Interactions of Porcine Circovirus Type 2 and Porcine Reproductive and Respiratory Syndrome Virus Avanti Sinha Iowa State University

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2011 Interactions of porcine circovirus type 2 and porcine reproductive and respiratory syndrome virus Avanti Sinha Iowa State University

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Interactions of porcine circovirus type 2 and porcine reproductive and respiratory syndrome virus

Avanti Sinha

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Microbiology

Program of Study Committee: Tanja Opriessnig, Major Professor Rodney Butch Baker Bradley Blitvich Cathy Miller Brett Sponseller

Iowa State University

Ames, Iowa

2011

DEDICATION

I dedicate this thesis to my parents Mrs. Mitra Sinha and late Dr. Arun K. Sinha.

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TABLE OF CONTENTS

CHAPTER 1. GENERAL INTRODUCTION ...... 1 Introduction ...... 1 Dissertation Organization ...... 6

CHAPTER 2. LITERATURE REVIEW ...... 7 Porcine Circovirus type 2 (PCV2) ...... 7 Taxonomy ...... 7 Viral Characteristic, Structure and Open Reading Frames ...... 9 Viral Entry and Replication ...... 10 Clinical Manifestation of PCV2 Infection in Pigs ...... 10 Pathogenesis ...... 14 Transmission ...... 21 PCV2 and Co-infections ...... 24 Prevention and Treatments ...... 29 Porcine Reproductive and Respiratory Syndrome Virus (PRRSV) ...... 34 Taxonomy ...... 34 Clinical Manifestation, Macroscopic and Microscopic Lesions ...... 37 PRRSV Pathogenesis ...... 39 PRRSV and Co-infections ...... 42 Interaction of PCV2 and PRRSV ...... 45

CHAPTER 3. PART 1. SINGULAR PCV2A OR PCV2B INFECTION RESULTS IN

APOPTOSIS OF HEPATOCYTES IN CLINICALLY AFFECTED GNOTOBIOTIC PIGS iv

Abstract ...... 51 Introduction ...... 52 Material and Methods ...... 55 Results...... 60 Discussion ...... 62 Conclusion ...... 65 Acknowledgments ...... 66 References ...... 66 Tables and Figures ...... 71

CHAPTER 3. PART 2. SUPPLEMENTAL DATA: APOPTOSIS IS NOT A FEATURE IN

LYMPHOID TISSUES FROM PIGS CO-INFECTED WITH PRRSV AND PCV2A OR

PCV2B

Abstract ...... 75 Introduction ...... 76 Material and Methods ...... 77 Discussion ...... 80 References ...... 81

CHAPTER 4. IN VITRO INTERACTION OF PCV2A OR PCV2B WITH PRRSV WITH EMPHASIS ON PCV2 SUBTYPE SPECIFIC OPEN-READING FRAMES 1 AND 2 Abstract ...... 87 Introduction ...... 88 Material and Methods ...... 90 Results...... 98 Discussion ...... 100 v

Concluding Remarks ...... 104 Acknowledgments ...... 104 References ...... 104 Tables ...... 113

CHAPTER 5. PRRSV INFLUENCES INFECTION DYNAMICS OF PCV2 SUBTYPES PCV2A AND PCV2B BY PROLONGING PCV2 VIREMIA AND SHEDDING Abstract ...... 120 Introduction ...... 121 Material and Methods ...... 123 Results...... 130 Discussion ...... 135 Conclusion ...... 140 Acknowledgments ...... 140 References ...... 140 Tables and Figures ...... 148

CHAPTER 6. PRRSV INFECTION AT THE TIME OF PCV2 VACCINATION HAS NO IMPACT ON VACCINE EFFICACY Abstract ...... 156 Introduction ...... 157 Material and Methods ...... 159 Results...... 164 Discussion ...... 167 Acknowledgments ...... 170 References ...... 171 Tables and Figure ...... 178 vi

CHAPTER 7. GENERAL CONCLUSIONS ...... 183 REFERENCES CITED ...... 189 ACKNOWLEDGMENTS ...... 215

CHAPTER 1. INTRODUCTION

Introduction

Porcine circovirus (PCV) is a non-enveloped DNA virus that contains a single- stranded, circular genome and belongs to the genus Circovirus and family Circoviridae.

PCV was first identified in 1974 as a contaminant of the continuous porcine kidney cell line

PK-15 (Tischer et al., 1974) and later found to be non-pathogenic based on experimental inoculation of pigs (Tischer et al., 1986; Allan et al., 1995). A wasting disease in pigs known as post-weaning multisystemic wasting syndrome (PMWS) was first observed in 1991 in western Canada and later associated with a PCV found to be distinct from the non- pathogenic PCV according to nucleotide sequencing (Allan et al., 1999a; Allan et al.,

1999b). To differentiate between the two PCV variants, it was decided to use the terminology PCV1 to indicate non-pathogenic isolates and PCV2 to indicate isolates associated with disease and PMWS (Ellis et al., 1998; Hamel et al., 1998; Meehan et al.,

1998; Allan et al., 1998). Later, PCV2 was also found to be associated with respiratory disease, enteritis, reproductive failure, and porcine dermatitis and nephropathy syndrome

(PDNS), which are now collectively known as porcine circovirus associated disease

(PCVAD). Retrospectively, PCV2 has been detected in archived tissues collected as early as

1962 in Germany (Jacobsen et al., 2009) and is now found globally. PCV2 has proven to be ubiquitous in nature, causing disease in varying percentages of animals within a group while the rest of the herd typically does not display clinical signs (Balasch et al., 1999). PCV2 can be further subdivided into subtypes 2a through 2e (Olvera et al., 2006; Gagnon et al., 2007;

Dupont et al., 2008; Wang et al., 2009; Jantafong et al., 2011). Starting in late 2004,

PCVAD prevalence and disease severity increased markedly in Canada, and PCV2b, a subtype previously not recognized in North America, was identified in the majority of the cases (Gagnon et al., 2007). Similar observations were made in Iowa, Kansas, and North

Carolina where PCV2b was identified as early as 2006 (Cheung et al., 2007).

The hallmark lesion associated with PCV2 infection is lymphadenopathy, or enlargement of most lymph nodes, which correlates with lymphoid depletion at the cellular level (Sorden, 2000). The exact mechanism of PCV2-associated lymphoid depletion is still unknown and there have been contradicting results. In a conventional pig model, the authors demonstrated the presence of PCV2 antigen within apoptotic lymphocytes using PCV2 specific immunohistochemistry (IHC) and terminal transferase dUTP nick end labeling

(TUNEL) (Shibahara et al., 2000). Apoptosis is a programmed mechanism used by multicellular organisms to keep a balance between cell proliferation and cell death which can be hijacked by viruses for their own replication and dissemination purposes (Teodoro et al., 1997). Presence of PCV2 DNA was demonstrated in hepatocytes of clinically affected pigs (Rosell et al., 1999) and in a follow-up study, 88% of the livers from pigs suffering of naturally occurring PMWS contained apoptotic hepatocytes with apoptotic bodies (Rosell et al., 2000). However, contradicting results were obtained by another research group in 2004 who investigated 21 pigs categorized in three lesional severity stages and five healthy control pigs (Resendes et al., 2004b). Apoptotic rates in lymphoid tissues were correlated inversely with viral load in serum and with severity of lesions (Resendes et al., 2004b). A study conducted in gnotobiotic pigs was unable to associate PCV2 with apoptosis in hepatocytes and concluded that apoptosis was not the chief pathway for hepatocytic cell loss

(Krakowka et al., 2004). Since the exact degree of involvement of PCV2 in host cell apoptosis is unknown, it became the primary objective of the first study. Moreover, as

PCV2b was associated with perceived enhancement of PCVAD under field conditions, and no study had determined the role of PCV2b in this pathway, the second goal of the first study was to also identify a potential difference between the PCV2a and PCV2b subtypes.

Previous experimental studies found differences between the two main methods to demonstrate apoptosis in tissues, the TUNEL method and the cleaved caspase-3 (CCasp-3)

IHC staining, when investigating the role of apoptosis in lymphoid tissues (Resendes et al.,

2004a), hence, our third goal was to identify potential difference in apoptotic detection systems. PCVAD is multi-factorial and the presence of co-infections can increase and intensify clinical PCVAD (Pallarés et al., 2002; Ellis et al., 2004). Porcine reproductive and respiratory syndrome virus (PRRSV), just like PCV2, has also been associated with apoptosis in infected host cells (Choi et al., 2002; Labarque et al., 2003). Since there was no known study that has examined apoptosis rates in PRRSV and PCV2 co-infected pigs, the goal of a pilot study was to elucidate apoptotic rates in pigs dually infected with PRRSV and

PCV2.

Increased disease severity is commonly observed when pigs are co-infected PCV2 and PRRSV (Pogranichniy et al., 2002; Pesch et al., 2003; Wellenberg et al., 2004). PRRSV is an enveloped RNA virus with single-stranded, positive-sense genome that belongs to the family Arteriviridae, genus Arterivirus (Cavanagh, 1997). Two PRRSV types, 1 and 2, which are commonly referred to as European and North American types respectively, exist based on genome sequence analysis (Wensvoort et al., 1992). In breeding age animals

PRRSV infection may manifest as increased numbers of late term abortions, mummified

fetuses, stillborn piglets, and live-born weak piglets (Christianson et al., 1993; Mengeling et al., 1994). In grow-finish pigs, PRRSV usually causes pyrexia, anorexia, severe respiratory disease, and varying percentages of mortality (Rossow et al., 1994). PRRSV infection renders the host unable to clear concurrent infections resulting in increased severity of disease (Kitikoon et al., 2009; Shi et al., 2010) and it is associated with enhanced PCV2 replication (Allan et al., 2000) which results in increased amounts of PCV2 DNA in coinfected pigs (Rovira et al., 2002). Both PCV2 and PRRSV viruses have a tropism towards pulmonary alveolar macrophages (PAMs) and both are capable of infecting PAMs

(Duan et al., 1997; Duan et al., 1998; Chang et al., 2006). The ability of PRRSV to replicate in PAMs (Bautista et al., 1993) and the inability of PAMs and dendritic cells to degrade

PCV2 (Gilpin et al., 2003; Vincent et al., 2003) might lead to modification of macrophage function that could contribute to immune deregulation. The objective of the second study was to establish a PCV2-PRRSV in-vitro co-infection model. PCV2 subtypes a and b differ from each other genetically in a signature motif located between nucleotides 1472 and 1476 of open reading frame 2 (ORF2) (Cheung et al., 2007). Chimeric DNA clones have been constructed with swapped signature motif regions that were used in in-vitro and in-vivo studies to determine the significance of the signature motif in virulence and it was concluded that clones containing the reciprocal signature motif had decreased virulence (Allemandou et al., 2011). Hence, an additional goal of the second study was to determine the importance of

ORF2 and the PCV2 signature motif located in ORF2 in PCV2-PRRSV co-infection in- vitro.

As field studies have proven that concurrent infection of pigs with PRRSV and

PCV2 increases their chance of developing PCVAD (Pogranichniy et al., 2002), PRRSV-

PCV2 co-infection has become a great concern. Spread of clinical PCVAD in North

America in 2005 and 2006 has also raised important questions on PCV2 transmission throughout the swine population. PCV2 DNA has been detected in oral, nasal, urine, and fecal secretions (Magar et al., 2000; Bolin et al., 2001; Shibata et al., 2003; Segalés et al.,

2005; Caprioli et al., 2006) and horizontal transmission has been proposed and confirmed as the primary mechanism of PCV2 infection (Dupont et al., 2009; Patterson et al., 2011). As

PRRSV enhances PCV2 replication, there is a potential of increased shedding of PCV2 and increased PCV2 viral load in PRRSV-PCV2 co-infected pigs that might lead to increased transmission rates of PCV2a or PCV2b in certain pig populations. No differences in shedding of PCV2a versus PCV2b was found in experimentally infected boars (Madson et al., 2008); however, PCV2a and PCV2b shedding has not been studied in growing pigs and no investigation has been undertaken to observe the shedding patterns of PCV2 when piglets are co-infected with PRRSV. Hence the objective of the third study was to investigate the effect of PRRSV on PCV2 shedding patterns under controlled experimental conditions and to characterize differences between PCV2 subtypes.

PRRSV has been reported to decrease the efficacy of swine influenza virus (SIV) vaccination (Kitikoon et al., 2009). PCV2 vaccination is one of the most widely used vaccination strategies in growing pigs to date and existing PRRSV infections at the time of

PCV2 vaccination is of concern to swine producers and veterinarians. Not only can neonates already be infected with PRRSV, but PRRSV infected pigs can remain viremic for extended periods of time (Lager et al., 1997). Since no information existed on the efficacy of PCV2 vaccination in PRRSV viremic pigs, the objective of our fourth study was to determine the effect of PRRSV infection on PCV2 vaccine efficacy in PRRSV viremic piglets.

Dissertation Organization

The present dissertation has been prepared as an alternate manuscript format. The dissertation contains an introduction, a literature review, four separate scientific manuscripts, and a conclusion. References for the introduction, literature review, and the conclusion are cited behind each chapter.

The first manuscript describes the detection of apoptosis in hepatocytes associated with

PCV2a or PCV2b in clinically affected gnotobiotic piglets, and has been published in

Research in Veterinary Science. Supplemental data are provided on apoptotic rates in lymphoid tissues in PCV2 and PRRSV coinfected conventional pigs. The second manuscript describes PRRSV and PCV2 replication and the release of the cytokines IFN-γ and IL-10 when PAMs are infected with PRRSV in conjunction with PCV2a or PCV2b, and is in preparation. The third manuscript describes the shedding dynamics of PCV2a and PCV2b when pigs are concurrently infected with PRRSV, and has been accepted in Veterinary

Microbiology. The fourth manuscript describes the efficacy of PCV2 vaccination when administered to PRRSV viremic pigs, and is published in Clinical and Vaccine Immunology.

CHAPTER 2. LITERATURE REVIEW

I. PORCINE CIRCOVIRUS TYPE 2

1. Taxonomy

1.1. Family, genus and members

Both porcine circovirus type 1 (PCV1) and porcine circovirus type 2 (PCV2) belong to the family Circoviridae which includes two genera, namely Circovirus and Gyrovirus

(Todd et al., 2005). The genus Gyrovirus has one member, chicken anemia virus (CAV), which has a negative-sense oriented genome, unlike the members of the genus Circovirus which all have an ambisense oriented genome (Gelderblom et al., 1989). Members of the genus Circovirus are host specific and include PCV1 (Tischer et al., 1986), PCV2 (Meehan et al., 1998), beak and feather disease virus (BFDV) (Niagro et al., 1998), canary circovirus

(CaCV) (Phenix et al., 2001), pigeon circovirus (PiCV), goose circovirus (GoCV) (Todd et al., 2001), duck circovirus (DuCV) (Hattermann et al., 2003), finch circovirus (FiCV), and gull circovirus (GuCV) (Todd et al., 2007).

In recent years, PCV2 has been subdivided into two main subtypes, PCV2a and

PCV2b (Gagnon et al., 2007; Olvera et al., 2006). PCV2a and PCV2b differ in their genome sizes with 1,768 nucleotides (nt) for PCV2a and 1767 nt for PCV2b (Olvera et al., 2006).

After comparison in conventional pig models, PCV2a and PCV2b have been found to be similar in their pathogenesis (Fort et al., 2008; Harding et al., 2010; Opriessnig et al.,

2008c). Retrospective studies have indicated that PCV2a was the most prevalent subtype in the pig population prior to 2003 (Allan et al., 2007), however recent data indicated that

PCV2b was more common in PCV associated disease (PCVAD) outbreaks from 2004 onwards (Gagnon et al., 2007). Today, PCV2b is the most prevalent subtype in the worldwide pig population (Gagnon et al., 2007). Additional, less prevalent PCV2 subtypes include PCV2c which has been found in archived samples from Danish pigs (Dupont et al.,

2008), and PCV2d and PCV2e which have been described in China and Thailand (Jantafong et al., 2011; Wang et al., 2009).

According to the previous studies it is imperative to examine the ability of PCV2b to cause differential pathogenesis compared to PCV2a. It is unknown whether PCV2b infection is increased via enhancement of apoptosis in the host, increased shedding of PCV2, immunomodulating of the host response system, decreasing the efficacy of PCV2 vaccines, or any combination thereof, but many of these mechanisms are poorly understood and are important to study.

1.2. Historical background

Porcine circovirus (PCV) was first identified in the continuous porcine kidney cell line PK-15 as a contaminant in 1974 (Tischer et al., 1974). Experimental inoculation of pigs with PCV did not result in any disease, thus PCV was considered a non-pathogenic virus

(Allan et al., 1995; Tischer et al., 1986). In 1991, a new disease termed post weaning multisystemic wasting syndrome (PMWS) emerged in Canada and was associated with progressive weight loss, respiratory disease and skin discoloration in pigs right after weaning (Harding and Clark, 1998). In the late 1990s, PMWS was linked to a variant strain of the non-pathogenic PCV (Allan et al., 1999b; Allan and Ellis, 2000; Morozov et al., 1998;

Segales et al., 2005a). Analysis and comparison of the genome sequence of the PMWS-

associated PCV and the non-pathogenic PCV revealed that there were distinct genetic differences between the two strains (Ellis et al., 1998; Hamel et al., 1998; Meehan et al.,

1998) with an overall sequence nucleotide identity of roughly 68% (Hamel et al., 1998). To distinguish between the two strains, the non-pathogenic strain was designated as PCV1 and the pathogenic strain was termed PCV2 (Allan et al., 1998; Meehan et al., 1998). At present,

PCV2 is linked to PCVAD, which encompasses a variety of disease manifestations

(Gillespie et al., 2009; Opriessnig et al., 2007a) further described in Section 4.

