellularly in E. coli for use in Ab detection. Purified NS3h showed reactivity with porcine serum in Western blot (WB) and indirect enzyme-linked immunosorbent assay (ELISA), and crude E2t (from bacterial inclusion bodies) was reactive in WB. Thus both, NS3h and E2t, appear to be immunogenic and are viable candidates for the further validation of ELISA methods.

Detection of PPgV RNA in serum and excretion samples of PPgV viremic pigs and their piglets revealed one piglet born of a sow, which cleared viremia by day 69 of gestation, was positive in serum directly after birth (before colostrum intake), indicating intrauterine infection. Lack of PPgV genome detection in excretion samples suggested that PPgV may rather be transmitted horizontally via the blood-borne route.

This study reports the first methodological steps towards investigating the immune response induced by PPgV and provides insights into possible viral transmission routes.

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Introduction

Porcine pegivirus (PPgV) was first discovered in 2016 in apparently healthy domestic pigs from Germany (Baechlein et al., 2016). It belongs to the genus Pegivirus, which comprises a group of positive-sense, single-stranded RNA viruses within the

Flaviviridae family (Smith et al., 2016). Persistent pegivirus infections have been found in humans and horses and were also present in three pigs from Germany, in which viral RNA was detected for up to 22 months (Berg et al., 1999, Tanaka et al., 1998,

Kapoor et al., 2013a, Baechlein et al., 2016). As in other pegiviruses, the pathogenicity of PPgV is so far unknown. Although PPgV RNA has been found in porcine serum samples from a farm with pigs exhibiting vesicular disease and lameness, there is no clear association between PPgV infection and disease, and most frequently the virus has been detected in apparently healthy pigs with no clinical signs of viral infection

(Yang et al., 2018, Baechlein et al., 2016, Kennedy et al., 2019).

PPgV is widely distributed and has been detected in domestic pigs from North

America, China, and different countries in Europe (Yang et al., 2018, Lei et al., 2019,

Chen et al., 2019, Baechlein et al., 2016, Kennedy et al., 2019). The highest RNA detection rate was reported in 11 of 67 (16.4%) clinical serum or tissue samples from

China (Chen et al., 2019). However, reports on PPgV-specific antibody (Ab) detection are lacking and viral genome detection alone may result in the underestimation of the abundance of PPgV occurrence in domestic pig herds and further potential hosts.

Ab detection is an important diagnostic tool for HPgV, as viremia and envelope protein 2 (E2) Ab response are mutually exclusive markers of infection in most individuals (Stapleton et al., 2011). Because of this, investigations on the exposure rate of HPgV require both E2-Ab and RNA detection (Gutierrez et al., 1997, Thomas et al.,

1998). Different serological assays have been established by expression of full-length or C-terminally truncated E2 protein in E. coli or mammalian cells (Chinese hamster

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ovary (CHO) and Baby hamster kidney (BHK-21) cells) (Dawson et al., 1996, Dille et al., 1997, Pilot-Matias et al., 1996a, Tacke et al., 1997a). In most developed countries,

HPgV RNA detection rates and HPgV E2-Ab-positive rates in volunteer blood donors amount to 1-4% and 5-13%, respectively, with higher rates in developing countries

(Blair et al., 1998, Gutierrez et al., 1997, Pilot-Matias et al., 1996a, Tacke et al., 1997b,

Mohr and Stapleton, 2009, Williams et al., 2004). HPgV RNA and Ab are more prevalent in high risk groups, such as intravenous drug users or people suffering from other blood-borne or sexually transmitted infections, reaching exposure rates over 80%

(Williams et al., 2004, Scallan et al., 1998, Tacke et al., 1997b, Stapleton et al., 2011).

Clearance of HPgV from blood occurs within two years in most immunocompetent individuals and coincides with the development of protein conformation-dependent, long-lasting anti-E2 Abs (Berg et al., 1999, Tanaka et al., 1998, McLinden et al., 2006,

Pilot-Matias et al., 1996b, Tacke et al., 1997b). Due to the fact that the Ab response appears to be restricted to E2, this antigen is assumed to possess the immunodominant epitopes (McLinden et al., 2006).

Equine pegivirus (EPgV) RNA has been detected in 0.8% to 14.2% of horses from the

United States, China and Brazil (Kapoor et al., 2013a, Lyons et al., 2014, Lu et al., 2016,

Agnello et al., 1999, Figueiredo et al., 2019). An ELISA for the detection of Abs against this virus was established using the non-structural protein 3 helicase domain (NS3h) and bacterial expression in E. coli (Lyons et al., 2014). In the study, 218 of 328 (66.5%) horses were positive for NS3h Abs in ELISA, of which 88% (192 of 218) were confirmed by Western blot (WB). Contrary to findings in HPgV with E2 Ab assays, in the NS3 Ab assay for EPgV, 10 of the 12 RT-PCR positive horses were also Ab positive (Lyons et al., 2014).

Transmission of human pegivirus can occur both parenterally through blood and sexual contact, as well as vertically (Dawson et al., 1996, Linnen et al., 1996, Schmidt et al., 1996, Feucht et al., 1996). The transmission of EPgV has not been studied in detail.

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However, transmission of Theiler’s disease associated virus (TDAV, Pegivirus E) between horses has been shown to be possible by experimental inoculation

(Chandriani et al., 2013). Similarly, pegiviruses can be transmitted to different species of New World monkeys by experimental infection via the blood-borne route (Stapleton et al., 2011). Transmission routes of pegiviruses infecting bats were not extensively studied; however, viral RNA was detected in saliva of viremic bats, suggesting that horizontal or even zoonotic transmission may be possible (Epstein et al., 2010).

To our knowledge, an assay for the detection of PPgV-specific Abs has not been described and studies on virus transmission between pigs have not been reported.

Therefore, we expressed PPgV proteins NS3h and C-terminally truncated E2 (E2t) intracellularly in E. coli and implemented WB and indirect ELISA methods for the detection of PPgV-specific serum Abs. A serological assay will permit the investigation of PPgV infection characteristics, including time point of seroconversion, relevance and relation of IgM, IgG, and IgA Abs, as well as maternally derived Abs. During the study, PPgV field infected domestic pigs were monitored alongside their piglets and serum and excretion samples were obtained every 2-3 weeks for evaluation of PPgV

RNA presence and potential serological responses. Taken together, this study provides the first insights into possible PPgV transmission routes and PPgV-specific Ab detection.

Materials and methods

Samples

Serum samples available for Ab detection originated from apparently healthy domestic pigs from Europe (Germany, Italy, Poland, Great Britain, Switzerland and

Serbia) and Taiwan, which were used in a previous study on PPgV RNA detection rate

(Kennedy et al., 2019). They were collected between 2014 and 2018 within the

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framework of national veterinary health management in concordance with national legal and ethical regulations.

Additionally, serum as well as nasal swabs, oral swabs, and fecal and urine samples were collected from six healthy, originally PPgV positive pigs (animals A-F) from a farm in Lower Saxony, Germany, and from initially PPgV negative control pigs

(animals G, H, J, and K) from the Farm for Education and Research in Ruthe of the

University of Veterinary Medicine Hannover. All animals were housed in the Clinic for Swine and Small Ruminants and Forensic Medicine and Ambulatory Service of the

University of Veterinary Medicine Hannover after the initial PPgV RNA screening and were sampled every 2-3 weeks for a maximum of 46 weeks (see Figure 3-4). Excretion samples were taken from all animals at each sampling time point from week 7 to week

46.