2. Virus characteristics, structure and open reading frames

PCV2 is a non-enveloped DNA virus containing single-stranded, covalently closed circular genome that varies in between 1767 to 1768 nucleotides depending on the genotype

(Larochelle et al., 2002; Meehan et al., 1998; Tischer et al., 1982). The PCV2 genome is organized into two major open reading frames (ORFs), ORF1 and ORF2, that encode the

35.7 kDA replicase (rep) protein and the 27.8 kDa capsid (cap) protein, respectively

(Cheung, 2003b; Mankertz et al., 1998a; Nawagitgul et al., 2000). ORF2, which encodes for the cap protein epitope B-133, is used for antigenic determination and as serological marker

(Truong et al., 2001). Recently, another protein encoded by ORF3, has been identified and was associated with apoptosis (Liu et al., 2006). Specifically, the ORF3 protein binds to the

E3 ubiquitin ligase (pPirh2) thereby preventing binding of cytochrome p53, resulting in excess accumulation of p53 within the cell that evokes cytochrome c release prior to caspase activation which finally induces apoptosis (Liu et al., 2007).

3. Viral entry and replication

PCV2 enters the host cell by binding to heparin sulfate and chondritin sulfate B glycosaminoglycans (GAG) for use as attachment receptors (Misinzo et al., 2006). After binding to the GAGs, PCV2 utilizes clathrin-mediated endocytosis, requiring an acidification step, to infect the host cell (Misinzo et al., 2005). After entry into the cell,

PCV2 uses a nuclear localization signal present in ORF2 to enter the nucleus (Liu et al.,

2001). PCV2 then utilizes rolling circle replication, an intermediate double-stranded replicative form, and host replication proteins to replicate within the nucleus (Cheung,

2003b; Faurez et al., 2009; Mankertz et al., 1998b). ORF1, located on the positive-sense viral strand, is differentially spliced into two transcripts that forms Rep and Rep’ proteins respectively, and are necessary for initiation of PCV2 viral replication (Cheung, 2003a;

Mankertz and Hillenbrand, 2001). A recent study confirmed that PCV2 replication also requires activation of two mitogen activated protein kinases (MAPK), the c-Jun amino- terminal kinase (JNK) and p38 MAPK (Wei et al., 2009).

4. Clinical manifestations of PCV2 infection in pigs

PCVAD encompasses a variety of disease syndromes in pigs including: postweaning mulitsystemic wasting syndrome (PMWS), porcine dermatitis and nephropathy syndrome

(PDNS), enteritis, pneumonia, reproductive failure, and neuropathy (Harding and Clark,

1998, Done et al., 2001, Kim et al., 2004, Harms et al., 2002; Kim et al., 2003, O'Connor et al., 2001; West et al., 1999, Seeliger et al., 2007).

4.1. Postweaning multisytemic wasting syndrome (PMWS)

Systemic PCVAD, also known as PMWS, was the first recognized disease manifestation associated with PCV2 infection (Harding and Clark, 1998). The six basic clinical signs for systemic PCVAD include wasting, dyspnea, enlarged lymph nodes, diarrhea, jaundice and pallor (Cottrell et al., 1999; Harding and Clark, 1998; Harms, 1999).

Apart from these main clinical signs, other less frequently seen signs include: coughing, pyrexia, gastric ulceration, meningitis and sudden death (Harms, 1999; Wellenberg et al.,

2000). Under field conditions, PMWS occurs typically at the late nursery or early grower stage and is thought to be attributed to declining levels of passively-derived antibodies

(McIntosh et al., 2006a).

Histopathological lesions associated with PMWS consist of marked lymphocyte depletion of follicles in lymphoid tissues, the hallmark lesion of PCV2 infection. Multifocal to diffuse histiocytic infiltration is frequently observed and numerous basophilic cytoplasmic inclusions can be occasionally detected in macrophages (Rosell et al., 1999;

Segalés et al., 2004). The presence of a moderate to abundant amount of PCV2 (Sorden,

2000) which can be easily demonstrated by assays such as immunohistochemistry (IHC) or in-situ-hybridization (ISH) is also characteristic of PMWS. Furthermore, granulomatous inflammation associated with abundant amount of PCV2 antigen can be seen in non- lymphoid tissues including lungs, liver, brain, kidney, and heart.

4.2. Porcine dermatitis and nephropathy syndrome (PDNS)

Clinical signs of porcine dermatitis and nephropathy syndrome (PDNS) include the presence of cutaneous pupura on the hindlimbs, forelimbs, abdomen, head and ear skin

(Done et al., 2001). The kidneys are often enlarged and have petechial hemorrhages on the surface. The most prominent microscopic lesion described for PDNS is acute necrotizing vasculitis affecting the dermis and kidneys (Choi and Chae, 2001). A link between PCV2 and PDNS was established in 2000 by a Spanish research group (Rosell et al., 2000b), although other pathogens like PRRSV and torque teno virus (TTV) have been associated with the disease and PDNS can be reproduced experimentally without presence of PCV2

(Krakowka et al., 2008; Segalés et al., 1998).

4.3. PCV2-associated enteritis

PCV2-associated enteritis commonly affects grow-finish pigs, is characterized by diarrhea, and can clinically resemble infections with Lawsonia intracellularis and

Salmonella sp. In 2004, an association of PCV2 and enteritis was established for the first time (Kim et al., 2004). Macroscopically, there is often thickening of the ileac mucosa

(Opriessnig et al., 2011a). The most consistent microscopic lesion is the presence of extensive amounts of PCV2 associated with areas of granulomatous enteritis (Kim et al.,

2004). Moreover, PCV2-associated enteritis is characterized by infiltration of the enteric mucosa and submucosa with macrophages, histiocytes and mononuclear cells which differentiates it from enteritis caused by Lawsonia intracellularis (Jensen et al., 2006).

4.4. PCV2-associated pneumonia

PCV2-associated respiratory disease is characterized by respiratory distress, sneezing and occasionally coughing in grow-finish pigs (Harms et al., 2002; Kim et al., 2003). It is often associated with porcine respiratory disease complex (PRDC) and concurrent infection

with other pathogens (Harms et al., 2002; Kim et al., 2003; Opriessnig et al., 2004b;

Thacker, 2001). Microscopically, the presence of an interstitial pneumonia characterized by pneumocytic type 2 hypertrophy and hyperplasia, often combined with marked macrophage infiltrates that cause alveolar septa thickening and areas of necrosis in the alveolar spaces is common (Kim et al., 2003). Additionally, there can be mild necrotizing bronchiolitis and, particularly in more chronic stages, marked peribronchiolar fibrosis.

4.5. PCV2-associated reproductive failure

PCV2-associated reproductive failure consists of late term abortions, increased numbers of stillborns, and increased numbers of mummified fetuses (O'Connor et al., 2001;

West et al., 1999). Affected fetuses have myocardial necrosis associated with edema and infiltration of macrophages and lymphocytes (West et al., 1999). In fetal tissues, PCV2 targets myocardiocytes for replication (Brunborg et al., 2007; Sanchez, Jr. et al., 2003) and demonstration of an abundant amount of PCV2 antigen in heart tissues is essential for making a diagnosis (Madson et al., 2009a).

4.6. PCV2-associated neuropathy

Clinical signs for PCV2-associated neuropathy consist of neurological disorders such as inability to balance, tremors, etc. PCV2 was associated with naturally occurring neuropathy and congenital tremors by using an indirect immunofluorescent assay and ISH

(Seeliger et al., 2007; Stevenson et al., 2001). Microscopically, acute hemorrhage and edema in the cerebellum and cerebellar vasculitis have been described (Seeliger et al., 2007).

5. Pathogenesis

5.1. Immune stimulation

An experimental study conducted in gnotobiotic pigs revealed that PCV2 by itself was not able to induce clinical PCVAD and lesions; however, clinical disease and lesions were reproduced in pigs when keyhole limpet hemocyanin (KLH) in incomplete Freund's adjuvant (ICFA) was used in PCV2 inoculated pigs (Krakowka et al., 2001). The possible enhancement of PCVAD by immune system stimulation was also investigated under field conditions using routine vaccinations or immune-modulating drugs (Kyriakis et al., 2002).

The authors concluded that immune system modulation increased PCV2 replication, thus leading to increased severity of clinical disease and lesions (Kyriakis et al., 2002).

Mycoplasma hyopneumoniae (M. hyopneumoniae) and Actinobacillus pleuropneumoniae (APP) vaccinations are given to pigs at 3 to 10 weeks of age, and

PCVAD typically occurs around 8 to 12 weeks of age in the U.S.; it is possible that immune system stimulation from the above vaccines might enhance the severity of PCVAD in the conventional pigs. To test this hypothesis, a group of researchers divided 61 piglets into four groups, vaccinated two groups of piglets with commercially available M. hyopneumoniae and APP bacterins at five and seven weeks of age, and inoculated all groups with PCV2

(Opriessnig et al., 2003). Diagnostic evaluations of the vaccinated and unvaccinated pigs were performed and PCV2 viremia load, duration of PCV2 viremia, lymphoid lesions were compared. The authors found that pigs in the vaccinated group displayed more severe lymphoid depletion, longer length of PCV2 viremia and higher loads of PCV2 DNA in serum and tissues compared to the non-vaccinated group (Opriessnig et al., 2003). A Danish study included twenty 3-week old piglets from a specific pathogen free (SPF) herd which

were divided into two groups of 10 piglets, one negative control group and one group inoculated with PCV2 (Ladekjær-Mikkelsen et al., 2002). Five piglets from each group were given KLH emulsified in ICFA for immune system stimulation and the groups were compared on the basis of elevated temperature, clinical signs, PCV2 DNA in serum and

PCV2 antigen in the tissues. In contrast to the above study, it was found that development of

PCVAD occurred in both immune stimulated and non immune stimulated pigs at the same rate and that there was no significant difference between the two experimental groups

(Ladekjær-Mikkelsen et al., 2002).

Since the above studies demonstrated contradictory results, it appears to be necessary to analyze the interaction of other viruses that are predominantly found at present in U.S. farms with PCV2 vaccines in piglets and induce PCV2 vaccines failure in the US farms.

5.2. Immunosuppression

Lymphoid depletion is the hallmark microscopic lesion of PCVAD and is one of the main characteristics of PCV2 infection in pigs (Clark, 1997; Rosell et al., 1999). Areas of lymphocyte depletion have been directly correlated with the presence and magnitude of

PCV2 antigen in lymphoid tissues (Chae, 2004; Darwich et al., 2002; Dorr et al., 2007;

Opriessnig et al., 2004b; Sarli et al., 2001; Segalés et al., 2005). It has been indicated that lymphoid depletion due to PCV2 infection may be related to immunosuppression leading to secondary infections by opportunistic pathogens (Carrasco et al., 2000; Clark, 1997; Segalés et al., 2001; Segalés and Mateu, 2006). A recent study investigated genomic expression in lymph nodes collected from pigs with lymphoid depletion and demonstrated that genes

implicated in immunosuppression, such as FGL2 and GPNMB, were up-regulated (Lee et al., 2010).

Several studies focused on B-cell and T-cell distribution in PCVAD affected pigs via flow-cytometric analysis and found that there was a decrease in CD4+and/or CD8+ T- lymphocytes (Sarli et al., 2001; Segalés et al., 2001; Shibahara et al., 2000). Moreover, it has been demonstrated that there was increased IL-10 in the peripheral blood mononuclear cells

(PBMC) of PCVAD affected pigs (Crisci et al., 2010; Darwich et al., 2008; Stevenson et al.,

2006) indicative of decreased IFN-α, IFN- γ and IL-12 production (Kekarainen et al., 2008).

An in-vitro study demonstrated that PCV2 decreased the efficiency of swine alveolar macrophages by inhibiting production of oxygen-free radicals and hydrogen peroxide needed for microbiocidal activity (Chang et al., 2006). Five oligonucleotides such as CpG motifs have been identified in the PCV2 genome and might be capable of modulating the host IFN response; however, only one CpG motif of PCV2 has been associated with inhibitory effect on IFN-α (Hasslung et al., 2003). Furthermore, the inhibitory effect of

PCV2 on IFN-α was found to be associated with the ability of PCV2 to be internalized by natural interferon producing cells via TLR-9, thereby interfering with IFN-α production

(Vincent et al., 2006).

PCV2a and PCV2b differ in a signature motif located between nucleotides 1472 and

1486 of ORF2, which corresponds to a six amino acid difference in the capsid protein (Cheung et al., 2007). Previously, when utilizing chimeric DNA clones with a swapped signature motif between PCV2a and PCV2b isolates (i.e. PCV2a/b and PCV2b/a) under both in-vitro and in-vivo conditions, it was found that PCV2 a/b and PCV2b/a clones had decreased virulence when

compared to PCV2a/a and PCV2b/b, respectively (Allemandou et al., 2011). Hence, it appears to be important to examine the significance of ORF2 in a PCV2 and PRRSV co-infection model.

5.3. PCV2 and its interactions with the immune system

Numerous studies have investigated the roles of the innate, cell-mediated and humoral immune responses in pigs during PCV2 infection

i. Innate immune response. Various investigations both in-vitro and in-vivo have examined the diminutive effects of PCV2 infection on cytokines. One in-vivo study focused on the cytokine release in PCV2 infected lymphoid tissues obtained from clinically affected pigs and found an increase in IL-10 mRNA in the thymus, a decrease in IFN-γ, IL-2, IL-4, and IL-12 in secondary lymphoid tissues, and an increase of IFN-γ in the tonsils (Darwich et al., 2003b). A second investigation undertaken by the same research group demonstrated that PCV2-infected PBMC showed increased IL-10 and IFN-γ production, and when the

PCV2-infected PBMCs were challenged with mitogen they were unable to produce IFN-γ,

IL-1β, IL-2, IL-4, or IL-8 (Darwich et al., 2003a). Another in-vitro study found that PCV2 infected alveolar macrophages were unable to produce oxygen free radicals and hydrogen peroxide implying a reduction in the microbiocidal activity (Chang et al., 2006). The same work described an increase in the levels of TNF-α and IL-8 and an upregulation in the mRNAs of alveolar macrophage chemotactic factor-II (AMCF-II), granulocyte colony- stimulating factor (G-CSF), monocyte chemotactic protein-I (MCP-I) and IL-8, all of which are acute-phase chemokines (Chang et al., 2006). An in-vitro study involving swine monocytic cells demonstrated that PCV2 induced the proliferation of monocyte derived macrophages (MDM), genesis of multi-nucleated giant cells (MGC), and increased

production of MIP-I and MCP-I (Yi-Chieh Tsai et al., 2010) which all have been found to play an important role in granulomatous inflammation (Kim and Chae, 2004). It has been demonstrated that the CpG-oligodeoxynucleotide (ODN) of PCV2b had an inhibitory affect on IFN-α; however, the CpG-ODN was also involved in inhibition of mRNA expression of

IL-6, IL-10, and the p40 subunit of IL-12 (Wikstrom et al., 2011).

ii. Cell mediated immune response. Few experimental studies have investigated the cell-mediated immune system in response to PCV2 infection. One recent investigation identified that IFN-γ secreting cells, which are essential for the secretion of IFN-γ (a cytokine that directs the maturation of monocyte-derived dendritic cells and enhances their polarization of CD4 T-cells into becoming TH1 effectors), were present in pigs subclinically infected with PCV2 and correlated with decreased viral load (Fort et al., 2009). This study reinforces the importance of IFN-γ secreting cells in combating PCV2 infection in pigs.

iii. Humoral immune response. Passively acquired IgG was sufficient to prevent

PCV2 infection in piglets up to 8 weeks of age (McKeown et al., 2005). However, after the decline of the passive immunity, anti-PCV2 IgM developed and increaseed between 8 and 9 weeks of age (Carasova et al., 2007). Under experimental conditions, it has been determined that anti-PCV2 IgM appeared 10 days after PCV2 exposure and peaked around 14 days.

Anti-PCV2 IgG appeared around 14 days post PCV2 exposure and continued to be present until 140 days post exposure, the last day of the study (Opriessnig et al., 2010b). Generally, the appearance of anti-PCV2 IgG is related to immunity but has also been suggested by some as indicator for development of PCVAD as pigs that seroconvert could develop

PCVAD (Okuda et al., 2003). Development of anti-PCV2 IgG antibody under field

conditions occurs after the maternal antibodies decline, usually between 7 and 15 weeks of age (Rodríguez-Arrioja et al., 2002).

Studies utilizing gnotobiotic pigs found that the induction of a neutralizing antibody response soon after successful PCV2 infection is of extreme importance to prevent development of clinical PCVAD (Meerts et al., 2005). Neutralizing antibodies have been identified as being vital for clearing PCV2 infection, and predominantly consist of IgG1 and

IgG2 isotypes (Meerts et al., 2006). If the neutralizing antibody response is ineffective,

PCV2 infection usually progresses towards PCVAD under both natural and experimental conditions.

As outlined above, the innate immune response to PCV2 is complicated and quite involved. As pulmonary alveolar macrophages are important for the innate immune system, it seems necessary to discern whether they are affected different by PCV2a and PCV2b subtypes for example in their ability to release IL-10 and IFN-γ.

5.4. Apoptosis

Apoptosis, or programmed cell death, is maintained in multi-cellular organisms for developmental equilibrium (Horvitz, 1999). An added benefit of apoptosis is that the cell is able to intervene with viral infection and initiate the apoptotic cascade so that the virus is impaired and the spread of the virus is disrupted. Nonetheless, numerous viruses have evolved ways to hijack the programmed cell death for their own benefit by encoding proteins that are able to inhibit or initiate apoptosis in the host cell. The phenomenon of apoptosis in PCVAD was first captured in a retrospective study where it was shown that

PCV2 utilized this pathway for B-lymphocyte depletion (Shibahara et al., 2000). Presence of

TUNEL-positive lymphocytes and apoptotic bodies in the B-lymphocyte areas were confirmed (Shibahara et al., 2000). Histopathologic evidence of severe liver damage with extensive hepatocytic cell death in PCVAD affected pigs was linked with apoptosis (Rosell et al., 2000a). However, contradictory results have been reported. For example, the reduction in B-cells has been attributed to the lack of T-cell and B-cell expansion rather than apoptosis (Mandrioli et al., 2004). In another research study conducted in PCV2-infected gnotobiotic pigs using TUNEL staining, the authors concluded that apoptosis induced by

PCV2 may not be the cause of lymphocyte depletion or hepatocyte loss in PMWS affected pigs (Krakowka et al., 2004). When both TUNEL staining and cleaved caspase-3 staining were used on formalin-fixed, paraffin-embedded lymphoid tissues, it was found that the apoptotic proportions were inversely correlated to the PCV2 load in the serum (Resendes et al., 2004). The severity of lesions was suggestive of more severe lymphoid depletion associated with lower apoptotic rates (Resendes et al., 2004). Interestingly, the same research group found that a higher level of apoptosis was present in hepatocytes that contained more PCV2 antigen; thus, concluding that the hepatitis observed in PCVAD pigs is caused by apoptosis of hepatocytes (Resendes et al., 2010). Incidentally, the ORF3 protein has been associated with apoptosis in the host according to several studies conducted in recent years (Liu et al., 2005; Liu et al., 2006; Liu et al., 2007). The ORF3 protein has been connected with apoptosis through its binding to porcine ubiquitin E3 ligase pPirh-2 which facilitated p53 expression and ultimately apoptosis of the host cell (Liu et al., 2007).

The contradictory study results obtained in the past should be used to drive further research to verify the exact roles of both PCV2a and PCV2b subtypes in causing apoptosis in pigs.

6. Transmission

6.1. Horizontal transmission

The main transmission route of PCV2 is the fecal-oral route (Segalés et al., 2005b).

Several research groups investigated the shedding characteristics of PCV2 from PCVAD affected experimentally-infected pigs. A controlled study conducted in gnotobiotic piglets concluded that PCV2 DNA is shed in nasal secretions and feces (Krakowka et al., 2000).

The transmission of PCV2 was investigated in specific pathogen free pigs intranasally inoculated with PCV2 and it was confirmed that nasal secretions are a means of transmitting

PCV2 (Magar et al., 2000). In another study, PCV2 was isolated from the nasal swabs, rectal swabs, tonsillar swabs, serum and urine of caesarian-derived, colostrum-deprived pigs, and direct contact of infected pigs with naïve pigs resulted in infection (Bolin et al., 2001).

When both experimental and natural PCV2 infection were investigated, PCV2 was found to be easily transmitted to naïve nursery pigs under field conditions and PCV2 was detected in various samples immediately after experimental inoculation (Shibata et al., 2003). Another study completed to determine if a PCR method can accurately detect PCV2 DNA in feces found that PCV2 is shed in high quantities in feces and suggested that feces might be important in the transmission of PCV2 (Yang et al., 2003).

A study analyzing the presence of PCV2 DNA in the nasal cavities, PCV2 viremia, and PCV2 antibody titers in pigs from several farms found that pigs in PCVAD affected farms had higher amounts of PCV2 DNA in the nasal cavities as well as serum compared to pigs from farms that were subclincially infected with PCV2 (Sibila et al., 2004). Similar conclusions were reached in another investigation where the authors examined the presence

of PCV2 DNA in serum, tonsillar, nasal, trachea-bronchial, urinary and fecal swabs of

PCV2-infected pigs and determined that PCVAD affected pigs had higher levels of PCV2

DNA in their nasal secretions, oral secretions, urine and fecal samples (Segalés et al.,

2005b). Colostrum-deprived pigs were experimentally infected with PCV2 at five weeks of age and PCVAD affected pigs had higher PCV2 DNA levels in peripheral blood leukocytes, serum, plasma, blood, fecal and tonsillar swabs (Caprioli et al., 2006). It was found that

PCV2 is transmitted easily through close pig-to-pig contact and that nose-to-nose contact might be a primary route of transmission (Dupont et al., 2009). A study on horizontal PCV2 transmission in conventional pigs indicated that PCV2 DNA can be detectable for 181 days in serum and for 69 days in nasal, oral and fecal specimens, concluding that naturally acquired PCV2 infections persist long term (Patterson et al., 2011a). Another study conducted by the same authors affirmed that horizontal transmission of PCV2 via nasal secretions, saliva, and feces is possible for at least 69 days after PCV2 infection (Patterson et al., 2011b).

Research undertaken previously demonstrated that PCV2 is shed in high quantities in nasal, oral and fecal contents in pigs and probably contributes to horizontal transmission in field situations. Therefore it appears to be important to analyze if there is subtype specific difference between PCV2a and PCV2bin viral shedding and transmission that might be associated with the higher prevelance of PCV2b in U.S. farms.

6.2. Vertical Transmission

i. Breeding herds. The presence of abundant PCV2 antigen associated with severe lesions in the cardiac muscle of fetuses and piglets born to sows with natural PCV2 infection

has initially demonstrated that abortions and stillbirths can occur in association with PCV2 infection (West et al., 1999). In follow up case reports, similar results were seen (Ladekjær-

Mikkelsen et al., 2001; O'Connor et al., 2001) indicating that vertical transmission of PCV2 occurs under certain situations. An experimental study where fetuses were inoculated with

PCV2 at different days of gestation resulted in typical myocardial changes associated with

PCV2 (Sanchez, Jr. et al., 2001). PCV2 antigen was found in various tissues obtained from mummified fetuses, stillborn piglets and subclinically infected piglets after intrauterine

PCV2 experimental infection of the fetuses (Johnson et al., 2002). A recent study has confirmed that fetuses inoculated with PCV2a or PCV2b had similar high titers of PCV2a and PCV2b (Saha et al., 2010). Investigators have been able to prove that conventional sows with high levels of PCV2 DNA in their serum but low anti-PCV2 antibody titers gave birth to viremic piglets, of which 25% eventually died (Calsamiglia et al., 2007). Real-time quantitative PCR was performed using feces from 100 PCVAD-affected conventional pigs and 101 PCVAD non-affected conventional pigs to identify a possible correlation between

PCV2 in fecal contents of lactating sows and suckling piglets (McIntosh et al., 2008). The authors found that PCVAD affected sows had higher quantities of PCV2 DNA in their feces which corresponded to higher PCV2 DNA in the feces of their piglets (McIntosh et al.,

2008). Presence of PCV2 in the milk of six sows infected experimentally with PCV2 indicated that PCV2 can be transmitted via milk to naïve piglets thus increasing PCV2 transmission in suckling pigs (Ha et al., 2009).

ii. Boars. An experimental study examined the detection rate of PCV2 DNA in boar semen and found that boars inoculated with PCV2 intranasally showed intermittent shedding of PCV2 in semen (Larochelle et al., 2000). When nested PCR was used to determine the

presence of PCV2 DNA in semen of naturally PCV2 infected boars, the presence of PCV2

DNA was found to be sporadic and continued for at least 27.3 weeks post-inoculation

(McIntosh et al., 2006b). Finally, possible differences in shedding patterns of PCV2a and

PCV2b in boar semen were investigated, and it was determined that high quantities of both

PCV2a and PCV2b DNA were detectable in serum and semen samples from experimentally infected boars (Madson et al., 2008). When sows were inseminated with semen spiked with

PCV2, dam and fetal infections occurred (Madson et al., 2009b; Madson et al., 2009a).

7. PCV2 and co-infections

Early studies have demonstrated that PCV2 is the etiological agent in the pathogenesis of PCVAD (Bolin et al., 2001; Ellis et al., 1999; Ladekjær-Mikkelsen et al.,

2002). An observational study established that PCV2 was consistently detected in clinically affected pigs and should be considered as the etiological agent for development of PCVAD

(Kim et al., 2002). However, experimentally, PCVAD has only been reproduced very efficiently with PCV2 in conjunction with other pathogens, and it is well established that the severity of disease in co-infected pigs is much more severe compared to pigs infected with

PCV2 alone.

7.1. Field investigations

Two conventional 2.5 month old pigs that suffered from clinical PCVAD were found to be co-infected with PCV2 and pseudorabies virus (Rodríguez-Arrioja et al., 1999). A retrospective study conducted on 484 U.S. field cases diagnosed with PCVAD found that

PRRSV was present in 52% of the cases, M. hyopneumoniae was present in 36% of the

cases, swine influenza virus was present in 5% of the cases, and PCV2 as the only pathogen was demonstrated in 2% of the cases (Pallarés et al., 2002). In Korea, investigation of 1,634 pig cases resulted in presence of concurrent PCV2 and Hemophilus parasuis infection in

32% of the cases and PCV2 and PRRSV in 29% of the cases (Kim et al., 2002). Detection rates of viral pathogens in proliferative and necrotizing pneumonia under field conditions in a retrospective study found PRRSV in 92%, PCV2 in 42%, swine influenza virus (SIV) in

2%, and a combination of PRRSV and PCV2 in 42% of the investigated cases (Drolet et al.,

2003).

A case control study conducted in the Netherlands involving 60 PCVAD pigs determined that 83% were co-infected with PRRSV. PCVAD and non-PCVAD pigs were evenly distributed among pigs co-infected with transmissible gastroenteritis virus (TGEV), porcine parvovirus (PPV), porcine respiratory coronavirus (PRCV) and SIV subtype H1N1

(Wellenberg et al., 2004). A field study involving Spanish indoor pig herds concluded that concurrent infection of pigs with PRRSV, SIV, and PCV2 was quite common in the herds thus predisposing the herds to development of clinical disease (Lopez-Soria et al., 2010). A

U.S. case-control study concluded that porcine enterovirus (PEV) is equally distributed between controls and cases and not involved in PCVAD (Pogranichniy et al., 2002).

A German case study done on 102 farms with a total of 171 samples observed that

8% of the pigs suffering from interstitial pneumonia were positive for SIV which was presumed to be a secondary pathogen and a consequence of PCVAD (Pesch et al., 2003). A case control study comprised of 69 PCVAD cases observed concurrent PCV2 and SIV infection in 55% of the pigs (Engle and Bush, 2006).

During the investigation of the prevalence of porcine torque teno virus (TTV) in

Spanish PCVAD pigs, it was determined that TTV was present along with PCV2 in PCVAD affected pigs (Kekarainen et al., 2006). A study conducted at a Swedish farm investigated the presence of PCV2, porcine TTV-1 or TTV-2, and bocavirus in PCVAD affected pigs and demonstrated that 71% of the clinically affected pigs contained all the three viruses

(Blomstrom et al., 2010). In Serbia a research work investigated conventional pigs that had died of PCVAD and established that the hepatitis in these pigs was associated with concurrent infection of PCV2, HEV and TTV (Savic et al., 2010).

A field study found that singular PCV2 infection was associated with enteritis; however, PCV2 was also indentified as a co-factor with Lawsonia intracellularis in ileitis

(Jensen et al., 2006). A recent case study investigated occurrence of bronchiolitis obliterans organizing pneumonia in PCVAD pigs and found PCV2 to be involved in this particular lesion (Cheng et al., 2011). Tests for other pathogens such as Pasturella multocida,

Streptococcus sp., Salmonella sp., APP, M. hyopneumoniae, PRRSV, pseudorabies virus

(PRV), classical swine fever virus (CSFV), PPV, cytomegalovirus, PEV, PCV1, PRCV, and

SIV were negative (Cheng et al., 2011).

The field investigations confirmed that concurrent infection with other pathogens increases PCVAD prevalence and severity in pigs and hence it appears to be imperative to examine the interaction of PCV2a or PCV2b with PRRSV under experimental conditions and to evaluate the effect of the later on the two subtypes of PCV2.

7.2. Experimental investigations

Several experimental studies have been conducted to understand the role of co- infection in PCVAD and are further described in the following section.

i. PRRSV. This is covered in: III. Interaction of PCV2 and PRRSV.

ii. Porcine parvo virus (PPV). When gnotobiotic pigs were experimentally infected with PCV2, incidentally it was found that a portion of the pigs were also infected with PPV which resulted in clinical PCVAD (Ellis et al., 1999). Several follow-up experimental studies were performed to further investigate the role of PPV in PCVAD. When colostrum- deprived piglets were co-infected with both PCV2 and PPV severe clinical disease was observed, whereas PPV or PCV2 alone were not able to produce any clinical signs and only mild lesions (Allan et al., 1999a). There was also wider tissue distribution of PCV2 antigen in the co-infected groups compared to the singularly infected pigs (Allan et al., 2000c).

When microscopic lesions and PCV2 antigen distribution in pigs singularly infected with

PCV2 or co-infected with PCV2 and PPV were compared, PCV2 and PPV co-infection was associated with severe microscopic lesions in a variety of organs compared to mild lesions in selected organs in pigs infected with PCV2 alone (Kennedy et al., 2000). A study involving segregated early weaned piglets investigated whether PCV2 and PPV co-infection augments PCVAD and whether vaccination against PPV was protective in the co-infected piglets (Opriessnig et al., 2004a). Clinical disease in the co-infected groups was mild-to- severe compared to the singular infected group that did not show any clinical signs. The study concluded that PCV2 viremia was increased and persisted for longer time in the coinfected group; however, PPV vaccination was not able to prevent clinical disease

(Opriessnig et al., 2004a).

iii. M. hyopneumoniae. An experimental study was conducted using 67 pigs to establish the effect of concurrent infections of M. hyopneumoniae and PCV2. The study determined that M. hyopneumoniae prolongs PCV2 infection and that dual infected pigs developed severe PCVAD, a direct contrast to singularly inoculated groups which only displayed mild clinical disease (Opriessnig et al., 2004b).

iv. SIV. SIV and PCV2 are often implicated in field cases of PRDC. A recent study undertaken by a group of researchers who wanted to verify a potential synergistic relationship between PCV2 and SIV, concluded that even though the PCV2-SIV co-infected group developed severe disease, no significant differences were found between the co- infected and singularly infected groups (Wei et al., 2010).

v. Porcine torque teno virus (TTV). Genotype 1 of porcine TTV has been implicated by some groups in potentiating PCVAD. One experimental study confirmed that pigs singularly infected with PCV2 or TTV group failed to show clinical disease whereas dual infected pigs developed clinical illness (Ellis et al., 2008).

vi. Lawsonia intracellularis. PCVAD is sometimes accompanied by diarrhea in field cases. Pigs that were singularly infected or co-infected with PCV2 and Lawsonia intracellularis developed enteritis implying that PCV2 was also capable independently cause enteritis (Opriessnig et al., 2011a).

vii. Salmonella typhimurium. When the effect of Salmonella typhimurium in PCVAD was investigated, it was found that Salmonella spp. antigen was detected sporadically along with PCV2 antigen in the same macrophages; however, there was no potentiation of PCV2 or PCVAD (Opriessnig et al., 2011a).

viii. Porcine epidemic diarrhea virus (PEDV). Field observations have indicated that

PCV2 and PEDV co-infection can cause severe disease in neonatal piglets. To further investigate this, neonatal piglets that were congenitally pre-exposed to PCV2 were infected with PEDV (Jung et al., 2006). The authors determined that PCV2 and PEDV co-infection caused severe histopathological lesions and prolonged clinical manifestation compared to the piglets infected with either PCV2 or PEDV alone and that PCV2 potentiates PEDV replication in the piglets (Jung et al., 2006).

8. Prevention and treatment

8.1. Commercially available vaccines

There are several commercial PCV2 vaccines available on the global market and each product has been shown to be very efficient in controlling PCVAD. The studies investigating PCV2 vaccine efficacy have been divided into field and experimental studies in this review.

i. Field studies. A field study was conducted to investigate the efficacy of a baculovirus-expressed PCV2 ORF2 subunit vaccine (Ingelvac® CircoFLEXTM; Boehringer

Ingelheim Vetmedica, Inc.) in a PRDC-affected pig herd (Fachinger et al., 2008). Sows were vaccinated against PPV, erysipelas, E. coli, Clostridium perfringens type A, and

Haemophilus parasuis before the study was initiated and piglets were divided into two groups: vaccinated (n=769) and non-vaccinated (n=773). The study demonstrated that, under field conditions, PRDC manifestation can be reduced if pigs are vaccinated against PCV2

(Fachinger et al., 2008). A different field study was conducted to evaluate the performance of another commercially available, baculovirus-expressed, PCV2-ORF2 subunit vaccine

(Circumvent® PCV; Intervet, Inc.) involving 485 pigs that were divided into vaccinated

(235) and nonvaccinated (250) pigs (Horlen et al., 2008). The authors found that there was dramatic reduction in PCV2 viremia and mortality in the vaccinated group and that the average daily weight gain in the vaccinated group was considerably higher compared to the control group (Horlen et al., 2008).

ii. PCV2 vaccination in the singular PCV2 challenge model. In a study undertaken to investigate the serological response in CIRCOVAC® (Merial, Inc.) vaccinated gilts, 23

PCV2 seronegative gilts were divided into two groups, a vaccinated and a non-vaccinated group (Charreyre et al., 2005). The piglets born to these gilts were challenged with PCV2 and then monitored for clinical signs, antibody development and the presence of lesions at the termination of the study. The authors concluded that the CIRCOVAC® (Merial, Inc.) vaccine had efficiently protected the piglets born to the vaccinated gilts (Charreyre et al.,

2005). Seventy-two piglets obtained from a commercial farm divided into vaccinated (4 groups) and non-vaccinated groups (4 groups) were used in an experimental study that examined the efficacy of Porcilis® PCV (Intervet Inc.) vaccination by challenging the pigs with one of four distinct PCV2 isolates (Fort et al., 2008). The study found that vaccination was able to induce a strong humoral response with high titers of neutralizing antibodies in the vaccinated pigs while reducing lymphoid lesions and PCV2 viremia (Fort et al., 2008).