Animals A and B, both sows, were inseminated in week 20 while both animals were

PPgV RNA positive. All 13 piglets from sow A (P1-P13) and 13 piglets from sow B

(P14-P26) were sampled after birth (before colostrum intake), and every 2 weeks thereafter for a maximum of 12 weeks. Samples taken from piglets included serum, nasal, oral and fecal swabs. For the detection of PPgV RNA in piglet excretion samples, swabs of three piglets of each sow were selected for each sampling week. Two vaginal swab samples were taken from each sow during the birth of their piglets. Sampling was approved by Lower Saxony’s official authorities (LAVES AZ 15A602) and was carried out in accordance with German legislation (TierSchVersV).

All swab samples were soaked in 1 ml cell culture media containing antibiotics for a minimum of 2 hours (h), after which media was used for RNA isolation, or stored.

Feces were diluted 1:10 in phosphate buffered saline (PBS), vortexed vigorously and centrifuged for 20 min at 4000 × g, and supernatant was stored. All samples were stored at -80 °C until use.

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RNA isolation and PCR

Viral RNA was isolated from 200 µl of serum (100 µl of serum from piglets in week 36 due to limited volume available), urine, fecal swab supernatant, or media using the

IndiMag Pathogen Kit (Indical Bioscience, Germany) and the KingFisher Duo Prime extractor (Thermo Scientific, Germany) and was eluted in 100 µl elution buffer.

Quantitative reverse-transcription PCR (qRT-PCR) was used to detect PPgV RNA as previously described (Kennedy et al., 2019). Control RNA was added to excretion samples and detected by qRT-PCR (primers EGFP-1-F

(5’-GACCACTACCAGCAGAACAC-3’), and EGFP-2-R

(5’-GAACTCCAGCAGGACCATG-3’) and probe EGFP-HEX

(5’-[HEX]-AGCACCCAGTCCGCCCTGAGCA-[BHQ1]-3’)) to confirm successful

RNA isolation, as previously described (data not shown) (Hoffmann et al., 2006).

Complementary DNA of the viral genomic RNA of PPgV isolate PPgV_903/Ger/2013

(GenBank accession number KU351669.1) was synthesized using SuperScript II

Reverse Transcriptase (Invitrogen, Germany). PCR for the amplification of the genomic regions encoding the predicted NS3 helicase domain and the truncated E2 was performed using primers PPgV/NS3H/fwd/19

(5’-ATTACTCGAGGTGGTCCCCTGGGCCAACATGCCTCAGGA-3’) and

PPgV/NS3H/rev/20

(5’-CCGCAGATCTGTCATACCACAATCAGTCACAGTGTCA-3’), or

PPgV/E2T/fwd/17 (5’-ATTACTCGAGCTTCTGCTCCTTGCTGCTGCTG-3’) and

PPgV/E2T/rev/18 (5’-CCGCAGATCTGTAGGAAACTGGTCTGTGTACTCAT-3’), respectively, containing appropriate restriction sites for subcloning (XhoI and BglII, underlined).

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Expression of PPgV NS3 helicase and truncated E2 recombinant proteins

The NS3h (811 bp) and E2t (994 bp) amplicons were cloned into a pET-19B vector

(Novagen, Germany) downstream of a polyhistidin tag. Plasmid and inserts were digested using appropriate restriction enzymes (Thermo Scientific) and were purified using the GeneJet Gel Extraction kit (Thermo Scientific) followed by ligation with T4 ligase (New England Biolabs, Germany). Recombinant E. coli Top10 clones were grown in LB-medium and were selected with ampicillin (50 µg/ml). Plasmid integrity was confirmed by PCR and sequencing.

For protein expression, One Shot BL21(DE3)pLysS chemically competent E. coli

(Invitrogen) were used according to the manufacturer’s instructions. Fresh cultures were added to 200 ml LB-medium containing 50 µg/ml ampicillin and cultured at 37

°C and 250 rpm. At an optical density at 600 nm of 0.6, bacterial expression was induced with 1 mM Isopropyl-beta-D-thiogalactopyranosid (IPTG) for 2 h. Bacteria were centrifuged at 4000 × g for 20 min at 4 °C. Cell pellets were weighed and frozen at 20 °C. B-PER complete protein extraction reagent (Thermo Scientific) was used for lysis of bacteria and soluble and insoluble cytoplasmic fractions were analyzed for the presence of NS3h and E2t proteins in Coomassie blue stained sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gels and Western blot. For E2t protein, the insoluble cytoplasmic fraction was washed four times with B-PER diluted

1:10 and the remaining pellet was resuspended in 1% SDS for use in Western blot.

Purification of PPgV NS3 helicase recombinant protein

Soluble proteins containing NS3h were diluted 1:10 in fast protein liquid chromatography (FPLC) buffer (20 mM sodiumhydrogenphosphate, 500 mM sodiumchloride, pH 7.4) and imidazole was added to a final concentration of 40 mM.

The diluted sample was filtered through a 0.22 µm filter membrane. Protein was purified using the Äkta Pure FPLC and a HisTrap excel 1 ml column (GE Healthcare,

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Sweden). After sample application (flow rate 1 ml / min), the column was washed with

20 ml FPLC buffer containing 40 mM imidazole and protein was eluted in 10 fractions of 0.5 ml each of FPLC buffer containing 500 mM imidazole. Elution fractions were analyzed by SDS-PAGE and Coomassie blue staining, as well as WB. Protein concentrations were determined using the Pierce BCA Protein Assay Kit (Thermo

Scientific). Fractions containing high concentrations of target protein were pooled and stored at 4 °C containing 0.1% sodium acid as preservative.

SDS-PAGE, Coomassie blue staining and Western blot

To visualize bacterial protein expression and to confirm FPLC-purification of NS3h protein, diluted bacterial lysate containing crude NS3h or E2t protein, as well as eluted fractions of NS3h, were analyzed on 12.5% SDS-PAGE gels stained with Coomassie blue or blotted onto PVDF membranes in a semidry electroblotting procedure.

Membrane blocking, Ab incubation and washing steps were carried out with TBS-0.1%

Tween20-buffer. PVDF membranes were blocked overnight with 2% Amersham ECL

Prime Blocking Reagent (GE Healthcare). Membranes were then incubated with Anti-

His6 mouse monoclonal Ab (Roche, Germany) diluted 1:100 for 1 h at RT and then washed four times for 15 min. Polyclonal Rabbit Anti-Mouse Immunoglobulins / HRP

(Dako, Denmark) diluted 1:2,000 was then added and incubated again for 1 h at RT, followed by four more steps of washing. Amersham ECL Prime Western Blotting

Detection Reagent (GE Healthcare) was used for the visualization of bound Ab.

Serum samples were tested on WB membranes obtained after SDS-PAGE and transfer of purified NS3h or crude E2t was performed as described above. Membranes were blocked as described above and incubated with serum diluted 1:500 in blocking reagent for 1 h on a shaker at RT. Membranes were washed and anti-pig IgG (whole molecule)-Peroxidase antibody produced in rabbit (Sigma Aldrich, Germany) diluted

1:30,000 was added for 1 h at RT, followed by another round of washes. Bound Ab was

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detected using ECL Prime or ECL Select Western Blotting Detection Reagents (GE

Healthcare).

ELISA

Nunc Medisorp ELISA plates (Thermo Scientific) were coated with 250 ng / well

(ELISA250) or 100 ng / well (ELISA100) of NS3h in 100 µl coating buffer (0.1 M NaCO3, pH 9.6) per well over night at 4 °C. Plates were washed three times with 360 µl PBS-

0.05% Tween20 (PBS-Tw) and blocked with 4% skimmed milk powder in PBS-Tw for

2 h at room temperature (RT) and subjected to another round of washes. Porcine serum samples were diluted 1:25 in 4% skimmed milk powder in PBS-Tw and 100 µl per well were incubated for 1 h at 37 °C. Plates were washed again and anti-pig IgG (whole molecule)-Peroxidase produced in rabbit (anti-pig IgG, Sigma Aldrich) diluted

1:35,000 in 4% skimmed milk powder in PBSM-Tw was added and incubated for 1 h at 37 °C. After washing, tetramethylbenzidine (Sigma Aldrich) was added and plates were incubated in the dark for 10 min at RT. The reaction was stopped with 1 M hydrochloric acid and optical densities (ODs) were determined automatically (TECAN

Sunrise Remote, Tecan, Switzerland) at a wavelength of 450 nm and a reference wavelength of 620 nm.

Results

NS3h protein purification

Purification of NS3h was achieved by immobilized metal ion chromatography (IMAC) in FPLC, which permitted elution of 0.67 mg protein / ml. Western blots and

Coomassie blue stained gels were used to assess the elution and the purity of the target protein. Though weak bands of unspecific protein were visible at larger protein sizes, purity of NS3h was high (Figure 3-1).

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Figure 3-1. Coomassie gel of NS3h protein before and after purification by IMAC. Lane 1 shows crude soluble protein that was applied onto the HisTrap column before dilution with FPLC buffer. Lane 2 shows flow through of FPLC, indicating proteins that did not bind to the column. Lanes 3, 4 and 5 show different fractions of eluted proteins with NS3h at a size around 37 kDa (compare Figure 3-2, lane 1: detection of NS3h by the anti-His6 Ab. Additional unspecific bands can be seen in lanes 3 and 4 at a protein size of ~70 kDa.

NS3h-specific serum antibody reactivity in Western blot and ELISA

PPgV RNA positive and negative serum samples were prescreened using ELISA250 to identify samples suitable for validation of WB and ELISA (data not shown). Further evaluation of 18 samples with varying reactivity in the preliminary ELISA250 was performed by WB and ELISA100. Clearly visible NS3h-specific bands at a protein size of ~37 kDa were seen following incubation with 5 of 18 samples (Figure 3-2). One sample evidenced unspecific bands at a protein size ~70 kDa, and variations in unspecific background reactions were visible, making evaluation of possible bands difficult in 6 samples. Differences between ODs of ELISA250 and ELISA100 ranged from a factor change of 0.6 (sample in lane 12, Figure 3-2) to 0.92 (sample in lane 3, Figure

3-2; Table 3-1).

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Figure 3-2. Western blots of purified NS3h protein incubated with serum samples as first antibody (Ab). Lane 1 shows the position of the NS3h-His fusion protein at a molecular weight of ~37 kDa using the anti-His6 Ab. Lanes 8, 11, 14, 17, and 18 evidence an NS3h-specific band. Background varies from low (lanes 7, 9, 13, 14, 15, and 19) to high (lanes 4, 5, and 18). Unspecific bands at a protein size ~70 kDa can be seen in lane 6.

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Lane no. WB WB ELISA250 ELISA100 WB PPgV in NS3h NS3h unspecific OD OD background RNA WB band bands 17 1.91 1.5 + + - 6 1.61 1.18 (+) + + - 14 1.58 1.19 + - + 18 1.57 1.28 + ++ + 11 1.45 1.08 + + - 4 1.08 0.84 (+) ++ - 2 1.04 0.8 (+) + + 8 0.93 0.61 + + + 3 0.82 0.75 (+) + + 5 0.82 0.64 (+) ++ - 19 0.76 0.52 - - 10 0.67 0.45 (+) + - 12 0.64 0.44 + - 15 0.38 0.23 - - 9 0.13 0.09 - + 16 0.11 0.1 + - 7 0.09 0.07 - + 13 0.07 0.04 - -

Table 3-1. Characterization of selected serum samples. ELISA optical densities (OD) of serum samples and their reactivity with NS3h and background Western blot, as well as PPgV RNA detection. Samples are arranged in descending order of their OD in ELISA250 (arrow). Detection of NS3h in WB is indicated as positive [+], uncertain [(+)], or negative [-].

E2t-specific serum antibody reactivity in Western blot

The expression of C-terminally truncated E2 was successfully achieved in the E. coli strain BL21(DE3)pLysS used here, and the protein was detectable in the insoluble cytoplasmic fraction of the bacterial lysate using the anti-His6 Ab in WB at a protein size of ~40 kDa (Figure 3-3, lane 1). Due to detection of the target protein E2t in the insoluble fraction indicative of expression in bacterial inclusion bodies, crude protein containing E2t was prepared for WB analyses by repeated washing of the insoluble protein pellet and final resuspension in 1% SDS.

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Two samples (one of which is seen in lane 17 of Figure 3-2; the other is not shown here) that evidenced NS3h-specific Ab reactivity in WB and ELISA were implemented in

WB with E2t crude protein (equivalent to isolation from 100 µl of bacteria), one of which evidenced an E2t-specific band (Figure 3-3, compare with lane 17 in Figure 3-2).

Figure 3-3. Western blot of crude E2t protein incubated with serum samples that showed NS3h-specific antibody (Ab) reactivity in Western blot and ELISA as first Ab. Lane 1 shows the position of the E2t-His fusion protein at a molecular weight of ~40 kDa using the anti-His6 Ab. An E2t-specific band is visible in lane 2, while unspecific bands at a protein size ~35 kDa are visible in lanes 2 and 3.

PPgV RNA in serum and excretion samples of pigs and piglets

Six fattening pigs from a farm in Lower Saxony, Germany, were found to be PPgV

RNA positive in serum and were moved to the Clinic for Swine and Small Ruminants and Forensic Medicine and Ambulatory Service of the University of Veterinary

Medicine Hannover. Animals A and B were continually tested PPgV genome positive

(samples taken every 2-3 weeks) by qRT-PCR for 38 and 27 weeks, respectively (Figure

3-4). Animals C, D, E, and F were PPgV RNA negative at the second sampling in week

3 and only animal D was again found PPgV RNA positive from weeks 13 to 33.

Additionally, excretion samples, including nasal and oral swabs, urine and feces, were taken at each time point (from week 7 to week 46) to assess possible routes by which

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PPgV might be spread, all of which were found to be negative for PPgV RNA. Initially

PPgV genome negative control animal G was found to be PPgV RNA positive in serum in week 17. Other negative control animals H, I and J were tested negative in serum and excretion samples throughout the study.