Since PCV2 is a ubiquitous virus and most sows have developed antibodies, an experimental study examined the presence and effect of maternal antibodies against PCV2 in piglets (Opriessnig et al., 2008b). The study used sixty 4-week old pigs divided into four groups containing piglets with varying levels of PCV2 antibodies. It was concluded that

Suvaxyn® PCV2 (Fort Dodge Animal Health Inc.) effectively protected the piglets from

PCV2-associated diseases and thus exhibited that maternal antibodies did not interfere with the PCV2 vaccine (Opriessnig et al., 2008b). A recent experimental study that analyzed the efficacy of commercially available vaccines used Suvaxyn® PCV2 (Fort Dodge Animal

Health Inc.), Ingelvac® CircoFLEX™ (Boehringer Ingelheim Vetmedica Inc.) and

Circumvent™ PCV2 (Intervet Inc.) and 60 SPF pigs divided into six groups (Opriessnig et al., 2009). Circumvent™ PCV2 and Suvaxyn® PCV2 were used as either single or two dose administration. After vaccination, all the groups except the negative group were challenged with PCV2 and monitored for the presence of clinical signs, presence and amount of PCV2

DNA in serum and tissues, and presence of antibodies against PCV2. No significant differences were identified between any of the vaccinated groups. All vaccines used induced high titer of neutralizing antibodies and, as a result, the amount of PCV2 DNA in sera and tissues was significantly reduced compared to the positive control group (Opriessnig et al.,

2009). Another recent study compared the effectiveness of passive versus active vaccination in the piglets. Seventy-eight piglets born to PCV2-vaccinated and non-vaccinated sows were divided into groups, vaccinated with different commercial vaccines (Ingelvac®

CircoFLEX™ and CIRCOVAC®) and challenged with PCV2b (Opriessnig et al., 2010a).

The study concluded that regardless of vaccination scheme, vaccinated pigs were protected against PCV2 challenge (Opriessnig et al., 2010a).

iii. PCV2 vaccination in the co-infection model. Since PRRSV and PCV2 co- infection is common, an experimental study examined the efficacy of PCV2 vaccination in pigs with concurrent PRRSV and PCV2 infection using 74 piglets divided into two vaccinated groups and four non-vaccinated groups (Opriessnig et al., 2008a). The vaccine used in this study was Suvaxyn® PCV2 One Dose (Fort Dodge Animal Health, Inc.) and it

was administered intramuscularly or intradermally. The study demonstrated that the vaccine was able to limit PCV2 infection by inducing a strong humoral response against PCV2

(Opriessnig et al., 2008a). Another co-infection model included one-dose and two-dose commercial vaccine regimens and an experimental vaccine and used 73 PCV2 viremic piglets. The pigs were divided into two control groups and five vaccinated groups that were challenged with PCV2, PRRSV and PPV (Shen et al., 2010). Vaccines used in the study were Suvaxyn® PCV2 (Fort Dodge Animal Health Inc.), Ingelvac® CircoFLEX™

(Boehringer Ingelheim Vetmedica Inc.) and Cicumvent™ PCV2 (Intervet Inc.) and a live chimeric PCV2 vaccine. The authors determined that all the vaccines were effective in reducing the PCV2 DNA load in the serum (Shen et al., 2010). Considering that M. hyopneumoniae is commonly present in field cases along with PCV2 infection and it potentiates PCV2 replication, a study was performed to investigate the efficacy of PCV2 vaccination in boars that were concurrently infected with PCV2 and M. hyopneumoniae

(Opriessnig et al., 2011b). Twelve boars were divided into four groups; two groups vaccinated and two groups were inoculated with M. hyopneumoniae. All groups were challenged with PCV2. The authors concluded that PCV2 vaccination was efficacious in reducing viremia in the vaccinated groups with considerably less shedding of PCV2 in the semen of vaccinated boars (Opriessnig et al., 2011b).

Commercially available vaccines have been shown to be efficacious in both field studies and under experimental conditions, in both a singular PCV2 challenge model and in the co- infection model. However, no work has been done to determine the efficacy of these vaccines when the animals are in the acute stage of infection at the time of vaccination, thus

it seemed imperitive to study the affect of an acute infection, such as PRRSV, on the efficacy of commercially available vaccines

8.2. Experimentally available vaccines

A study that used experimental DNA and subunit vaccines based on PCV2 ORF2 was able to demonstrate that the vaccinated pigs were protected from subsequent PCV2 challenge and that PCV2 replication was prevented (Blanchard et al., 2003). In another study, the authors produced their own vaccine by cloning ORF2 of PCV2a into a PCV1 infection clone, thereby creating a chimeric PCV1-2a vaccine candidate that they tested in conventional pigs challenged with PCV2 (Fenaux et al., 2004). The study concluded that the chimeric virus was able to provide protection against the PCV2 challenge. A study that investigated the efficacy and safety of autogenous adjuvanted PCV2 vaccines utilized tissue homogenates of PCVAD affected pigs and found that the autogenous PCV2 vaccines contained infectious PCV2 (Baker et al., 2009). Vaccine efficacy was not investigated in this study (Baker et al., 2009). In a different study the authors constructed a chimeric PCV2 vaccine candidate based on PCV2 subtype b and tested the efficacy of the vaccine in conventional pigs (Beach et al., 2010). The authors found that both PCV2a and PCV2b load was significantly reduced in challenged pigs compared to non-vaccinated pigs (Beach et al.,

2010).

8.3. Strategies to control PCVAD

A very important method of reducing PCVAD is by maintaining good management practices. Madec proposed a 20-point plan (Madec et al., 1999; Madec et al., 2000) for

PCVAD affected farm that includes the farrowing units, post-weaning facilities and grow-

finish facilities. 1) All-in-all-out production system with thorough cleaning and disinfecting of the barn between batches. 2) Treatment for parasites in sows before farrowing as well as washing the sows 3) Limited cross-fostering. 4) Small post-weaning pens should be separated by solid partitions. 5) Emptying, cleaning and disinfection of pens at a regular basis. 6) Stocking should be only 0.33 m2 /pig. 7) Feeder space should be more than 7 cm. 8)

Air quality at levels of less than 10 ppm ammonia, less than 0.1% CO2 and humidity less than 85%. 9) Control temperature. 10) Avoid mixing batches. 11) Small grow-finish pens separated by solid partitions. 12) Emptying, cleaning and disinfection of pens at regular basis in grow-finish barns with all-in-all-out rule. 13) No mixing of post-weaning pigs. 14)

No mixing between the finishing pens. 15) Stocking should be only .75 m2 /pig. 16) Good air and temperature. 17) Vaccination regimen should be appropriate. 18) Controlled airflow in the buildings and animal flow within buildings. 19) Stringent hygiene protocol should be followed. 20) Ill animals should be removed or euthanized quickly.

Apart from the above requisite steps to keep PCVAD under control it is also essential to investigate the effect of PCV2 vaccinations in piglets that are already PRRSV viremic at 3-week old age.

II. PORCINE REPRODUCTIVE AND RESPIRATORY SYNDROME VIRUS

(PRRSV)

1. Taxonomy

1.1. Family, genus, members

PRRSV is an enveloped RNA virus containing positive-sense, single-stranded genome belonging to the family Arteriviridae and the genus Arterivirus of the order

Nidovirales (Cavanagh, 1997; Meulenberg et al., 1993). Genomic analysis of sequences showed existence of two PRRSV types, type 1 and type 2, commonly known as European and North American genotypes respectively, which share 64-67% nucleotide sequence identity (Meng et al., 1995; Nelsen et al., 1999). The other three members of the genus

Arterivirus include equine arteritis virus (EAV), lactate dehydrogenase elevating virus

(LDV), and simian hemorrhagic fever virus (SHFV) (Snijder and Meulenberg, 1998).

1.2. Historical background

PRRSV was first identified as the etiological agent of porcine reproductive and respiratory syndrome (PRRS) in 1991 (Benfield et al., 1992; Wensvoort et al., 1991) after the disease had been known as “mystery disease” in the U.S. (Keffaber, 1989). PRRSV causes high economic losses to the worldwide swine industry which was estimated in 2005 to equal 560 million dollars in the U.S. alone (Neumann et al., 2005).

1.3. Viral characteristics and structure

i. Viral Entry. PRRSV enters porcine monocytic cells in lymphoid tissues and the lungs (Duan et al., 1997a; Duan et al., 1997b). Multiple studies have been conducted on the entry of the virus into its permissive cells over the last years. Heparan sulfate was one of the first entry mediators to be identified for attachment to pulmonary alveolar macrophages

(Delputte et al., 2002). Heparan sulfate is present on most cells and is part of the proteoglycans that form the glycosaminoglycans, which are important for adhering and modulating the action of extra-cellular ligands. However, heparan sulfate is not the only attachment mediator. Sialoadhesin, another transmembrane glycoprotein, is also able to bind

to PRRSV (Vanderheijden et al., 2003). An α 2-3-linked sialic acid on the PRRSV envelope has been implicated in successful PRRSV infection of pulmonary alveolar macrophages since the sialic acids on the membrane protein/glycoprotein-5 interact with the sialoadhesin receptors on pulmonary alveolar macrophages and trigger the cell to internalize the virus

(Van et al., 2010a; Van et al., 2010b). After attachment, PRRSV is taken in the cell via the clathrin-mediated endocytosis which moves the virus into an early endosome. In the early endosome, PRRSV interacts and binds to CD163, a scavenger receptor, potentially via glycoproteins 2 and 4, to uncoat the viral envelope (Das et al., 2010). This results in lowering of the pH further enabling PRRSV to uncoat its genome (Nauwynck et al., 1999).

ii. Replication and open reading frames (ORF). Several researchers have worked on elucidating the mechanism of replication of PRRSV. PRRSV has a positive-sense single- stranded RNA genome which has a length between 13 to 15 kilobases (Kb) and contains eight ORFs (Snijder and Meulenberg, 1998). ORF1 encodes for the replicase protein, which functions as RNA-dependent-RNA-polymerase that comprises three fourths of the PRRSV genome, and is located at the 5’ end of the genome (Snijder and Meulenberg, 1998). ORF1 is subdivided into the ORF1a and ORF1b regions that encode for 13 non-structural proteins

(nsp) that have replicase, polymerase and protease functions. Recently, nsp1α, nsp1β, nsp2, nsp4 and nsp11, have demonstrated their ability to inhibit an IFN response and are now considered to be important in immune evasion (Beura et al., 2010). ORF2 through ORF7, located at the 3’ end of the PRRSV genome, give rise to a nested set of six subgenomic mRNAs with a common leader sequence at their 5’ end from which the structural proteins are encoded (Snijder and Meulenberg, 1998). ORF2 is further subdivided into ORF2a and

ORF2b which code for glycoproteins 2a and 2b respectively, minor structural proteins that

are N-glycosylated (Wu et al., 2001). Glycoproteins 3 and 4, encoded by ORF3 and ORF4, are also N-glycosylated minor structural proteins (Meulenberg, 2000; Wu et al., 2005).

ORF5 encodes for glycoprotein 5 which is another N-glycosylated protein and which represents the major structural protein present on the envelope. Glycoprotein 5 is disulfide bonded to the non-glycosylated matrix (M) protein, the other major structural protein that traverses the viral membrane three times, and is encoded by ORF6 forming a heterodimer which is necessary for PRRSV infection (Mardassi et al., 1996; Snijder et al., 2003). ORF7 encodes for the nucleocapsid (N) protein which is found bound to the genome of PRRSV

(Yoo and Wootton, 2001) and is the most immunogenic of all PRRSV proteins (Murtaugh et al., 2002).

2. Clinical manifestation, macroscopic and microscopic lesions

The severity and clinical manifestation of disease associated with PRRSV depends on the age (Mengeling1998a, Van der Linden et al 2003) and breed of infected pigs (Halbur et al., 1998; Vincent et al., 2005) as well as the virus genotype and virulence (Christopher-

Hennings et al., 2001; Halbur et al., 1995; Halbur et al., 1996; Mengeling et al., 1996;

Mengeling et al., 1998).

2.1. Breeding age females

Breeding age females infected with PRRSV may display late term abortions, increased numbers of mummified fetuses, stillborn piglets, and live-born weak piglets, or show delayed return to estrus (Christianson et al., 1993; Mengeling et al., 1994). In addition,

PRRSV is able to cross the placenta and infect fetuses where it has been shown to replicate

primarily in the fetal thymus (Rowland, 2010). The piglets that survive fetal PRRSV infection may develop chronic PRRSV infection with virus persisting in tonsil and lymph nodes and can serve as a source for PRRSV spread (Rowland et al., 2003).

2.2. Suckling pigs

PRRSV infected neonatal and nursery pigs commonly display tachypnea, dyspnea, pyrexia, anorexia and lethargy in varying degrees that can be associated with increased mortality rates (Rossow et al., 1994; Rossow, 1998). PRRSV associated enteritis, characterized by profuse diarrhea, is a common finding in PRRSV infected suckling piglets.

2.3. Growing pigs

Growing pigs infected with PRRSV exhibit clinical signs such as sneezing, nasal discharge, fever, and decreased weight gain. PRRSV often culminates in severe respiratory disease that can cause growth impediment and death (Johnson et al., 2004). Gross lung lesions vary depending on the PRRSV isolate involved and concurrent infection with other viruses or bacteria. Lung lesions are usually characterized by failure to collapse and discoloration ranging from no coalescence to diffusely mottled-tan lung surfaces (Done and

Paton, 1995; Halbur et al., 1996; Rossow et al., 1995). Lymph nodes can be enlarged.

Microscopic PRRSV lesions in lung tissues are characterized by hyperplasia of type 2 pneumocytes and septal thickening due to the presence of mononuclear cells and alveolar effusion consisting of macrophages (Collins et al., 1992; Halbur et al., 1995; Halbur et al.,

1996).

3. PRRSV pathogenesis

PRRSV infection is characterized by prolonged presence of the virus in the host and is associated with persistant existence in the host lymph nodes and tonsils. PRRSV is also able to cross the placenta and infect fetuses. Several mechanisms have been implicated for this.

3.1. Apoptosis

The ORF5 encoding glycoprotein 5 of PRRSV has been implicated in apoptosis both in vitro and in vivo (Sirinarumitr et al., 1998; Suarez et al., 1996; Sur et al., 1998). A recent study has elucidated the process via which PRRSV initiates apoptosis (Lee and Kleiboeker,

2007). The study demonstrated the presence of phosphatidylserine on the outer leaflet of infected cell which represented the early apoptotic stage. Caspase 8, 9 and 3 activities were demonstrated and indicated apoptosis along with the presence of PRRSV infection in the same cells. The study concluded that Bax/Bcl-2 proteins of the programmed cell death family of proteins were involved in the apoptosis of the cells infected with PRRSV (Lee and

Kleiboeker, 2007).

3.2. Immune suppression

PRRSV was found to have a tropism towards alveolar macrophages whose function the virus alters resulting in enhanced concurrent infection with other pathogens (Galina et al., 1994; Renukaradhya et al., 2010; Van et al., 2001). It appears that PRRSV infection renders the host unable to clear concurrent infections resulting in increased severity of disease (Allan et al., 2000a; Kitikoon et al., 2009; Shi et al., 2010; Thanawongnuwech et al.,

2000). However, contradicting these reports, other research groups have not found any correlation between PRRSV and immunosuppression (Albina, 1997; Beilage, 1995; Drew,

2000).

3.3. Mechanisms of immune evasion

The immune response to PRRSV infection is very complicated as PRRSV has acquired several mechanisms for immune evasion. This is an important reason why a successful vaccination protocol has not been accomplished to date and PRRSV eradication still proves to be quite difficult.

i. Innate immune response. PRRSV is capable of down-regulating the innate immune response in the host by the inhibition of interferons such as IFN-α, IFN-β, TNF-α and IL-1 which are the first cytokines to be released in host defense against viral infections (Van et al., 1999; Van and Nauwynck, 2000). PRRSV has dexterously employed tactics that inhibit the induction of IFN-α and IFN-β at the mRNA levels by its non-structural proteins (Beura et al., 2010; Chen et al., 2010). At different levels, the nsp1α, nsp1β, nsp2, nsp4 and the nsp11 all inhibit the IFN-β promoter activation (Beura et al., 2010). Both IRF-3 and NFκB were activated by the TLR-3 signaling pathway when PRRSV used TLR-3 as receptor; however, nsp1β of PRRSV specifically inhibited both IRF-3 and NFκB activation thus subverting the host innate immune system (Beura et al., 2010). When IL-1β, IL-8 and IFN-γ were induced during early PRRSV infection, the host was capable of eliminating the virus successfully from the system (Lunney et al., 2010). PRRSV was also found to be involved in the up-regulation of IL-10 in dendritic cells which circumvented the immune response towards a TH2 response (Charerntantanakul et al., 2006; Flores-Mendoza et al., 2008).

ii. Cell mediated immune response. IL-1β and IFN-α are pro-inflammatory cytokines that are necessary for activation of T-cells to exhibit cytotoxicity and secrete IFN-γ.

However, since PRRSV inhibits pro-inflammatory cytokines such as CD-25, TNF-α remains at a minimal level, and the T-cells are not only not activated but appear suppressed

(Charerntantanakul et al., 2006). Specifically, it was found that active PRRSV infection in monocyte-derived dendritic cells caused down-regulation of MHC class I, MHC class II,

CD-14 and CD11b/c, and T-cells were not activated (Wang et al., 2007). PRRSV specific

IFN-γ cells appeared a relatively long time after initial PRRSV infection with the majority being CD4+/CD8+ cells whereas the CD8+ cytotoxic cells, which are necessary for viral clearance, were present in very low numbers (Meier et al., 2003).

iii. Humoral immune response. An appropriate humoral immune response against

PRRSV is complicated since both innate and the cellular immunity, the first steps towards an adaptive immune response, is not activated properly. Antibodies directed against the

PRRSV N and M proteins appear at 7-9 days post-infection and are non-neutralizing

(Labarque et al., 2000; Lopez et al., 2007). Neutralizing antibodies develop later and those directed against GP-5 are the most protective (Ansari et al., 2006). An extensive study examined the glycosylation sites on the GP-5 domain to determine how mutations affected the immune response and formation of neutralizing antibodies (Plagemann et al., 2002). It was found that N30, N33, and N44/N51 mutations resulted in high titer of neutralization antibodies against PRRSV (Plagemann et al., 2002), which did not occur when glycans were present. It was postulated that PRRSV uses decoy epitopes and that PRRSV infection and shedding would not be prevented since antibodies form against these epitopes and are not neutralizing (Fang et al., 2006; Ostrowski et al., 2002).