Animals A and B were inseminated in week 20. While animal A remained PPgV RNA positive for the duration of the pregnancy and until 12 days post-partum, animal B was last found PPgV genome positive in week 27, 50 days after insemination. In week

36, both sows bore 13 piglets each. PPgV RNA was detected in serum of one piglet (P8) directly after birth (before colostrum intake) in week 36 and contained 1,412 RNA copies / ml serum (Cq value 37.61). All other piglet serum samples were tested PPgV

RNA negative by qRT-PCR. For each sampling week, excretion samples were selected from three piglets of each sow for detection of PPgV genome, and samples from serum

RNA positive piglet P8 in week 36 were additionally included. However, all excretion samples from piglets were found to be PPgV RNA negative.

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Figure 3-4. PPgV viral genome quantity in serum (RNA positive results only) during the course of infection in domestic pigs. Sampling periods of each animal, gestation of animals A and B, and piglet sampling are indicated below. Animals A-F were PPgV genome positive at the first sampling time point (week 0). Animals A and B were not sampled during the first four weeks of gestation. Negative control animals G-J were genome negative at their respective first sampling time point. Animals E, F, H, I, and J remained PPgV RNA negative for the duration of sampling, and animal G and piglet P8 were PPgV genome positive only in sampling week 17 and week 36 (piglet sampling directly after birth, before colostrum intake), respectively.

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Discussion

Since the discovery of PPgV in recent years, studies have shown that the virus is widely distributed and RNA detection rates ranged from 0% in Serbia, Taiwan and

Switzerland in one study to 15.1% in another study from the United States (Kennedy et al., 2019, Yang et al., 2018). Though PPgV infections can persist, genome detection alone may lead to underestimation of virus distribution, as animals with transient and past infections can be overlooked (Baechlein et al., 2016, Dawson et al., 1996).

However, an assay for the detection of Abs induced during PPgV infection, which would aid not only in the estimation of PPgV exposure, but also give deeper insights into the immune response, has not been described so far. Therefore, in this study, PPgV proteins were expressed intracellularly in E. coli to allow for the detection of PPgV- specific Abs using Western blot and indirect ELISA.

The helicase domain of PPgV NS3 was expressed in the soluble cytoplasmic fraction of E. coli, which facilitated the purification with IMAC under native conditions. Purity of the protein was optimal at 40 mM Imidazole during washing and binding and yielded elution with high protein concentrations. NS3h has a predicted protein size of

33.5 kDa, which coincides with results obtained here showing NS3h in Western blot at a size of ~37 kDa.

PPgV NS3h-specific Abs were detectable in porcine sera using Western blot, showing that PPgV NS3 can induce an Ab response in the porcine host. Results obtained here indicate that viremia and NS3h Ab response can be detected simultaneously, similar to observations in EPgV (Lyons et al., 2014). WB with purified NS3h protein evidenced unspecific bands, as they can also be seen in Coomassie blue stained SDS-PAGE gel

(Figure 3-1), in one of 18 samples shown here (lane 6 in Figure 3-2). All other serum samples reacted only with NS3h, suggesting that protein purity is sufficient for the methods implemented here.

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Comparison of WB and ELISA results indicates that samples showing a clear band in

WB also have high ODs in ELISA. Nonetheless, ODs over 0.93 in ELISA250 were found in serum samples that did not detect a clear NS3h-specific band in WB. So far, it is unclear whether such high ELISA reactivities may be caused by ELISA background

(compare lane 2 in Figure 3-2) or whether WB background conceals NS3h-specific bands. To yield better differentiation between Ab positive and negative samples, ongoing experiments target further improvement of ELISA and WB conditions to reduce background caused by serum components binding to ELISA plates and WB membranes unspecifically, such as changing washing conditions (i.e. higher concentrations of Tween20). Furthermore, E2t appears to be a promising immunogenic

PPgV protein for implementation in indirect Ab ELISA. The necessity of measuring anti-E2 Abs, specifically those that are protein conformation-dependent, is evident in

HPgV serology (Berg et al., 1999, Tanaka et al., 1998, McLinden et al., 2006, Pilot-Matias et al., 1996b, Tacke et al., 1997b). Experiments pertaining to the expression of soluble

E2t or purification of E2t under denaturing conditions followed by refolding are ongoing.

This study also investigated possible transmission routes of PPgV. Analysis of PPgV genome in nasal discharge, saliva, urine and feces showed that these are not probable routes of virus shedding from persistently or transiently infected pigs. Likewise, vaginal swab samples of animals A and B taken during the birth of their piglets did not contain PPgV RNA, though animal A was viremic at the time, indicating that transmission during birth may be unlikely. However, detection of PPgV RNA in the serum of piglet P8 directly after birth suggests intrauterine infection. Sows were housed separately for birth of piglets and P8 was born of sow B, which cleared viremia before day 69 of gestation, making cross contamination of the piglet sample highly unlikely. Intrauterine infection of P8 likely took place before day 69 of the gestation, when PPgV RNA was detectable in serum of the sow. Clearance of the virus from serum of P8 by week 38 (at 16 days of age) could have been facilitated by maternally 52

Chapter 3

derived Abs. None of the other piglets were found PPgV RNA positive for up to 10 weeks of age, suggesting that transmission from sow A, which was viremic at the time, to piglets did not occur. The fact that this phenomenon was only found in one of 26 piglets, and that the Cq value was relatively high (37.61), indicates that intrauterine infection is possible, but that the vertical transmission route is not efficient in pegivirus infection of pigs. This coincides with higher PPgV genome detection rates found in older animals than in piglets, as was seen in previous studies (Lei et al., 2019, Kennedy et al., 2019). Blood-borne transmission between pigs could for instance take place iatrogenically during vaccinations, drug injections or other treatments, through direct blood contact of wounded pigs, or due to cannibalism.

Results obtained here show that PPgV is most probabaly not transmitted by virus shedding in excretion, but that horizontal transmission by the blood-borne route and occasional intrauterine transmission, as can be found in HPgV, are more likely

(Dawson et al., 1996, Linnen et al., 1996, Schmidt et al., 1996, Feucht et al., 1996). This study also provides first insights into the PPgV-specific immune response by detection of NS3 and E2-specific Abs in PPgV genome positive and negative porcine serum samples. The establishment of reliable indirect ELISAs will allow for high throughput

Ab detection in serum samples and will increase the understanding of the immune response induced during pegivirus infection. Furthermore, investigation of the Ab presence in other species with improved test systems will give deeper insights into the host range and elucidate possible reservoir hosts like wild boar, where PPgV genome has not been detected so far (Kennedy et al., 2019).