PRRSV has a cellular tropism towards alveolar macrophages that can cause immune suppression in hosts and thus enhance concurrent infection, PRRSV is capable of inhibiting pro-inflammatory cytokines such as IFN-α, IFN-β, TNF-α and IL-1 that are necessary for viral eradication by host immune and up-regulates IL-10 that suppresses immune response in the hosts, and PRRSV also circumvent the production of neutralizing antibodies in the host against itself thus rendering the host defenseless against an onslaught from PRRSV infection and as well as secondary infections. It is therefore very important to examine the interaction of PRRSV with PCV2 and to analyze if the immune suppression induced by

PRRSV would eventually lead to higher PCV2 load in the host and increased severity of

PCVAD.

4. PRRSV and co-infections

Coinfections of pigs with PRRSV and other pathogens occur commonly under field conditions. A selection of PRRSV co-infections studies which were investigated under experimental conditions are summarized below.

4.1. M. hyopneumoniae

An in-vitro study was conducted to examine proinflammatory cytokines produced by pulmonary alveolar macrophages during PRRSV and M. hyopneumoniae co-infection. In this study, the authors used tracheal ring ex-plants from pigs infected with M. hyopneumoniae and pulmonary alveolar macropahges that were infected with PRRSV singularly and as well as co-cultured with M. hyopneumoniae and PRRSV for 24 hours

(Thanawongnuwech et al., 2001). Semi-quantitative RT-PCR and ELISAs were used to

detect the mRNA of soluble cytokines. It was found that the dual infected group had increased levels of IL1α, IL1β, IL8 and TNF mRNA. It was concluded that the production of pro-inflammatory cytokines like IL1α, IL1β, IL8 and TNF-α possibly increases the severe and chronic pneumonia experienced under field conditions (Thanawongnuwech et al., 2001).

An in-vivo study by the same group investigated the cytokine profile in pigs dually infected with PRRSV and M. hyopneumoniae (Thanawongnuwech and Thacker, 2003). Seventy 2- week old crossbred pigs negative for antibodies to PRRSV and M. hyopneumoniae were divided into four treatment groups consisting of negative controls, pigs inoculated with M. hyopneumoniae, pigs inoculated with PRRSV and pigs co-infected with PRRSV and M. hyopneumoniae. ELISA were used to determine protein levels of IFN-γ and IL-10 in the bronchoalvolear lavage fluid. The authors found that IL-10 was released in increased levels in the dually infected as well as singularly infected groups demonstrating a TH2 biased immune response (Thanawongnuwech and Thacker, 2003).

4.2. PRCV and SIV

The potential of PRRSV to interfere in the replication process of porcine respiratory corona virus (PRCV) and SIV was investigated in a study (Van et al., 1996). Thirty-six conventional 10-week old pigs were divided into negative controls, PRRSV infected pigs,

PRCV infected pigs, SIV infected pigs, PRRSV- PRCV co-infected pigs, and PRRSV-SIV co-infected pigs. The authors found that dually inoculated groups displayed more severe disease than the pigs infected with PRRSV alone; however, PRRSV did not interfere with

PRCV and minimally affected SIV replication (Van et al., 1996). The same research group conducted another study using conventional pigs and cesarean-derived, colostrum-deprived

pigs for singular and dual infections with PRRSV and SIV in three different experiments

(Van et al., 2001). In the first experiment, 19 pigs were divided into four groups with one group serving as negative controls. The three remaining groups were inoculated with

PRRSV which was followed by H1N1 inoculation three days later, seven days later, or 14 days later. In the second experiment, 11 pigs were assigned to three groups. Again, one group served as the negative control group and the two remaining groups were inoculated with PRRSV followed by H1N1 inoculation three or seven days later. In the third experiment, 15 cesarean-derived, colostrum-deprived pigs were divided into three groups.

Two groups were inoculated with either PRRSV or H1N1 influenza virus. The last group was inoculated with PRRSV and seven days later with H1N1. Overall, the conventional pigs displayed higher body temperatures and weight loss, however, Streptococcus suis was isolated from one of the pigs in this group. The cesarean-derived, colostrum-deprived pigs did not show severe clinical disease in the dual infected groups and hence, the authors were unable to conclude if dual infection causes more severe disease in this type of pigs (Van et al., 2001).

4.3. Streptococcus suis

When the effect of concurrent S. suis infection on PRRSV was investigated, it was found that the animals had a greater susceptibility to S. suis septicemia and disease, which was possibly due to immune suppression exerted by the PRRSV infection

(Thanawongnuwech et al., 2000). The study used 80 3-week old pigs that were divided into six groups with one negative control group, a group vaccinated with a PRRSV modified live vaccine, a group infected with S. suis, a PRRSV-modified live vaccinated groups infected

with S. suis, a group infected with PRRSV, and a group infected with PRRSV and S. suis.

The authors found that the PRRSV-S. suis dual infected group displayed severe clinical disease compared to all the other groups with a mortality rate of 88% (Thanawongnuwech et al., 2000). Another study used piglets which were divided into four groups, with one group intranasally inoculated with S. suis type 2, one group infected with PRRSV, and one group inoculated with PRRSV followed by S. suis. The fourth group served as uninoculated control group. The authors discovered that the dually infected PPRSV- S. suis pigs displayed a more severe form of clinical disease than did the singularly infected pig groups (Galina et al., 1994).

It is well known that PRRSV is able to cause severe disease in combination with selected other pathogens. It istherefore necessary to analyze the interaction of PRRSV and

PCV2 to confirm if co-infection with PRRSV and PCV2 causes increased PCV2 load in the host that may lead to increased PCVAD severity .

III. INTERACTION OF PCV2 AND PRRSV

1.1. Field studies

A Spanish study investigated the presence of PRRSV in cases of PCVAD and concluded that PRRSV potentiates PCVAD manifestation and increases disease severity compared to pigs infected with PCV2 alone (Segalés et al., 2002). A case control study done in the Netherlands found that 83% of the PCVAD pigs were dually infected with PCV2 and

PRRSV, whereas 35% of the clinically healthy pigs were also co-infected with PCV2 and

PRRSV (Wellenberg et al., 2004). Concurrent PCV2 and PRRSV infection was related to a more severe clinical presentation compared to PCV2 alone infection (Wellenberg et al.,

2004). In a retrospective Spanish study, porcine proliferative and necrotizing pneumonia cases were investigated (Grau-Roma and Segalés, 2007). The authors found that PCV2 alone was associated with proliferative and necrotizing pneumonia in Spain; however, PCV2 and PRRSV were found in 41% of the proliferative and necrotizing pneumonia cases (Grau-

Roma and Segalés, 2007). This study is in contrast to a study done in Canada where 60 lungs with lesions consistent with proliferative and necrotizing pneumonia were investigated

(Drolet et al., 2003) and 42% (55/60) of the cases were positive for both PRRSV and PCV2.

It was determined that PRRSV alone can cause proliferative and necrotizing pneumonia as

92% (55/60) of the lungs were positive for PRRSV without involvement of PCV2 (Drolet et al., 2003). Twenty-eight Italian pigs suffering from proliferative and necrotizing pneumonia were selected and 43% (12/28) of the pigs were positive for PCV2 antigen, 68% (19/28) of the pigs were positive for PRRSV, and 29% (8/28) of the pigs were co-infected with PCV2 and PRRSV. Hence, it was concluded that proliferative and necrotizing pneumonia in Italy was due to co-infection of pigs with PRRSV and PCV2, even though the prevalence of

PRRSV in affected pigs was higher (Morandi et al., 2010).

An investigation of 31 cases and 56 controls obtained from the U.S. field assessed the relationship of PCV2 and major swine viruses (Pogranichniy et al., 2002). No significant difference in the distribution of PPV, PEV, SIV, PRCV and porcine lymphotropic herpesvirus type 1 (PLHV-1) was found between cases and controls. On the other hand,

PCV2 had the highest association with PCVAD, and it was concluded that pigs with PRRSV and PCV2 co-infections had the highest odds ratio in developing PCVAD (Pogranichniy et al., 2002). An epidemiological study investigated the infection dynamics of PCV2 and

PRRSV in 10 swine producing systems with PCVAD (Calsamiglia et al., 2003). It was

determined that when a farm was experiencing concurrent infection with both PCV2 and

PRRSV, the disease severity in the pigs increased significantly (Calsamiglia et al., 2003). A separate epidemiological assessment of co-infecting pathogens investigated 583 pigs obtained from a single integrated pig production system in the U.S. (Dorr et al., 2007). The group constructed several age models to examine the timing of co-infection of PCV2 with different pathogens and concluded that PRRSV and PCV2 co-infection was most prominent in 3-week old piglets (Dorr et al., 2007). In 2003, Pesch et al. investigated the presence of

PCV2, PRRSV, PPV, Chlamydia suis, SIV, M. hyopneumoniae and APP in PCVAD affected pigs from 102 different farms. The pathogens most commonly identified were

PRRSV and PCV2 and 60% of the cases were found to be co-infected with PCV2 and

PRRSV. When these two viruses circulated at the same time in a farm then the incidence of

PCVAD-associated disease increased (Pesch et al., 2003).

1.2. In vivo experiments

Concurrent PCV2-PRRSV infection was investigated in colostrum-deprived piglets

(Allan et al., 2000b). Seventeen piglets were randomly divided into four groups: five control pigs, three pigs inoculated with PCV2, four pigs inoculated with PRRSV, and five pigs inoculated with PCV2 and PRRSV. It was found that there was a higher prevalence of PCV2 antigen positive pigs in the group inoculated with both PCV2 and PRRSV compared to the singularly inoculated groups leading to the conclusion that PRRSV potentiated PCV2 replication in the dually infected groups (Allan et al., 2000b). In another experimental study using the cesarean-derived-colostrum deprived pig model, sixty-one 3-week old pigs were divided into four groups including a negative control (n=12), a PCV2 inoculated group

(n=19), a PRRSV inoculated group (n=13) and a PCV2-PRRSV dual inoculated group

(n=17) (Harms et al., 2001). The results of this study demonstrated that PCV2 and PRRSV co-infected group displayed more severe lymphoid depletion and interstitial pneumonia compared to the singular PCV2 and PRRSV infected groups. Another study investigated

PCV2-PRRSV dual infection in twenty-four 5-week old conventional pigs which were distributed into a control group (n=5), a PRRSV inoculated group (n=5), a PCV2 inoculated group (n=7) and a dual infected PRRSV-PCV2 (n=7) group (Rovira et al., 2002). The piglets in the PRRSV group and the dual infected PRRSV-PCV2 group were infected with

PRRSV one week prior to PCV2 challenge. The PRRSV-PCV2 dual infected group developed a more severe form of PCVAD compared to the other groups and there was a higher load of PCV2 DNA in the serum compared to the group infected with PCV2 alone.

In-situ hybridization (ISH) demonstrated that more cells positive for PCV2 nucleic acid and more severe microscopic lesions were found in the PRRSV-PCV2 co-infected group compared to the singular PCV2 inoculated group. This study demonstrated that when conventional pigs are infected with both PRRSV and PCV2, resulting lesions can be quite severe (Rovira et al., 2002).

1.3. In vitro experiments

Few in vitro experiments have been conducted to investigate the interaction of

PRRSV and PCV2 in pulmonary alveolar macrophages as this cell type is the known in vivo target cell for PRRSV. A research group established the in vitro interaction of PCV2 and

PRRSV and their singular and combined effect on alveolar macrophages (Chang et al.,

2003). The authors found that in contrast to PRRSV infection, PCV2 singular infection did

not induce any cytopathic effect. The dually-infected group showed a decreased infectious rate and cytopathic effect compared to the group singularly infected with PRRSV and the authors concluded that PCV2 reduced the infectivity and replication of PRRSV in the alveolar macrophages (Chang et al., 2003). Experimental work by the same group further addressed the in vitro interaction of PRRSV and PCV2 and examined the IFN-α and TNF-α production in the cell culture supernatant at an interval of 18, 36, 54, 72, 90, and 108 h post- infection (Chang et al., 2005). The authors determined from the results that PCV2 induced a potent IFN-α secretion from the alveolar macrophages cells which might inhibit PRRSV action in the PCV2-PRRSV group compared to the PRRSV alone group. The authors also concluded that PCV2 was a weak inducer of TNF-α whereas PRRSV was a strong inducer of TNF-α. Concurrent infection of PRRSV and PCV2 possibly reduces PRRSV infection in alveolar macrophages which might elevate the pulmonary inflammatory response in the pigs, thus increasing the severity of pneumonia (Chang et al., 2005). The same research group conducted another experimental study where they investigated PCV2-PRRSV concurrent infection and the expression of Fas and Fas ligand on the surface of splenic lymphocytes, macrophages and peripheral blood lymphocytes (Chang et al., 2007). It was observed by the authors that the splenic lymphocytes and macrophages in the dually infected group displayed a significantly higher amount of Fas and FasL expression on the cell surfaces compared to the group singularly infected with PCV2. Therefore, lymphocyte depletion in vivo might partly be the result of apoptosis due to Fas and FasL expression

(Chang et al., 2007).

Field studies, and in-vivo and in-vitro experimental studies have proven that PRRSV causes increased PCV2 replication, increased PCVAD severity, and a higher incidence of

PCVAD. It appears imperative to examine and analyze the interactions of PRRSV and

PCV2 in the host to detect PCV2 vaccine failure caused by PRRSV, increased PCV2 pathogenesis, and/or if PRRSV is involved in any immune-modulation that would increase

PCV2 shedding and transmission in naïve populations.

CHAPTER 3: SINGULAR PCV2A OR PCV2B INFECTION RESULTS IN

APOPTOSIS OF HEPATOCYTES IN CLINICALLY AFFECTED GNOTOBIOTIC

PIGS

A paper published in

Research in Veterinary Science doi:10.1016/j.rvsc.2010.10.013, 2010

Avanti Sinha, Shayleen Schalk, Kelly M. Lager, Chong Wang, Tanja Opriessnig

ABSTRACT

Porcine circovirus type 2 (PCV2) is clinically associated with respiratory disease, failure-to- thrive, hepatitis, and diarrhea; however, the precise pathogenesis of PCV2-associated disease and in particular its involvement in apoptosis is still controversial. The objectives of this study were (1) to determine whether PCV2 is associated with apoptosis by examining and comparing hepatic tissues from clinically affected or unaffected gnotobiotic pigs that were experimentally infected with PCV2, (2) to determine if there are differences between

PCV2a and PCV2b in inducing hepatocyte apoptosis, and (3) to determine if there are differences between apoptosis detection systems. Forty-eight gnotobiotic pigs were separated into five groups based on inoculation status and development of clinical disease:

(1) sham-inoculated, clinically-unaffected (n=4), (2) inoculated with PCV2a, clinically- unaffected (n=10), (3) inoculated with PCV2a, clinically-affected (n=6), (4) inoculated with

PCV2b, clinically-unaffected, (n=13) and (5) inoculated with PCV2b, clinically-affected

(n=15). Formalin-fixed, paraffin-embedded sections of liver from all pigs were analyzed for signs of apoptosis [presence of single strand DNA breaks in the nucleus by the terminal transferase dUTP nick end labeling (TUNEL) assay or presence of intra-nuclear cleaved caspase 3 (CCasp3) demonstrated by CCasp3 immunohistochemistry (IHC)]. In addition, the liver tissues were also tested for presence of cytoplasmic and intra-nuclear PCV2 antigen by an IHC assay. Specific CCasp3 and TUNEL labeling was detected in the nucleus of hepatocytes in PCV2a and PCV2b infected pigs with significantly (P < 0.05) higher levels of apoptotic cells in clinically-affected pigs. Regardless of PCV2 genotype (PCV2a;

PCV2b), there were higher levels of PCV2 antigen in clinically-affected pigs compared to clinically-unaffected pigs. There was no significant difference in detection rate of apoptotic cells between the TUNEL assay and CCasp3 IHC. When high amounts of PCV2 antigen were present, the incidence of CCasp3 and TUNEL staining also increased regardless of the

PCV2 genotype. This suggests that PCV2-induced apoptosis of hepatocytes is important in the pathogenesis of PCV2-associated lesions and disease.

1. Introduction

Porcine circovirus (PCV) is a small, non-enveloped, circular, single stranded DNA virus. There are two main types of PCV defined as PCV1 and PCV2 (Meehan et al., 1998).

Phylogenetic analysis of PCV2 isolates has shown that PCV2 can be further divided into two main genotypes, PCV2a and PCV2b, which are both distributed worldwide (Gagnon et al., 2007; Olvera et al., 2007).

Initially, PCV2 was identified in cases of post-weaning multisystemic wasting syndrome (PMWS) in Canada in 1991 (Ellis et al., 1998; Harding and Clark, 1997). PCV2 is associated with several disease manifestations in pigs commonly summarized as porcine circovirus associated disease (PCVAD) (Opriessnig et al., 2007). PCVAD includes systemic infection in nursery age pigs commonly characterized by wasting and general failure to thrive (Chae 2004), hepatic disease (Rosell et al., 2000a), respiratory disease (Harms et al.,

2002), enteric disease (Kim et al., 2004), reproductive failure (Ladekjær-Mikkelsen et al.,

2001; O’Connor et al., 2001) and porcine dermatitis and nephropathy syndrome (PDNS)

(Rosell et al., 2000b). PCVAD has since emerged globally (Segalés et al., 2005) and now has become one of the most economically important diseases throughout pig producing countries (Opriessnig et al., 2007).