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Acknowledgements

We thank Vesna Milićević (Belgrade, Serbia), Ming-Chung Deng and Chia-Yi Chang

(New Taipei City, Taiwan), Serum Bank of the Federal Food Safety and Veterinary

Office (Bern, Switzerland), Animal and Plant Health Agency (Weybridge, UK),

Francesco Feliziani and Gian Mario De Mia (Perugia, Italy), Katarzyna Podgórska and

Katarzyna Stępniewska (Puławy, Poland), Hua-Ji Qiu and Yuan Sun (Harbin, China),

Sarah Derking and Jörg Tenhündfeld (Vreden, Germany) and Thomas Große Beilage

(Essen/Oldenburg, Germany) for providing porcine serum samples. Special thanks to

Alexander Postel and Inga Grotha for sample library preparation.

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4 Overall Discussion

4.1 PPgV RNA detection in domestic pig serum samples from Europe and

Asia

New high-throughput sequencing technologies are facilitating the rapid discovery of ever more viral genome sequences, expanding our knowledge of the world’s virome and presenting us with new viruses for exploration at an increasing rate.

Characterization of such newly found viruses is crucial for understanding their impact on the host. At the beginning of this project, PPgV had only recently been discovered and the information that was available was limited to three full-length genomic sequences, RNA detection rate of 2.2% found in 255 German domestic pigs, and observation of persistent infection in three pigs (Baechlein et al., 2016). Consequently, this project aimed at broadening our knowledge of pegivirus infection in the porcine host through the investigation of virus distribution, tissue tropism, transmission routes, and antibody response.

Firstly, the establishment of a reliable qRT-PCR, combined with the creation of an in vitro-transcribed RNA copy standard, was necessary for the detection and accurate quantification of PPgV genome. Dual-labeled-probe-based qRT-PCR using TaqMan polymerase (TM qRT-PCR) is a highly specific PCR method, that evidenced efficiency of over 96% as measured using the in vitro-transcribed RNA copy standard in this study (Kennedy et al., 2019). Additionally, no lack in sensitivity was observed when comparing the newly developed TM qRT-PCR and the previously described SYBR-

Green-based qRT-PCR (Baechlein et al., 2016), indicating that sensitivity is not impaired due to higher specificity caused by the addition of a probe. Moreover, partial

NS3 sequences acquired in the study after detection of PPgV by TM qRT-PCR (see

Chapter 4.3) evidenced up to three mismatches in the probe sequence (one of which

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was at the third position from the 5’-end), suggesting sufficient sensitivity of TM qRT-

PCR even with a certain degree of genetic diversity in the target viral RNA sequence.

Investigation of serum samples originating from apparently healthy domestic pigs from Europe and Asia revealed that PPgV viremia was present in 2.7% of animals, though the range covered 0% (Switzerland, Serbia, Taiwan) to 10.3% (Great Britain)

(Kennedy et al., 2019). As mentioned in Chapter 2, these detection rates coincide with findings of PPgV RNA in pigs from Germany (2.2%) and China (7.25%), and are somewhat lower than the rate found in USA (15.1%) (Baechlein et al., 2016, Lei et al.,

2019, Yang et al., 2018, Kennedy et al., 2019). Due to the apparently wide distribution of PPgV, it seems unlikely that the virus is not present in the pig populations of

Switzerland, Serbia and Taiwan. The absence of detectable PPgV may rather be attributable to limited sample size combined with low prevalence, and limitations of the methods used here (Kennedy et al., 2019). Follow-up investigation of additional samples, as well as Ab detection, will clarify whether PPgV may be present in pigs in these countries. To date, the reasons for variance in the detection rates of PPgV RNA between the different countries are unknown. The findings presented in this project highlight that PPgV distribution within different age groups is uneven, increasing from piglets to sows and boar, as was also reported for pigs in China (Kennedy et al.,

2019, Lei et al., 2019). This shows that factors like herd management could influence

PPgV spread within and between herds. Future research targeting virus entry into pig populations and infection dynamics within herds is necessary to understand the spread of PPgV.

4.2 Phylogenetic analyses of PPgV

At the beginning of this project, three full-length genome sequences were available from PPgV-infected domestic pigs from Germany, which occupied a separate branch in the phylogenetic tree of pegiviruses (Baechlein et al., 2016). In addition, three further

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near full-length PPgV sequences from pigs from the United States, which shared high nucleotide identitities among them (96.2-97.0%) and somewhat lower identities with sequences from Germany (83.7-89.0%), were reported (Yang et al., 2018). These six sequences were included in phylogenetic analyses using a partial NS3-coding genome region in this project, which additionally analyzed PPgV originating from Germany,

Italy, Poland, Great Britain and China. Clusters of sequences from geographically close regions, as well as identical sequences obtained from pigs of the same herd indicated local spread of PPgV (Kennedy et al., 2019). Additionally, global virus spread was indicated by high similarity of sequences obtained from geographically distant regions of the world (i.e. from China and Great Britain), and might be facilitated by trade of pigs or pig products. As is seen in pegivirus infection of horses (Pegivirus D and E) and humans (Pegivirus C and H), distinct pegivirus species can be found within the same host species (Smith et al., 2016). Similarly, further pegivirus species that can infect pigs but are distinct from Pegivirus K may so far be undiscovered.

In comparison to the relatedness of Pegivirus A isolates originating from bats and primates, sequences clustering with PPgV (Pegivirus K) have solely been isolated from pigs, which indicates specific host tropism (Baechlein et al., 2016, Kennedy et al., 2019,

Lei et al., 2019, Yang et al., 2018, Smith et al., 2016).

4.3 No detection of PPgV RNA in wild boar

In our study, 800 wild boar serum samples from Lower Saxony, Germany, tested negative for PPgV RNA. As mentioned in Chapter 2, future investigation of wild boar serum in Ab assays will help in determining whether wild boar are indeed negligible as possible (reservoir) hosts for porcine pegiviruses. However, shared infection of wild boar and domestic pigs can be seen in other members of the Flaviviridae, like CSFV and

Atypical porcine pestivirus, and also in other viruses like African swine fever virus or

Pseudorabies virus (Cagatay et al., 2018, Postel et al., 2018, Moennig, 2015, Guinat et

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al., 2016, Ruiz-Fons et al., 2008). Transmission routes of viruses between domestic pigs and wild boar include direct contact of infected animals, but also indirect transmission via inanimate vectors, for instance by the consumption of contaminated feed (Ruiz-

Fons et al., 2008, Guinat et al., 2016, Fritzemeier et al., 2000, Moennig, 2015). As mentioned in Chapter 3 and further discussed in Chapter 4.6, PPgV transmission may require contact with PPgV-infected blood or products containing such blood or blood components, possibly reducing the probability of viral transmission between wild boar and domestic pigs in the case of PPgV.

In any case, HPgVs have been suggested to be ancient viruses, as they are well-adapted to their host, are widely distributed (even among highly isolated populations in humans), with evidence of co-migration, and may well be a symbiont or commensal of humans (Chivero and Stapleton, 2015, Simmonds and Smith, 1999, Pavesi, 2001,

Sharp and Simmonds, 2011). Furthermore, bats may be ancient hosts of pegiviruses, according to the phylogenetic diversity of pegiviruses found in the different species from diverse parts of the world (Quan et al., 2013). These are indicators that pegiviruses could be present in the wild boar population. Due to hygiene control measures implemented in pig production and to reasons mentioned above, transmission of pegiviruses between wild boar and domestic pigs may be limited, and pegiviruses in wild boar may by genetically distant. Consequently, wild boar pegiviruses may be undetectable by RNA detection methods used in this study

(Kennedy et al., 2019). Once again, Ab assays may be a useful tool in the further investigation of pegivirus infection in different wild boar populations.

4.4 Persistent and transient PPgV infections

Persistence of pegivirus infection in pigs could be shown here for five, six, and nine months (animals A, B, and D in Chapter 3, see Figure 3-4), and was observed for 22 months in a previous study (Baechlein et al., 2016). These observations coincide with

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findings in HPgV infection, in which immunocompetent individuals typically clear viremia within two years (Gutierrez et al., 1997, Tanaka et al., 1998).

One longitudinal study (sampling time points several months to years apart) described

HPgV viral load (VL) dynamics with increased VL directly from onset, steady VL level during persistence, and no reduction in VL in the years preceding clearance of viremia, though one individual showed a VL reduction in the months prior to clearance (Lefrere et al., 1999). The observations made in this project with PPgV viremia, as seen in

Chapter 3, largely coincide with this description. However, VL increase and decrease during onset and clearance, respectively, were evident within four weeks in this study

(Chapter 3, Figure 3-4, PCR negative to steady VL in animal D, and steady VL to PCR negative in animals A, B, and D). These phenomena in PPgV infection might have remained undetected, had samples been acquired less frequently. Therefore, sampling every two to three weeks, as was performed in this project, was adequate to obtain detailed insights into VL dynamics.

In the context of first investigations, the highest PPgV genome load in serum was detected in a sample from Germany (2 × 106 copies / ml), and the average genome load overall was lower for PPgV (3.8 × 105 copies / ml) than for HPgV (average plasma concentration > 1 × 107 copies / ml) (Kennedy et al., 2019, George et al., 2003, Chivero et al., 2014). However, higher genome loads were detected later (maximum 1.1 × 107 copies / ml; Chapter 3, Figure 3-4), thus reaching serum concentrations closer to those observed in individuals infected with HPgV and similar to findings reported for EPgV infection (George et al., 2003, Chivero et al., 2014, Kapoor et al., 2013a). In this project, transient infection was evident in animal G, and may also have been the case in animals C, E and F (Chapter 3, Figure 3-4). Intriguingly, the mean VL was higher in persistently infected animals (2.4 × 106 copies / ml) than in those with apparently transient infection (4.0 × 103 copies / ml). This shows that PPgV seems to establish an efficient persistent infection, with constantly high viral replication at an, as of now,

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unknown replication site (possibly PBMCs, liver or thymus; see Chapter 4.5). Animals

A-F originated from the same herd and were of the same age group. Moreover, they appear to have been infected with the same PPgV strain, according to preliminary

PPgV genome sequencing results, in which sequences acquired from animals A-F in week 0 were identical (sequencing was performed as described in Chapter 2 (Kennedy et al., 2019)). This indicates no clear tendency towards host or viral factors contributing to this distinction. Persistent infections were also observed in EPgV infection in two horses over 3.5 years, while four other horses apparently cleared viremia (Kapoor et al., 2013a). Factors contributing to pegivirus persistence in immunocompetent hosts are largely unknown. Concordantly, why certain animals clear PPgV infection immediately, while others become persistently infected, at least for some time, remains to be elucidated.

Here, one piglet was found PPgV RNA positive directly after birth, before colostrum intake, indicating an intrauterine infection that took place before day 69 of the gestation, while the sow was viremic. This strongly suggests that the piglet was persistently infected for the remainder of the gestation after initial infection, and that it cleared the infection within two weeks of birth, perhaps due to uptake of maternally derived neutralizing Abs (Chapter 3).

Furthermore, short transient infections with detectable viremia for only days, as was seen in animal G in Chapter 3, may be common in the porcine host, thus leading to lower RNA detection rates. In HPgV, RNA and E2-Ab detection rates are added to calculate the total exposure, because E2-specific Abs coincide with viral clearance

(Tacke et al., 1997, Gutierrez et al., 1997, Thomas et al., 1998). Should this also be the case in PPgV infection, RNA detection alone would clearly underestimate PPgV exposure. Future research should include Ab detection for examination of seroprevalence among pig populations to resolve this possible issue.

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4.5 Investigation of PPgV tissue tropism

Determination of the tissue tropism is an essential step for better understanding the pathogenesis and viral life cycle of PPgV. Detection of PPgV RNA in PBMCs (Chapter

2, Figure 2-2) by FISH and qRT-PCR shows parallels to HPgV cell tropism (Fogeda et al., 1999, George et al., 2003, Chivero and Stapleton, 2015, Kennedy et al., 2019).

Moreover, the detection of PPgV in the thymus of infected animals coincides with lymphotropism. It would be intriguing to investigate whether the thymus may be a primary site of PPgV replication, and if so, whether age-related thymic involution leads to a reduction in viral replication or to a shift in the replication site.

In this project, PPgV RNA detection in PBMCs, liver and thymus may be explained by infection of hematopoietic precursor cells. It has been speculated that the primary permissive cell type(s) for HPgV may be hematopoietic stem cells that carry the virus throughout differentiation, leading to reduced activation and proliferation of lymphocytes (Chivero and Stapleton, 2015, Chivero et al., 2014).

As has been found in HPgV, serum concentrations of PPgV RNA are higher than those detected within organs, including the liver (Pessoa et al., 1998, Kennedy et al., 2019).

HPgV was initially thought to be hepatotropic, and indeed viral RNA was present in the liver of infected patients, but further research indicated that the virus did not in fact replicate in the liver (Chivero and Stapleton, 2015, Fan et al., 1999, Pessoa et al.,

1998, Tucker et al., 2000, Berg et al., 1999, Laskus et al., 1997, Laras et al., 1999). PPgV

RNA was detectable in the liver of three pigs, which could indicate hepatotropism, but could also be attributed to accumulation of viral RNA in the liver and virus replication within lymphocytes present in liver tissue, as mentioned above. HPgV has been detected in the brain tissue of patients with encephalitis and it was suggested that the virus may cross the blood-brain barrier passively in lymphocytes (Balcom et al., 2018,

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Tuddenham et al., 2019). Similar modes of passive transportation of PPgV to various tissues may take place within infected PBMCs (Kennedy et al., 2019).

Findings described in Chapter 2 indicate that PBMCs are promising candidates for the further exploration of PPgV replication sites (Kennedy et al., 2019). Analysis of PPgV replication in PBMCs from infected animals ex vivo or infection of PBMCs from PPgV

RNA negative animals may result in enhanced understanding of the life cycle of pegiviruses in pigs. Likewise, detection of negative-strand RNA in samples acquired during the course of this project may facilitate the clarification of replication sites for

PPgV.

4.6 Insights into PPgV transmission routes

At present, little is known about the transmission of animal pegiviruses. HPgV has been shown to be transmitted by exposure to infected blood, sexually, and vertically from mother to child (Bhanich Supapol et al., 2009, Hino et al., 1998, Kleinman, 2001,

Lin et al., 1998, Ohto et al., 2000, Stapleton, 2003). Similarly, transmission of EPgV was successful by inoculation of naïve animals with virus-containing serum, and the virus was present in commercially available horse sera (Chandriani et al., 2013, Postel et al.,

2016).