Currently there are contradicting reports about whether or not PCV2 plays a vital role in inducing apoptosis in various pig tissues. Examples of viruses that induce apoptosis include: adenovirus, human immunodeficiency virus (HIV-1), porcine reproductive and respiratory syndrome virus (PRRSV), influenza virus A/B, rotavirus (Roulston et al., 1999),

African swine fever (Oura et al., 1998), classical swine fever (Summerfield et al., 1998) and many others. Microscopically, apoptosis can be demonstrated by dUTP labeling of single strand DNA breaks in the nucleus (Krakowka et al., 2004) or by demonstrating the presence of cleaved caspase 3 (CCasp3) by immunohistochemistry (IHC) staining (Resendes et al.,

2004a, b). A novel open reading frame (ORF3) was identified in PCV2 infected PK15 cells in 2005. It was discovered that ORF3 protein is not essential for viral replication but is involved in activating caspase-8 and caspase-3 pathways which in turn induce apoptosis

(Liu et al., 2005).

A study using naturally infected conventional pigs found that within lymphoid tissues, apoptosis of lymphocytes had occurred, which was determined by the use of the terminal transferase dUTP nick end labeling (TUNEL) stain (Shibahara et al., 2000). Within the same study PCV2 was stained via IHC and it was found that PCV2 was present in the nuclei of macrophages and apoptotic lymphocytes, as well as apoptotic bodies phagocytosed by macrophages, providing evidence that PCV2 was the cause of B lymphocyte depletion

(Shibahara et al., 2000). Similarly in BALB/c mice experimentally inoculated with PCV2, it was found that PCV2 replicated in the spleen, lymph nodes and Peyer’s patches of infected mice and that the replication of PCV2 within these tissues was associated with apoptosis which was confirmed by the TUNEL assay (Kiupel et al., 2005). It was also noted that upregulation of caspase 8 and 3 in the spleens of infected mice might be the mechanism through which PCV2 induces apoptosis in mice (Kiupel et al., 2005).

In contrast, a recent study using pigs naturally affected with PCVAD noted that there was an inverse relationship between amounts of PCV2 antigen and apoptosis in the thymus and most peripheral lymphoid tissues (Resendes et al., 2004b). It was also found that pigs with severe lymphoid depletion had less apoptosis (Resendes et al., 2004b). In Italy, pigs with varying clinical presentations of PMWS were classified into three classes based on the severity of the disease presentation (Mandrioli et al., 2004). Apoptosis in lymphoid tissues was analyzed and there was a significant decrease in apoptosis in lymphoid tissues when control pigs were compared to pigs at the initial stage of infection; however, no significant differences were found in pigs in the intermediate and final stages of the disease (Mandrioli et al., 2004).

PCV2 has been found to replicate in a number of different tissues and the induction of severe immunosuppression using cyclosporine in PCV2 infected gnotobiotic piglets increased the replication of PCV2 within hepatocytes (Krakowka et al., 2002). Livers from

100 pigs with naturally occurring PMWS were examined for lymphohistiocytic hepatitis, and apoptotic bodies; of these 100 livers 88% were found to have apoptosis and 12% had no microscopic liver lesions. It was also determined that hepatocytes were a target for PCV2 infection and replication (Rosell et al., 2000a). Similarly another study using 15 pigs from five different farms with previous confirmation of PMWS found PCV2 antigen and DNA in hepatocytes of the livers from all pigs that were PCV2 positive (Rosell et al., 1999).

The objectives of this study were to determine (1) whether PCV2 is associated with apoptosis of hepatocytes by comparing clinically affected or unaffected gnotobiotic pigs that were experimentally infected with PCV2, (2) if there are differences between PCV2a and

PCV2b in inducing hepatocyte apoptosis, and (3) if there are differences between apoptosis detection systems exist.

2. Materials and methods

2.1. Pig source and housing

Forty-eight gnotobiotic piglets acquired surgically from crossbred sows free of

PCV2 were used. Directly after surgical removal pigs were placed into sterile stainless steel isolators which were mounted with polyvinyl canopies and maintained under positive pressure. The isolator units were cleaned with sodium hypochlorite and water before sterilization with ethylene oxide prior to the start of the experimental study. Isolators were sustained at a temperature of 32 ˚C and the temperature was progressively decreased to a

subsistence temperature of 27 ˚C with HEPA filtration of the intake and exhaust air. The pigs were initially fed a pasteurized milk diet from a commercially available source

(Esbilac® Kansas City, Kansas, USA) three times per day, with increasing quantities and a reduction to feeding twice a day to a maximum of 370 ml per feeding. All animal manipulations were approved by the Institutional Animal Care and Use Committee

(IACUC) according to the guidelines set by the National Animal Disease Center. The pigs were tested on a weekly basis and found to be negative for aerobic and anaerobic bacteria from rectal swabs by routine culture methods. In addition, all pigs tested negative for porcine parvovirus (PPV), PRRSV, swine hepatitis E virus, and bovine viral diarrhea virus

(BVDV) as determined by routine PCRs on serum collected on the day of necropsy.

2.2. Inoculation

The pigs were inoculated intranasally between seven and 10 days of age. Control pigs (n=4) were inoculated with 1 ml PCV2-free cell culture medium. The remaining pigs were either inoculated with 1 ml of PCV2a strain DQ629114 (Cheung et al., 2007) at a dose

-2 -4 of 10 to 10 TCID50/ml (PCV2a, n=16) or with 1 ml of PCV2b strain DQ629115 (Cheung

-2 -4 et al., 2007) at a dose of 10 to 10 TCID50/ml (PCV2b, n=28). The infectious dose of the inocula was determined by IHC as previously described (Cheung, 2003).

2.3. Study design

For the purpose of this study, the pigs were separated based on inoculation status and development of clinical disease resulting in five groups: (1) Sham-inoculated, clinically- unaffected (n=4), (2) PCV2a, clinically-unaffected (n=10), (3) PCV2a, clinically-affected

(n=6), (4) PCV2b, clinically-unaffected, (n=13) and (5) PCV2b, clinically-affected (n=15).

Liver tissue from all 48 pigs was collected at the time of necropsy between 23 and 41 days post inoculation, evaluated for microscopic lesions, and scored for presence of apoptotic cells as determined by the TUNEL assay or CCasp3 IHC stain and for presence of PCV2 antigen by IHC.

2.4. Tissue preparation

At necropsy several sections of different liver lobes were collected and immediately immersed in 10% buffered formalin. After the tissues were fixed for 24 h, the tissues were dehydrated, paraffin-embedded, sectioned at 4 µm and mounted on a glass slide followed by routine hematoxylin/eosin staining as described (Krakowka et al., 2004).

2.5. Histopathology

Microscopic lesions were evaluated by a veterinary pathologist (TO) blinded to treatment groups. The sections of liver were evaluated for the presence of lymphohistiocytic inflammation (0=none; 3=severe), suppurative hepatitis (0=none; 3=severe) and presence of hepatocyte degeneration (0=none, 3=diffuse, severe) and areas of necrosis (0=none,

3=diffuse, severe). The individual lesion scores were summed and the lesion sum scores were compared between groups.

2.6. Detection of PCV2 antigen

IHC for detection of PCV2-specific antigen was performed using a rabbit polyclonal antiserum as described previously (Sorden et al., 1999). PCV2-antigen scoring was done by

a veterinary pathologist (TO) blinded to treatment groups. The following scoring protocol was used: 0=no signal or no visible stain, 1=staining was observed in individual inflammatory cells, 2=staining was observed in moderate numbers of inflammatory cells and hepatocytes, 3=abundant staining in inflammatory cells and hepatocytes. Although this system is subjective, the same veterinary pathologist scored treatment groups in an earlier study in which a high correlation was found with digital computer image analysis

(Opriessnig et al., 2004).

2.7. TUNEL assay

In brief, liver tissue sections were dewaxed, rehydrated in graded alcohol, incubated in 3% hydrogen peroxide for 10 min at 22 ºC and rinsed twice with phosphate buffered saline (PBS). This was followed by treatment with Proteinase K (Ambion, Inc., Foster City,

USA) for 20 min at 30º C. After two washing steps with PBS, the tissues were incubated in

10% sheep serum (Sigma, St. Louis, Missouri, USA) for 20 min at 22 ºC and rinsed twice with the PBS. TUNEL staining was performed using a commercially available kit (in-situ cell death detection kit, POD; Roche Diagnostics, Indianapolis, Indiana, USA). The manufacturers’ instructions were followed and DAB chromogen (Dako North America Inc.,

Carpentaria, California, USA) was applied for 5 min and then counterstained with hematoxylin.

2.8. Cleaved Caspase-3 (CCasp3) immunohistochemistry (IHC)

The tissue sections were dewaxed, rehydrated in graded alcohol and incubated in 3% hydrogen peroxide for 10 min at 22 ºC, rinsed three times in Tris buffered saline (TBS)

treated with 0.05% protease (Sigma Aldrich) for 2 min and washed three times with TBS buffer. A CCasp3 IHC (Cell Signaling Technology®, Danvers, Massachusetts, USA) was applied at a concentration of 1:100 for 1 h at 22 ºC. Further processing was done by using a streptavidin-biotin detection kit (Dako North America Inc.).

2.9. Apoptosis controls

Known negative and positive controls were included in each TUNEL and CCasp3 run. As a positive control, sections of murine intestines from a mouse experimentally inoculated with live mycobacterium paratuberculosis followed by dextran sulfate sodium treatment were used (Courtesy of Dr. C. Johnson). This procedure produced colitis in the mouse which resulted in epithelial cell apoptosis as described previously (Vetuschi et al.,

2002).

2.10. Apoptotic cell scoring

Apoptotic cell counts were performed by scoring 10 random hepatic areas using a

25× lens and a 10× eyepiece, yielding a final magnification of 250× with an Olympus® microscope (Leeds Precision Instrument, Inc; North Minneapolis, Minnesota, USA). The apoptotic cell score ranged from 0 to 3 (0=average of 0.0-0.9 apoptotic cells/10 fields,

1=average of 1-4.9 apoptotic cells/10 fields; 2=average of 5-14.9 apoptotic cells/10 fields;

3=average of 15 or more apoptotic cells/10 fields).

2.11. Statistical Analysis

The statistical analysis on the data was performed using the SAS software version

9.2, SAS system for windows (SAS Institute Inc., Cary, North Carolina, USA). A Kruskal-

Wallis non-parametric analysis of variance (ANOVA) was used to determine differences in median scores between groups. A P value of 0.05 or less was considered statistically significant. If significant, association between categorical variables in paired situations was evaluated by using Fisher’s exact test.

3. Results

3.1. Microscopic lesions

Microscopic lesions in liver sections were observed in 88.6% (39/44) of the experimentally inoculated pigs, no microscopic lesions were observed in sham inoculated pigs (Fig. 1). Hepatic lesions were characterized by infiltration with low-to-high numbers of mixed inflammatory cells (mainly lymphocytes, macrophages, and a few neutrophils). In addition, there was mild-to-severe vacuolar degeneration and loss of hepatocytes in 58.3%

(28/48) of the pigs (Fig. 1). Affected pigs also had severe lymphoid depletion and histiocytic replacement of follicles in lymphoid tissues (tonsil, spleen, inguinal lymph node, tracheobronchiolar lymph node (data not shown). Regardless of clinical disease status and genotype, PCV2 inoculated pigs had significantly (P < 0.003) more severe lesions compared to sham-inoculated pigs (Table 1). The association between disease status (clinically- affected, unaffected) and lesion sum score was significant (P = 0.003) and clinically- affected pigs had significantly (P = 0.001) more severe lesions compared to clinically-

unaffected pigs. No significant (P = 0.228) difference of lesion score was observed between pigs inoculated with PCV2a or PCV2b (Table 1).

3.2. PCV2 antigen in liver

Low to high amounts of PCV2 antigen was detected in 24/48 pigs (Table 1; Fig. 1).

There were significantly (P < 0.001) higher amounts of PCV2 antigen in clinically-affected pigs when compared to clinically-unaffected pigs; however, there was no significant (P =

0.402) difference between pigs inoculated with PCV2a and PCV2b (Table 1). PCV2 antigen was detected in the cytoplasm of macrophage-like cells in areas of inflammation but also in the cytoplasm and nuclei of hepatocytes.

3.3. Detection of apoptotic cells by the TUNEL assay

The distribution of the final TUNEL scores in the different treatment groups (sham- inoculated, PCV2a, PCV2b) is summarized in Table 2 and representative images are shown in Fig. 1. Twenty-two of 48 (45.8%) liver tissues had a score of 0, 22.9% (11/48) had a score of 1, 20.8% (10/48) had a score of 2 and 10.4 % (5/48) had a score of 3. There was a significant (P < 0.001) association between disease status (clinically-affected, unaffected) and TUNEL scores and a significant (P < 0.001) association between lesion sum scores and

TUNEL scores. Overall, TUNEL scores were significantly (P < 0.001) higher in clinically- affected pigs (Table 1). There was also a significant (P < 0.001) association of TUNEL scores and PCV2 IHC scores with an increase in TUNEL score associated with a higher

PCV2 IHC score (Table 3). There was no significant (P = 0.886) association between

TUNEL scores and PCV2 genotype (PCV2a, PCV2b) (Table 1). Comparison of apoptotic

rates as determined by TUNEL assay at different DPIs showed that there was no age dependent difference in the severity of apoptosis (data not shown).

3.4. Detection of apoptotic cells by CCasp3 immunohistochemistry

The distribution of the final scores in the different treatment groups is summarized in

Table 2 and representative images are shown in Fig. 1. Twelve of 48 (25.0%) pigs had no detectable CCasp3 staining in sections of liver (score 0), 39.6% (19/48) of the pigs were found to have low amounts of CCasp3 staining (score 1), 18.8% (9/48) had moderate amounts of CCasp3 staining (score 2), and 16.7% (8/48) of the pigs had large amounts of

CCasp3 staining in sections of liver (score 3). Clinically-affected pigs had significantly (P <

0.001) higher CCasp3 IHC scores compared to clinically-unaffected pigs. There was also a significant (P = 0.03) association between lesion sum scores and CCasp3 IHC scores.

Although CCasp3 staining was significantly (P = 0.023) higher in clinically-unaffected pigs inoculated with PCV2a compared to clinically unaffected pigs inoculated with PCV2b and sham-inoculated pigs, a difference in CCasp3 staining was not observed between clinically- affected pigs inoculated with PCV2a or PCV2b (Table 1). However, there was a significant

(P < 0.001) association of CCasp3 IHC scores and PCV2 IHC scores (Table 3). Comparison of apoptotic rates as determined by CCasp3 IHC at different DPIs demonstrated that there was no age dependent difference in the severity of apoptosis (data not shown).

4. Discussion

PCV2 infection in gnotobiotic pigs has been found to result in liver failure associated with icterus which becomes the imminent cause of death in pigs with this manifestation of

PCVAD (Krakowka et al., 2004). The current study used experimentally PCV2-inoculated gnotobiotic pigs with and without signs of clinical disease and hepatic failure. The severity of microscopic lesions in hepatic tissue was significantly (P < 0.05) higher in the clinically- affected pigs when compared to the clinically-unaffected and sham-inoculated control pigs

(Fig 1) which is in agreement with previous studies in conventional pigs (Rosell et al.,

2000a), further emphasizing the association of PCV2 with hepatic lesions and clinical disease.

The main goal of this study was to determine if there is an association between

PCV2 and apoptosis of hepatocytes. We found that the numbers of apoptotic cells increased with increased lesion severity and increased amount of PCV2 antigen. The pigs used were essentially germ-free gnotobiotic pigs and the presence of other pathogens or factors can be ruled out. This is in contrast to field studies, where association of specific microscopic changes such as apoptosis with a specific pathogen is difficult as pigs in the field are commonly infected with more than one pathogen. Therefore, results obtained on field cases need to be viewed with caution. Studies conducted by several research groups have identified apoptosis by TUNEL assay (Shibahara et al., 2000; Krakowka et al., 2004;

Mandrioli et al., 2004; Resendes et al., 2004a; Seeliger et al., 2007) and CCasp3 IHC

(Resendes et al., 2004a, b; Seeliger et al., 2007) in conventional, clinically-affected pigs

(Rosell et al., 1999, 2000b). A recent investigation on conventional pigs using the TUNEL assay, CCasp3 IHC and PCV2 in-situ hybridization (ISH) provided evidence of virus- induced apoptosis in endothelial cells in pigs diagnosed with PCV2-associated cerebellar vasculitis (Seelinger et al., 2007). Similarly, a study using conventional clinically-affected pigs, the TUNEL assay and PCV2 ISH found that diminished lymphoid cell proliferation

was the actual cause of cell depletion (Mandrioli et al., 2004). Another group investigating conventional PMWS-affected pigs using CCasp3 IHC determined that apoptosis was lower in clinically-affected pigs when compared to clinically non-affected pigs (Resendes et al.,

2004b). More recently, a study looking at apoptosis in pigs with hepatitis caused by naturally induced PCV2-associated PMWS concluded that there were higher levels of apoptosis in hepatocytes in pigs with higher PCV2 viral loads (Resendes et al., 2010).

While no information is currently available on PCV2-associated apoptosis in conventional specific pathogen free (SPF) pigs, colostrum-deprived (CD) pigs, or caesarian- derived colostrum-deprived (CDCD), investigations to associate PCV2 with apoptosis have been conducted in gnotobiotic pigs. Little to no apoptosis was found in archived paraffin- embedded blocks of formalin-fixed and cold ethanol-fixed liver, lymph nodes, spleen and thymus obtained from clinically-affected and unaffected gnotobiotic pigs (Krakowka et al.,

2004). This is in contrast to the outcomes obtained in the current study and may be due to the fact that lower numbers of pigs were used in the previous study (Krakowka et al., 2004) as well as different challenge virus and different tissues were evaluated. In addition, virus dependent factors such as PCV2 isolates can be closely related and yet be exclusive, differences in detection systems for apoptosis and PCV2 may influence results, and host- dependent factors such as host susceptibility and immune response have been shown to be important (Opriessnig et al., 2006, 2007, 2009).