In HPgV infection, RNA and Ab detection rates increase together with the probability of infection from healthy blood donors in developed countries, to blood donors in developing countries, and finally to high-risk groups (i.e. intravenous drug users), where exposure rates (RNA plus Ab positive rate) can be nearly universal. PPgV was detected in 0% - 10.3% in this study, and up to 15.1% in another study from the United

States, which may indicate that RNA and Ab detection rates are similar to those of

HPgV in healthy blood donors rather than individuals of high-risk groups.

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Results obtained in the presented study reveal that PPgV is probably not transmitted by shedding of the virus by excretion (Chapter 3). Rather, as PPgV RNA is found most abundantly in serum of infected animals, blood-borne horizontal transmission, sexual transmission and transmission from sow to piglet appear more likely. As was shown by the detection of PPgV RNA in one piglet directly after birth, before colostrum intake, intrauterine infection appears to be possible, though maybe ineffective. This suggests that horizontal transmission may be more prominent. Sexual transmission from boar to sows may be possible during artificial insemination, while transmission in both directions could occur during natural mating. Close contact and poor health condition of pigs may increase the probability of blood-borne PPgV transmission, and pig typical behavior, such as cannibalism, which can be induced by stress and high animal density, may also contribute. PPgV particles or infectious microvesicles containing PPgV genome, as they have been described in HPgV (Chivero et al., 2014), may accumulate in the blood of infected animals and could facilitate blood-borne transmission. Further evaluation of the above-mentioned PPgV transmission routes is necessary and will aid in the understanding of PPgV infection dynamics within herds.

4.7 Antibody reactivity in Western blot and ELISA

As mentioned above, PPgV Ab detection will be a helpful tool for the further investigation of prevalence in pig populations, host tropism and dynamics of the immune response in persistent and transient infections, as well as of the relevance of different antibody types and maternally derived Abs. First insights into serum Ab reactivity were obtained in this project, indicating that PPgV NS3 and E2 are viable candidates for the establishment of such assays (Chapter 3).

Mammalian expressed HPgV E2 was the preferable antigen for HPgV-specific Ab detection (Dille et al., 1997), and HPgV E2-specific Abs become detectable coinciding with clearance of viremia, are long-lasting, and provide a certain degree of protection

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from reinfection (Gutierrez et al., 1997, Tacke et al., 1997, Tanaka et al., 1998, Thomas et al., 1998). In comparison, EPgV-specific Abs were detected using bacterially expressed NS3h, and over 58% of horses tested in one study evidenced Ab reactivity with this antigen (Lyons et al., 2014). Based on these previous reports, in this project,

NS3h and C-terminally truncated E2 were produced in E. coli and evaluated for their reactivity with porcine serum Abs (Chapter 3).

High purity of PPgV NS3h was achieved by purification with IMAC in FPLC and limits background induced by unspecific bacterial proteins in Western blot and ELISA assays. As E2t was expressed in the insoluble cytoplasmic fraction (in bacterial inclusion bodies), purification under denaturing conditions followed by refolding of the protein are currently ongoing. Alternatively, alterations in bacterial expression and production of soluble E2t (i.e. induction at lower temperatures or expression in bacterial strains specifically designed for production of soluble protein) may aid in purification of the protein in its native form.

Although a lack of detectable human serum Abs against HPgV antigens other than E2 has been described (McLinden et al., 2006, Pilot-Matias et al., 1996a, Pilot-Matias et al.,

1996b, Fernandez-Vidal et al., 2007), preliminary results obtained here show that PPgV

NS3h is immunogenic. Multiple PPgV RNA positive and negative porcine serum samples evidenced reactivity with this antigen. Results found here for PPgV NS3h coincide with evidence of equine serum Ab reactivity observed with EPgV NS3h

(Lyons et al., 2014).

Comparison of Ab reactivity with PPgV NS3h versus with E2t is interesting and will shed light on which roles these antigens play in the immune response of pigs infected with PPgV. Porcine serum samples acquired here will aid in determining the dynamics of the immune response induced in PPgV infection, and it will be interesting to discern whether detection of E2t-specific Abs coincides with clearance of viremia, as is seen in

HPgV infection (Gutierrez et al., 1997, Tacke et al., 1997, Tanaka et al., 1998, Thomas et 68

Chapter 4

al., 1998). Reinfection, or reoccurrence of viremia, was observed in one animal

(Chapter 3, animal D). A possible explanation for this observation includes the production of neutralizing Abs leading to viral clearance, followed by a drop in Ab levels thus being insufficient to prevent reinfection. Alternatively, it is possible that

PPgV remained within the animal though viremia was cleared or under the detection limit of the TM qRT-PCR. In either case, this is the first description of such a reinfection

(or reoccurrence of viremia) event for PPgV infection. In HPgV, dropping of E2-Ab levels and subsequent reinfection have been described, and such causal relations between Ab levels and reinfection may also be plausible for PPgV (Devereux et al.,

1998). The circumstances under which PPgV infection is eliminated need to be further studied, including the specific mechanisms of the humoral and cellular immune response regarding clearance of the virus from the replication site and from blood.

Efforts for the validation of NS3- and E2-based Ab detection assays are currently ongoing and offer a promising step towards expanding our knowledge of pegivirus infections in pigs. Firstly, this will provide information on the seroprevalence of PPgV in pig populations from different countries, including those in which PPgV RNA has not been detected to date, which will give a more accurate estimation of the exposure of pigs to this recently discovered virus. Secondly, Ab presence in wild boar and in additional potential host species will give insights into the host tropism of Pegivirus K and into possible reservoir hosts. Lastly, dynamics and properties of the humoral immune response induced in PPgV infection can be determined using such Ab assays.

An additional, enormous advantage for the further characaterization of PPgV would be the ability to grow the virus in cell culture. Difficulties in culturing of HPgV have been described, and to date, the best results are obtained using primary human PBMCs

(Chivero and Stapleton, 2015). As mentioned above, the usefulness of porcine PBMCs will need to be evaluated.

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In conclusion, the results obtained in this project provide important first insights into the biological characterisitics of PPgV, including the virus distribution, tissue tropism, infection dynamics, and transmission routes, as well as the host immune response.

Future studies should focus on evaluating the impact that this recently discovered and widely distributed virus has on porcine health.

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Summary

5 Summary

Title: Biological characterization of porcine pegivirus

Author: Johanna Kennedy

Porcine pegivirus (PPgV) is a recently discovered member of the genus Pegivirus within the Flaviviridae family. The genus comprises a group of enveloped, positive- sense, single-stranded RNA viruses that can cause persistent infections in their hosts, but, thus far, appear to be mostly apathogenic.

In this project, the investigation of domestic pig serum samples from Europe and Asia

(n = 1,736) revealed a wide distribution of the virus and presence of PPgV RNA in 2.7% of animals and 15.2% of pig herds. In contrast, PPgV genome was not detectable in serum samples originating from wild boar hunted in Lower Saxony, Germany (n =

800). Phylogenetic analyses of a highly conserved genome region within PPgV non- structural protein 3 (NS3) revealed pairwise nucleotide identities > 90% within PPgV sequences, and suggest that PPgV is spread locally, but can also be disseminated across long distances.

In tissue samples of PPgV-viremic pigs, viral RNA was found most abundantly in the liver, thymus and in peripheral blood mononuclear cells (PBMCs), not only by qRT-

PCR, but also using fluorescence in situ hybridization. These results indicate hepato- and lymphotropism, and suggest that further investigations of possible replication sites of PPgV should focus on PBMCs and hepatocytes as possible candidates.

Close examination of the PPgV genome content in serum and excretion samples obtained from PPgV-positive pigs and their piglets longitudinally over the course of

11 months showed that shedding of the virus via excretion routes is unlikely, and that intrauterine infection appears to be possible, though it may represent a rather

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Summary

ineffective form of transmission. Instead, PPgV may be transmitted most effectively horizontally via the blood-borne or sexual routes, similar to human pegiviruses.

As assays for the detection of PPgV-specific antibodies have not been described to date, in this project, partial PPgV NS3 and E2 proteins were expressed in E. coli and evaluated for their suitability in the establishment of an antibody-detection assay.

Preliminary results indicate the reactivity of porcine serum antibodies with both proteins. The establishment and validation of an antibody enzyme-linked immunosorbent assay are currently ongoing.

Taken together, the results obtained during the course of this project provide valuable insights into the epidemiology and biological properties of porcine pegivirus infection, and the methods developed here will aid in the further characterization of this newly discovered and widely distributed virus.

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Zusammenfassung

6 Zusammenfassung

Titel: Biologische Charakterisierung des porzinen Pegivirus

Autor: Johanna Kennedy

Das porzine Pegivirus (PPgV) wurde erst kürzlich identifiziert und gehört zum Genus

Pegivirus der Familie Flaviviridae. Das Genus umfasst behüllte, positiv-strängige RNA-

Viren, die persistente Infektionen verursachen können, aber in den meisten Fällen apathogen zu sein scheinen.

In diesem PhD-Projekt wurde PPgV RNA in Serumproben von Hausschweinen aus

Europa und Asien (n = 1.736) bei 2,7% der Einzeltiere und in 15,2% der Bestände nachgewiesen, was die weite Verbreitung des Virus verdeutlicht. Hingegen konnte das Virus nicht in Serumproben von Wildschweinen aus Niedersachsen in

Deutschland (n = 800) detektiert werden. Phylogenetische Analysen konservierter

Genomregionen des PPgV Nicht-Strukturprotein 3 (NS3) zeigten innerhalb der PPgV

Sequenzen paarweise Nukleotididentitäten von über 90% und lassen vermuten, dass

PPgV sowohl regional als auch über weite Distanzen verbreitet werden kann.

Die höchsten Konzentrationen an PPgV RNA wurden mittels qRT-PCR und

Fluorezenz-in-situ-Hybridisierung in Leber- und Thymusgewebe und in peripheren mononukleären Blutzellen (PBMCs) virämischer Tiere nachgewiesen. Diese

Ergebnisse deuten auf einen Leber- bzw. Lymphtropismus hin. Aus diesem Grund bieten sich PBMCs und Leberzellen als vielversprechende Kandidaten für die zukünftige Untersuchung der Replikation von PPgV an.

Die Untersuchung von Serum und Ausscheidungen PPgV-positiver Schweine und ihrer Ferkel über einen Zeitraum von 11 Monaten hinweg zeigte, dass das Virus höchstwahrscheinlich nicht aktiv ausgeschieden wird. Intrauterine Infektionen

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Zusammenfassung

scheinen möglich, wenn auch wenig effektiv zu sein. Stattdessen wird PPgV höchstwahrscheinlich eher horizontal über Blut und venerisch übertragen, wie es auch beim humanen Pegivirus der Fall ist.

Testverfahren zum Nachweis PPgV-spezifischer Antikörper wurden bislang nicht beschrieben. Aus diesem Grund wurden in diesem Projekt Teile des PPgV NS3-

Proteins und des Hüllproteins E2 in E. coli exprimiert und deren Verwendbarkeit für die Etablierung eines serologischen Verfahrens untersucht. Erste Ergebnisse deuten darauf hin, dass porzine Serumantikörper beide Proteinen binden können. Die

Etablierung und Validierung eines PPgV-Antikörper-ELISA sind Bestandteile aktuell laufender Experimente.

Im Laufe dieses PhD-Projekts wurden wertvolle Einsichten in die Epidemiologie und die biologischen Eigenschaften der PPgV-Infektion gewonnen, und die hier entwickelten Methoden werden die weitergehende Charakterisierung dieses neu entdeckten und weit verbreiteten Virus vorantreiben.

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References

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Acknowledgements

Acknowledgements

First of all, I would like to thank Prof. Paul Becher, Prof. Karl-Heinz Waldmann and

Prof. Eike Steinmann for their excellent supervision and the very helpful supervisor meetings during the course of my PhD project, and for the opportunity of conducting this work at the Institute of Virology. Additionally, I want to thank Dr. Imke Steffen for stepping in to evaluate this thesis.

I want to mention especially Christine Bächlein, PhD – your door was always open and your support in all matters was endless – thank you!

I want to thank all my colleagues for teaching me in the ways of the virologists, and for their input during many valuable discussions. Special thanks to Inga Grotha, Doris

à Wengen and Ester Barthel, who never failed to provide help with the unending searches for samples, equipment and answers. Thanks to Dr. Alexander Postel and Dr.

Denise Meyer for many ideas and helpful comments. I would also like to thank Regina

Behre, as well as Prof. Beatrice Grummer, Dr. Tina Selle and Tanja Czeslik from the

HGNI, for their kindness and helpfulness in all PhD matters and the organization of congresses and meetings.

Thanks to the staff from the Clinic for Swine and Small Ruminants of the TiHo, especially Dr. Doris Höltig and Klaus Schlotter, and all others who cared for the pigs that were part of this project and who acquired samples. Additionally, thanks to the colleagues from the Institute of Pathology, Dr. Vanessa Maria Pfankuche, PhD,

Malgorzata Ciurkiewicz, PhD, Dr. Florian Hansmann, PhD, and Prof. Wolfgang

Baumgärtner, for their expertise in pathology and for conducting fluorescence in situ hybridization.

I want to thank our late colleague, Prof. Ludwig Haas, who was not only my favorite professor during vet school, but who inspired me to choose this career path.

Acknowledgements

Thanks also to Joe Brannan for proofreading – I hope that your somewhat random knowledge of pegiviruses will come in handy one day.

A huge thanks goes to all my friends, who made these past years special (and the downfalls bearable) through shared laughter, frustration and coffee – in particular

Kirsten Hülskötter, Dr. Lena Baron, Franziska Holzapfel, Gökce Nur Cagatay, PhD,

Nele Gremmel and Dr. Oliver Suckstorff. You made our little office on the 5th floor the coffee-haven that it is, and the past years would not have been the same without you.

Lastly, I want to thank my whole family and especially my parents, Micaela Kennedy and Jens Kunze, for their never-ending support and patience, my brother, Dr. Ricardo

Kennedy, for creating footsteps to follow in, and Lars Reise, for his constant love, compassion and understanding.