Differences in pathogenicity between PCV2 genotypes have been suggested; however, definitive experimental proof is lacking to date. While differences in virulence between PCV2 isolates exist (Opriessnig et al., 2006), this appears not to be confined to genotype (Opriessnig et al., 2008). To our knowledge, this is the first study comparing

apoptosis in pigs infected with PCV2a or PCV2b. It has been shown through our study that

PCV2 is involved in apoptosis; however, there was no significant difference found between the numbers of apoptotic cells within PCV2a or PCV2b inoculated pigs. PCV2a and PCV2b differ in a small region in ORF2 (Cheung et al., 2007) encoding the capsid protein

(Nawagitgul et al., 2000), whereas ORF3 which has been demonstrated to be associated with apoptosis (Liu et al., 2005, 2006, 2007), is located within ORF1 and is more conserved among PCV2 isolates, which may explain the results of this study.

A recent study compared the differences between the TUNEL and the CCasp3 IHC assays on lymphoid tissues of healthy conventional pigs of different ages (Resendes et al.,

2004a); however, to the authors’ knowledge no experimental studies have been performed to compare these assays in hepatic tissues. Our study concluded that both methods detected significantly (P < 0.05) higher numbers of apoptotic cells in clinically-affected pigs compared to clinically-unaffected pigs and sham-inoculated control pigs. Nevertheless, when comparing the TUNEL assay directly with the CCasp3 assay we found no differences in detection rates of apoptotic cells (Fig 1) indicating that the same type of cell was labeled by both techniques as described by others using a double staining technique (Mirkes et al.,

2001). However, in order to further confirm this observation, staining of serial sections would be necessary.

5. Conclusion

In summary, higher amounts of PCV2 antigen were demonstrated to be associated with more severe microscopic lesions and increased apoptosis in the hepatic tissues from

PCV2a or PCV2b singularly-infected, clinically-affected gnotobiotic pigs. No differences

were observed between PCV2a- or PCV2b-infected pigs. The TUNEL assay and CCasp-3

IHC appear to be equivalent in their ability to detect apoptotic cells in liver tissues from

PCV2-infected gnotobiotic pigs.

Acknowledgements

We thank Dr. HuiGang Shen for critical review of the manuscript.

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Table 1. Median group scores (minimum, maximum) for amount of PCV2 antigen as determined by PCV2 IHC stains and numbers of apoptotic cells as determined by cleaved caspase-3 (CCasp3) IHC and the TUNEL assay in sham-inoculated control pigs and unaffected and clinically affected pigs inoculated with PCV2a or PCV2b. Clinical Status Inoculation PCV2 IHC* CCasp3** TUNEL** Microscopic lesions*** Non-Affected Sham (n=4) 0 (0, 0)A*** 0 (0, 0)A 0 (0, 0)A 0 (0, 0)A PCV2a (n=10) 0 (0, 2)A 1 (0, 2)B 0.5 (0, 2)A 3 (0, 4)B PCV2b (n=13) 0 (0, 1)A 0 (0, 1)A 0 (0, 1)A 1 (0, 4)B Affected PCV2a (n=6) 3 (0, 3)B 2.5 (1, 3)C 2 (0, 3)B 4 (3, 8)C

PCV2b (n=15) 2 (0, 3)B 2 (1, 3)C 2 (0, 3)B 3 (1, 9)C A, B,C Different superscripts indicate significant (P < 0.05) differences between group medians within the same column. *0 = negative, 1= low number, 2 = moderate numbers and 3 = large numbers. **0 = average of 0.0-0.9 apoptotic cells/10 fields, 1 = average of 1-4.9 apoptotic cells/10 fields; 2 = average of 5-14.9 apoptotic cells/10 fields; 3 = average of 15 or more apoptotic cells/10 fields. ***The lesions sum score for microscopic lesions ranged from 0 = no lesions to 12 = severe lesions

Table 2. Prevalence of scores ranging from 0 to 3 for PCV2 antigen as determined by PCV2 IHC and apoptosis as determined by cleaved caspase-3 (CCasp3) IHC and the TUNEL assay on liver tissues obtained from sham-inoculated controls or pigs inoculated with PCV2a or PCV2b. Score PCV2 IHC* CCasp3** TUNEL** range Controls PCV2a PCV2b Controls PCV2a PCV2b Controls PCV2a PCV2b 0 4/4 9/16 10/28 4/4 1/16 7/28 4/4 6/16 12/28 1 0/4 1/16 5/28 0/4 9/16 10/28 0/4 5/16 6/28 2 0/4 2/16 6/28 0/4 3/16 6/28 0/4 3/16 7/28 3 0/4 4/16 7/28 0/4 3/16 5/28 0/4 2/16 3/28 * 0 = negative, 1 = low number, 2 = moderate numbers, 3 = large numbers. **0 = average of 0.0-0.9 apoptotic cells/10 fields, 1 = average of 1-4.9 apoptotic cells/10 fields; 2 = average of 5-14.9 apoptotic cells/10 fields; 3 = average of 15 or more apoptotic cells/10 fields.

Table 3. Mean CCasp-3, TUNEL and microscopic lesion scores (±SE) at different levels of PCV2 antigen as determined by PCV2 IHC. PCV2 IHC scores* CCasp3** TUNEL** Microscopic lesions*** 0 (n=23) 0.6±0.1 0.3±0.1 1.7±0.4 1 (n=6) 1.0±0.3 0.5±0.3 1.8±0.5 2 (n=8) 1.6±0.2 1.4±0.4 2.4±0.4 3 (n=11) 2.6±0.2 2.3±0.2 5.3±0.7 *0 = negative, 1 = low number, 2 = moderate numbers, 3 = large numbers **0 = average of 0.0-0.9 apoptotic cells/10 fields, 1 = average of 1-4.9 apoptotic cells/10 fields; 2 = average of 5-14.9 apoptotic cells/10 fields; 3 = average of 15 or more apoptotic cells/10 fields. ***The lesions sum score for microscopic lesions ranged from 0 = no lesions to 12 = severe lesions.

Fig 1. Histological comparison of liver sections obtained from a negative control pig (1A-D) and a pig infected with PCV2b and severely affected by porcine circovirus associated disease (PCVAD) (2A-D) (Bars 10 µm). (1A) Hematoxylin & eosin (HE) stain: Normal hepatic architecture; (1B) PCV2 immunohistochemistry (IHC): No visible staining; (1C) Cleaved caspase-3 (CCasp-3) IHC: No visible staining; (1D) TUNEL staining: No visible staining. (2A) HE stain: Diffuse moderate multifocal degeneration of hepatocytes; (2B) PCV2 IHC: Diffuse moderate staining in nuclei and cytoplasm of hepatocyte-like cells; (2C) CCasp-3 IHC: Diffuse moderate to abundant staining of hepatocyte nuclei; (2D) TUNEL staining: Diffuse moderate to abundant staining of hepatocyte nuclei.

CHAPTER 3 - SUPPLEMENTAL DATA: APOPTOSIS IS NOT A FEATURE IN

LYMPHOID TISSUES FROM PIGS CO-INFECTED WITH PRRSV AND PCV2A

OR PCV2B

Abstract

The objective of this study was to determine the effect of porcine reproductive and respiratory syndrome virus (PRRSV) infection and porcine circovirus type 2 (PCV2) subtypes a (PCV2a) or b (PCV2b) on apoptosis rates in lymphoid tissues obtained from co- infected pigs. Forty-four, 3-week-old, conventional PCV2 and PRRSV naïve pigs were randomly divided into one of three groups: Negative controls (n=8), pigs co-infected with

PCV2a and PRRSV (n=18) and pigs coinfected with PCV2b-PRRSV (n=18). Formalin- fixed, paraffin-embedded sections of lymph nodes, tonsils, and spleen from all pigs were analyzed for presence of apoptosis by detecting intra-nuclear cleaved caspase 3 (CCasp3) with immunohistochemistry (IHC). In addition, PCV2 IHC was done and microscopic lesions and amount of PCV2 antigen were compared among the groups. While negative pigs remained negative for PCV2, 28/36 PCV2 infected pigs had low amounts of PCV2 antigen in lymphoid tissues. CCasp-3 staining patterns were observed in all groups, regardless of infectious status and including negative control pigs, indicating that lymphocyte depletion via apoptosis is not a mechanism utilized by PCV2-PRRSV co-infection in subclinically infected pigs.

Introduction

Porcine circovirus type 2 (PCV2) is recognized as an important disease causing agent worldwide (Chae, 2004) and, if uncontrolled, is a major cause of economic loss to the swine industry. PCV2 is a single stranded, non-enveloped icosahedral virus with a genome length of 1767-1768 nucleotides (Hamel et al., 1998; Meehan et al., 1998). PCV2 was first associated with post-weaning multisystemic wasting syndrome (PMWS) in swine herds in western Canada (Ellis et al., 1998). Today, porcine circovirus associated disease (PCVAD) includes systemic infection, respiratory disease, enteric disease, reproductive failure, and porcine dermatitis and nephropathy syndrome (Harding et al., 1998; Allan et al., 2000a;

Rosell et al., 2000b; O'Connor et al., 2001; Kim et al., 2003; Jensen et al., 2006). PCV2 can be further subdivided into PCV2a and PCV2b (Gagnon et al., 2008). The PCV2 genome contains three distinct open reading frames (ORF), where ORF1 and ORF2 encode for the replication and capsid proteins, respectively (Mankertz et al., 1998; Nawagitgul et al., 2000).

ORF3 has been recently associated with a protein that induced apoptosis in host cells (Liu et al., 2006).

Apoptosis is a programmed cell death mechanism used by mammalian cells to maintain homeostasis in the cellular development, and thus have a balance between cell proliferation and death. Apoptosis occurs via two pathways: the extrinsic pathway involving cell death receptor and ligand, and the intrinsic pathway that includes the mitochondrial pathway (Suliman et al., 2001; Wajant, 2002). Both pathways eventually lead to cleavage of

Caspase-3 which induces subsequent poly-ADP ribose polymerase (PARP) cleavage that inhibits DNA repair and facilitates DNA fragmentation (Thornberry, 1998). Many viruses hijack the host programmed cell death pathway to serve their own purpose of propagation as

well as dissemination by inhibition or induction of apoptosis (Teodoro et al., 1997). PCV2 and PRRSV both have been associated with apoptosis in host cells (Suarez et al., 1996;

Sirinarumitr et al., 1998; Choi et al., 2002; Labarque et al., 2003; Liu et al., 2005). Different techniques such as in situ hybridization, immunohistochemistry (IHC), and terminal transferase dUTP nick end labeling (TUNEL) have been utilized to confirm apoptosis in

PRRSV infected pigs (Sur et al., 2000). Moreover, glycoprotein 5 of PRRSV has been demonstrated to be involved in the apoptosis of host cells via its 118 amino acid residues at the N-terminal end (Fernandez et al., 2002).

The main lesion associated with PCV2 infection is lymphoid depletion (Segalés et al., 2004) which occurs via apoptosis of B-cells (Shibahara et al., 2000). Furthermore, hepatic failure in PCVAD-affected pigs was associated with apoptosis of the hepatocytes

(Rosell et al., 2000a). Recently, an association of PCV2 and apoptosis in hepatocytes was confirmed by using dual PCV2 IHC and TUNEL, or dual PCV2 IHC and CCasp-3 IHC

(Resendes et al., 2010). The present study was conducted to further investigate a possible role of PRRSV and PCV2 in induction of apoptosis in lymphoid tissues.

Materials and methods

Formalin-fixed, paraffin embedded lymphoid tissues, tonsil, and spleen were obtained from 44 conventional pigs which came from a farm known to be free of PCV2 and

PRRSV, and were divided into three groups with eight to 18 pigs in each group. The experimental protocol was approved by the Iowa State University Institutional Animal Care and Use Committee. A portion of the pigs were inoculated with PCV2a and PRRSV (n=18) or PCV2b and PRRSV (n=18) and the remaining pigs were kept as non-infected controls

(n=8) following previously described protocols (Sinha et al., 2010). Twelve days after experimental infection, all pigs were euthanized humanely using an overdose of sodium pentobarbital (Vortech Pharmaceuticals, Dearborne, Michigan, USA) and necropsy was performed. Sections of lymph nodes, tonsil, and spleen were collected and 10% neutral buffered formalin fixed and routinely processed for histological examination. The tissues were dehydrated, paraffin-embedded, sectioned at 4µm and mounted on a glass slide followed by routine hematoxylin/eosin staining.

Microscopic lesions were evaluated by a veterinary pathologist (TO) blinded to the treatment groups. Lymph nodes, tonsils and spleen were evaluated for the presence of lymphoid depletion ranging from 0 (normal) to 3 (severe) and histiocytic inflammation and replacements of follicles ranging from 0 (normal) to 3 (severe) (Opriessnig et al., 2004).

Lymphoid lesions were either not observed (control pigs) or were characterized by mild depletion of follicles and mild granulomatous lymphadenitis in all coinfected groups.

Significant differences in lymphoid lesion scores were not observed among the groups (data not shown).

PCV2-specific antigen was detected by IHC using a rabbit polyclonal antiserum as described previously (Sorden et al., 1999). PCV2 antigen scoring was conducted by a veterinarian pathologist (TO) blinded to the treatment groups. The scoring protocol used in this study was as follows: 0 = no signal or visible stain, 1 = staining in individual inflammatory cells, 2 = staining observed in moderate numbers of inflammatory cells, 3 = abundant staining in inflammatory cells. All control pigs were negative for PCV2 antigen by

IHC. A low amount of PCV2-antigen in lymphoid tissues was detected in 16/18 pigs

inoculated with PCV2a and 12/18 pigs inoculated with PCV2b. The prevalence of PCV2 antigen in lymphoid tissues was independent of PCV2 subtype.

Unlike previous studies where the average trial length ranged from 21 to 32 days

(Allan et al., 2000c; Rovira et al., 2002), this trial was terminated at day12 to evaluate

PRRSV-induced lung lesions which tend to be most severe between 10 and 14 days after experimental infection. At the time of necropsy, all pigs were clinically normal. The outcome of the co-infection on clinical disease in the later stages of infection remains unknown. As expected, and similar to a previous study (Yu et al., 2007), the pathological lesions associated with PCV2 were either not present or they were mild; however, PCV2 antigen was detected in most tissues in co-infected pigs.

For demonstration of CCasp-3 specific staining, the tissue sections were dewaxed, rehydrated in graded alcohol and incubated in 3% hydrogen peroxide for 10 min at 22 ºC, rinsed three times in Tris buffered saline (TBS) treated with 0.05% protease (Sigma Aldrich) for 2 min and washed three times with TBS buffer. A CCasp3 IHC (Cell Signaling

Technology®, Danvers, Massachusetts, USA) was applied at a concentration of 1:100 for 1 h at 22 ºC. Further processing was done by using a streptavidin-biotin detection kit (Dako

North America Inc.). Apoptotic cell counts were performed by scoring 10 random using a

45× lens and a 10× eyepiece, yielding a final magnification of 450× with an American

Optical Spencer microscope (Leeds Precision Instrument, Inc; North Minneapolis,

Minnesota, USA). The apoptotic cell score ranged from 0 to 3 (0=average of 0.0-0.9 apoptotic cells/10 fields, 1=average of 1.0-3.9 apoptotic cells/10 fields; 2=average of 4.0-9.9 apoptotic cells/10 fields; 3=average of 10.0 or more apoptotic cells/10 fields). Statistical analysis of the data was performed using the JMP® software version 8.0.1 (SAS Institute,

Cary, NC). A non-parametric Kruskal-Wallis ANOVA followed by pairwise Wilcoxon tests was used. A p-value of less than 0.05 was set as a statistically significant level throughout this study. The average CCasp-3 scores (lymph node, tonsil, and spleen combined) were

3.88±0.3 for control pigs, 4.33±0.3 for PCV2a-PRRSV infected pigs and 4.56±0.3 for

PCV2b-PRRSV infected pigs. There were no significant differences among groups.

Discussion

PCV2 has been associated with severe disease mainly in situations when there has been some immunostimulation or co-infection (Krakowka et al., 2000; Krakowa et al.,

2008). Numerous experimental studies and field observations have concluded that PCV2 and co-infection increases the severity of PCVAD in affected pigs (Allan et al., 1999;

Kennedy et al., 2000; Pallarés et al., 2002). Experimental studies in piglets have concluded that PRRSV actually increases PCV2 replication which enhances the severity of clinical disease of PCVAD in these pigs (Allan et al., 2000b; Allan et al., 2007). In fact, PRRSV has been found to have the highest correlation with PCVAD and PCV2 in a case control study conducted with field observations (Pogranichniy et al., 2002). All these observations suggest that PRRSV and PCV2 might be interacting in-vivo to increase the severity of PCVAD disease. In the current study we were unable to confirm that increased apoptosis rates are linked with concurrent PCV2 and PRRSV infection which may be related to the duration of the study and the lack of clinically affected pigs which is different to our previous study

(Sinha et al., 2010) where clinically affected and unaffected gnotobiotic pigs were compared.

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CHAPTER 4: EFFECT OF IN VITRO INTERACTION OF PRRSV AND PCV2 ON

VIRUS REPLICATION AND IFN-γ AND IL-10 PRODUCTION IN PORCINE

ALVEOLAR MACROPHAGES

A paper prepared for submission

Avanti Sinhaa, Kathy Lina, Michelle Hemanna, Huigang Shena, Nathan M. Beachb, Carmen

Ledesmab, Xiang-Jin Mengb, Chong Wanga,c, Patrick G. Halbura and Tanja Opriessniga,*

aVeterinary Diagnostic and Production Animal Medicine, College of Veterinary Medicine, Iowa

State University, Ames, Iowa, USA bDepartment of Biomedical Sciences and Pathobiology, Center for Molecular Medicine and

Infectious Diseases, College of Veterinary Medicine, Virginia Polytechnic Institute and

State University, Blacksburg, Virginia, USA cDepartment of Statistics, College of Liberal Arts & Sciences, Iowa State University, Ames,

Iowa, USA

Running Title: In vitro interaction of PRRSV and PCV2

*Corresponding author: 1600 S. 16th Street, College of Veterinary Medicine, Iowa State

University, Ames, IA 50011-1250. Phone: +1 515-294-1137. Fax: +1 515-294-3564. E- mail: [email protected].

Abstract

Porcine circovirus type 2 (PCV2), genotype b, is now the predominant subtype of PCV2 worldwide. Severe disease is commonly reported in field cases where PCV2b is found in combination with porcine reproductive and respiratory syndrome virus (PRRSV). The objectives of this study were to advance the understanding of PCV2 and PRRSV interactions by: 1) establishing a PCV2 and PRRSV in-vitro co-infection model, and 2) further investigating the influence of PCV2 ORF2 and the signature motif in ORF2 on PCV2 replication kinetics and cytokine expression. To achieve these objectives, we constructed chimeric viruses composed of different combinations of ORF1 and ORF2 of PCV2a or

PCV2b (chimera PCV2aORF1-2bORF2 and chimera PCV2bORF1-2aORF2). Pulmonary alveolar macrophages (PAMs) were infected singularly or with combinations of PCV2b,

PCV2a, chimeric PCV2aORF1-2bORF2, and chimera PCV2bORF1-2aORF2, and one of five PRRSV isolates: type 1 (SD 01-08) or type 2 (NC-16845, VR-2332, MN-184, JA-142).

Supernatant harvested at 24, 48, 72 and 96 hours post inoculation (hpi) was evaluated by real-time PCR for the presence and amount of PRRSV RNA, PCV2 DNA, and PCV2 mRNA. The levels of IFN-γ and IL-10 were also determined by quantitative ELISAs. PCV2 replication was observed in all groups at 24 hpi; however, at 48 hpi it was limited to groups inoculated with PCV2 strains containing ORF1 of PCV2a (PCV2a, chimera PCV2aORF1-

2bORF2) concurrently infected with PRRSV. Compared to the non-infected controls, IFN-γ was found to be down-regulated 50-100% in all groups except for PAMs co-inoculated with type 1 PRRSV and PCV2 (regardless of strain), and PAMs infected with JA-142 and chimera PCV2bORF1-2aORF2. In summary, in vitro differences in PCV2 or PRRSV replication or IFN-γ and IL-10 production in PAMs inoculated with one of 30 PCV2-

PRRSV combinations were independent of PCV2 ORF2 origin. PCV2a-ORF1 dependent in- vitro enhancement of PCV2 replication was observed when the PCV2a strain in combination with PRRSV (regardless of strain) was utilized further emphasizing the enhancing effect of

PRRSV on PCV2 infection.

Keywords: alveolar macrophages; porcine circovirus type 2 (PCV2); porcine reproductive and respiratory syndrome virus (PRRSV); interaction.

1. Introduction

Porcine circovirus type 2 (PCV2) is a non-enveloped, single stranded, covalently closed circular DNA virus in the genus Circovirus of the family Circoviridae, which was first identified in pigs suffering from postweaning multisystemic wasting syndrome

(PMWS) in Canada in 1991 (Allan et al., 1998; Harding and Clark, 1997; Todd et al., 2005).

PCV2 has become a globally important viral disease of pigs now commonly referred to as porcine circovirus associated disease (PCVAD) (Gillespie et al., 2009). Clinical manifestations of PCVAD include poor body condition, generalized lymphadenopathy, respiratory disease, porcine dermatitis and nephropathy syndrome, enteritis in grow-finish pigs, and reproductive failure (Harms et al., 2002; Kim et al., 2003; Rosell et al., 2000; West et al., 1999).

PCV2 is further divided into three main subtypes, PCV2a, PCV2b, and PCV2c

(Olvera et al., 2006; Segalés et al., 2008; Wang et al., 2009). No differences in virulence between PCV2a and PCV2b isolates were observed in pigs singularly infected with PCV2

(Gillespie et al., 2009; Opriessnig et al., 2008b). Genetically, PCV2a and PCV2b differ in a signature motif located between nucleotides 1472 and 1486 of ORF2, which correlates to a six amino acid difference in the capsid protein (Cheung et al., 2007; Cheung and Greenlee,

2011). Inactivated chimeric DNA clones of PCV2a and PCV2b isolates (i.e. chimera

PCV2aORF1-2bORF2 and chimera PCV2bORF1-2aORF2 ) have been constructed, using the signature motif of one genotype and the remainder ORF2 of the other genotype, to determine the contribution of the signature motif to virulence (Allemandou et al., 2011).

Under in-vitro and in-vivo conditions, clones with swapped signature motifs had decreased virulence when compared to the parental strains PCV2a and PCV2b, respectively

(Allemandou et al., 2011).

PCVAD is multi-factorial disease and the presence of co-factors can enhance PCV2- associated lesions (Ellis et al., 1999; Ellis et al., 2004; Pallarés et al., 2002). Infectious co- factors that have been experimentally documented to enhance PCV2 replication include porcine parvovirus (PPV) (Allan et al., 1999; Kennedy et al., 2000; Krakowka et al., 2000),

Mycoplasma hyopneumoniae (Opriessnig et al., 2004), porcine reproductive and respiratory syndrome virus (PRRSV) (Allan et al., 2000; Harms et al., 2001; Rovira et al., 2002), and porcine torque teno virus (TTV) (Ellis et al., 2008).

PRRSV, a single stranded, enveloped RNA virus of the genus Arterivirus and the family Arteriviridae, was first reported in 1987 and can be divided into two major genotypes: type 1 (European strains) and type 2 (North American strains) (Rossow, 1998).

In breeding age animals, PRRSV infection may manifest as increased numbers of late term abortions, mummified fetuses, stillborn piglets, and live-born weak piglets (Christianson et al., 1993; Mengeling et al., 1994). In grow-finish pigs, PRRSV is associated with pyrexia,

anorexia, severe respiratory disease, and varying levels of mortality (Rossow et al., 1994).

Under field conditions, pigs coinfected with PRRSV and PCV2 were more likely to develop severe disease (Pallarés et al., 2002; Pogranichniy et al., 2002). Experimental studies have shown that the effect of PRRSV on the augmentation of PCV2-associated lesions is due to an enhanced replication of PCV2 in coinfected pigs (Harms et al., 2001; Opriessnig et al.,

2008a; Rovira et al., 2002). The ability of PRRSV to replicate in porcine alveolar macrophages (PAMs) (Bautista et al., 1993; Mengeling et al., 1996), and the inability of macrophages and dendritic cells to degrade PCV2 (Gilpin et al., 2003; Vincent et al., 2003) might lead to macrophage alterations such as an uncontrolled release of cytokines and formation of oxygen radicals, nitric oxides, and proteases which have been previously reported with PCV2 infection (Chang et al., 2006a) and could alter the immune response.

Studies using live pigs are often limited due to space and resource constraints. One of the main advantages of in-vitro comparison studies is the ability to include several PCV2-

PRRSV combinations. The objectives of this study were to: 1) establish a PCV2-PRRSV in- vitro co-infection model and 2) further investigate the significance of PCV2 ORF2 and the

PCV2 signature motif in PCV2/PRRSV interaction by utilizing chimeric viruses between

PCV2a and PCV2b that contain the ORF2 of the reciprocal virus.

2. Materials and Method

2.1. PCV2 inocula

PCV2a. PCV2a isolate 40895 (designated in this study as PCV2a) was isolated from a pig diagnosed with PCVAD on a farm in Iowa in 1998 (Fenaux et al., 2000; Opriessnig et al., 2006) and used to construct an infectious DNA clone (Fenaux et al., 2002). The clone

was used to transfect PK-15 cells and was propagated to a titer of 104.5 50% tissue culture infectious doses (TCID50) per ml.

PCV2b. PCV2b isolate NC-16845 (designated in this study as PCV2b) was isolated from a farm in North Carolina with a history of severe PCVAD in 2006 (Opriessnig et al.,

2008b). An infectious clone was constructed as described (Beach et al., 2010b; Beach et al.,

2010a) and used to transfect PK-15 cells for propagation of the inocula (Opriessnig et al.,

4.0 2008b). The final virus stock used for this study had an infectious titer of 10 TCID50 per ml.

Construction of chimera PCV2aORF1-2bORF2 and chimera PCV2bORF1-2aORF2 infectious clones. Both chimeric PCV2 clones were propagated to an infectious dose of 104.5

TCID50. Chimera PCV2aORF1-2bORF2 and chimera PCV2bORF1-2aORF2 infectious clones were assembled by overlap-extension PCR as previously described (Beach et al.,

2010a). Full-length chimeric genomes were assembled from three overlapping PCR fragments to assemble chimera PCV2aORF1-2bORF2 (primers 1 and 2, primers 3 and 4, and the PCV2a template; primers 5 and 6 and the PCV2b template; Table 1) and chimera

PCV2bORF1-2aORF2 (primers 7 and 2, primers 8 and 4, and the PCV2a template; primers

9 and 10, and the PCV2b template; Table 1). Each amplicon was generated using Platinum®

PCR Supermix High Fidelity mastermix (Invitrogen) with the same amplification parameters (95°C 3 min; 35 cycles of 95°C 30 sec, 54°C 30 sec, 68°C 1.5 min). A fusion

PCR (35 cycles of 95°C 30 sec, 60°C 30 sec, 68°C 1.5 min) was used to assemble fragments of the chimeric DNA clones (primers 2 and 4; Table 1). The chimeric fusion products were digested with SacII, and cloned into pBluescript II SK(+). Each chimeric DNA clone was completely sequenced to confirm that no undesired changes had been introduced. Infectious

virus stocks of PCV2a, PCV2b, and chimeric viruses PCV2aORF1-2bORF2 and

PCV2bORF1-2aORF2 were generated by transfection of PK-15 cells with dimerized or concatomerized infectious DNA clones as previously described (Fenaux et al., 2002; Fenaux et al., 2003).

All four viral infectious clones as outlined in Fig 1. were found to produce infectious virus after transfection into PK-15 cells, and each infectious virus stock was titrated by immunofluorescent assay (IFA) as described previously (Fenaux et al., 2003).

2.2. PRRSV inocula

The type 1 PRRSV isolate used in this study was the SD 01-08 (kindly provided by

Dr. Y. Fang), which has been well characterized (Fang et al., 2004; Fang et al., 2006; Fang et al., 2007; Ropp et al., 2004). In addition, several other type 2 PRRSV isolates were also used in this study including VR-2332, JA-142, MN-184 (kindly provided by Dr. M.

Murtaugh), and NC16845 which was isolated from pigs in North Carolina in 2006. All isolates were propagated on MARC-145 cells in culture media (RPMI-1640) (SIGMA®,

Sigma Aldrich), supplemented with 10% fetal bovine serum (FBS) (Advantage ®, Atlanta

Biologicals, Lawrenceville, GA, USA) and 1% 100× antibiotic-antimycotic (AB/AM)

(Invitrogen, Carlsbad, CA, USA). The same medium was also used for inoculation of

3.8 4.0 PAMs. The isolate titers were calculated to be 10 TCID50 per ml for SD 01-08; 10

6.8 TCID50 per ml for VR-2332, JA-142 and NC16845; 10 TCID50 per ml for MN-184. All isolates were adjusted to 0.01 multiplicity of infection (MOI) for inoculation of the PAMs.

The PRRSV isolates used in this study had an overall nucleotide sequence identity of 61-

91% and an overall amino acid sequence identity of 55-92% in ORF5.

2.3. Isolation of pulmonary alveolar macrophages (PAMs)

PAMs were collected from lung bronchoalveolar lavage (BAL) fluid as previously described (Mengeling et al., 1995) with the following slight modifications. Two healthy pigs serologically free of PCV2, PPV, PRRSV, swine influenza virus, and M. hyopneumoniae were humanely euthanized by intravenous pentobarbital sodium overdose (Fatal Plus ®,

Vortech Pharmaceuticals, LTD, Dearborn, MI, USA), the trachea was aseptically severed, and lungs were removed from the thoracic cavity. The BAL fluid consisted of 100 ml of 1×

Hanks’ Balanced Salt Solution (HBSS) (Cellgro®, Mediatech Inc., Manassas, VA, USA) supplemented with 100× AB/AM (Invitrogen) and was dispensed into the lung with the aid of 50 ml pipette. The trachea was clamped shut and the lung was massaged gently for approximately 4 min. The BAL fluid containing the PAMs was transferred to 50 ml sterile polypropylene conical tubes (Fisherbrand®, Fisher Scientific Inc., Pittsburgh, PA, USA) and the process was repeated three times. The recovered BAL fluid from each pig was pooled and centrifuged at 600 x g for 10 min at 4 C. The supernatant was removed and 8 ml of

HBSS (Mediatech Inc.) was used to re-suspend the PAMs. Three ml of HISTOPAQUE®-

1077 (Sigma Aldrich, St. Louis MO, USA) was added into a 15 ml plug seal cap polypropylene conical tube (Corning Incorporated, Corning, NY 14831) and the PAMs were overlaid slowly while holding the tube at a 45 degree angle. The mixture was centrifuged at

400 x g for 30 min at room temperature. The upper layer was aspirated to approximately 0.5 cm of the opaque surface interface using a Pasteur pipette and discarded. The opaque interface, containing the PAM layer, was transferred to a sterile 50 ml conical tube (Fisher

Scientific Inc.). Ten ml HBSS (Mediatech Inc) was added to the PAM layer, mixed by

inversion, and centrifuged at 600 x g for 10 min. The supernatant was aspirated and discarded, and the PAM pellets were re-suspended in 10 ml of RPMI-1640 (SIGMA®,

Sigma Aldrich).

2.4. Preparation and inoculation of PAMs

The PAMs were counted with a hemocytometer and assessed for viability using trypan blue stain. The PAMs from both animals were pooled and cells were adjusted to final concentration of 2×107 cells per ml in RPMI-1640 (Sigma Aldrich) with 1%AB/AM

(Invitrogen). Immediately post evaluation, 20 plates with 96 wells each, accounting for six inoculation combinations per plate, one set of plates for each time point, three wells for each inoculation combination and time point, were seeded with approximately 5×104 cells per well in culture media and incubated at 37ºC in 5% CO2 for 3 h. Afterwards, the plates were washed once with HBSS (Mediatech Inc.) with 1% of 100× AB/AM (Invitrogen), and infected singularly or with different combinations of viruses (outlined in the experimental design in Table 2) at a MOI of 0.01 in culture media without FBS. The plates were incubated at 37ºC in 5% CO2 for 1 h, washed three times with HBSS (Mediatech Inc.) with

1% of 100× AB/AM (Invitrogen) and 300 µl of culture medium was added to each well.

2.5. Experimental design

The experimental design is summarized in Table 2. For the inoculation of PAMs, one of four PCV2 isolates (Fig 1) and one of five genetically distinct PRRSV isolates (VR-2332,

MN-184, JA-142, NC-16845, and the SD 01-08) were used, which resulted in a total of 30 possible combinations and groups. Each of the described groups was done in three replicates

in three different wells. Quantities of 200 µl of culture supernatants were collected at 24, 48,

72 and 96 hours post inoculation (hpi) and used immediately for total nucleic acid extraction and subsequent real-time PCR analysis or ELISA. At each time point, the supernatant of the three replicates for each group was combined, for a total volume of 600 µl. A portion of the pool was used for testing and the remaining supernatant was immediately frozen at -80ºC for long term storage.

2.6. Total nucleic acid extraction

Total nucleic acids were extracted using the MagMaxTM Viral RNA Isolation Kit

(Applied Biosystems, Life Technologies, Carlsbad, CA, USA) on the KingFisher Flex

System (ThermoFisher Scientific, Pittsburgh, PA, USA).

2.7. Quantitative PCR assays

PCV2 DNA. Detection and quantification of PCV2 DNA was conducted by a quantitative real-time PCR as previously described (Opriessnig et al., 2003).

PCV2a and PCV2b differentiation. Differentiation of PCV2a and PCV2b DNA was done by a multiplex real-time PCR using a primer probe combination designed to target the highly conserved region of ORF2 that distinguishes between PCV2 subtypes (Opriessnig et al., 2010).

PCV2 mRNA. The sequences of primers and probe for quantification of PCV2 capsid

(Cap) mRNA were described previously (Yu et al., 2007). The probe was labeled with Cal

Fluor® Orange 560 (Biosearch Technologies, Novato, CA, USA). The reaction was performed using the QIAGEN OneStep RT-PCR Kit (Qiagen, Valencia, CA, USA) in a 25

µl reaction volume including 5 µl of 5×OneStep RT-PCR buffer, 1 µl of (0.4 mM) dNTP mix, 1 µl of RT-PCR enzyme mix, 0.5 µl of 1:10 diluted ROX™ reference dye (Invitrogen),

1 µl (0.4 µM) of each primer, 0.5 µl (0.2 µM) of probe, 10 µl of DNase/RNase-free water and 5 µl extracted sample. The reactions were completed in a 7500 Fast Real-Time PCR system (ABI, Foster City, CA) under the following conditions: 50°C for 30 min, 95°C for 15 min, followed by 40 cycles of 94°C for 15 s, 55°C for 30 s and 60°C for 1 min. Five progressive 1:10 dilutions of RNA standards (Yu et al., 2007) were used to generate the standard curve.

PRRSV RNA. Real time RT-PCR was conducted on the extracted RNA samples to detect and quantify North American and European PRRSV. For North American PRRSV strains, a forward primer NA15096F (5'- TAGTGAGCGGCAATTGTGTCTG-3'), a reverse primer NA15229R (5'- GGCGCACAGTATGATGCGT-3') and a probe NA15173P (5'-

FAM-CAGGGAGGATAAGTTACACTGTGGAGTTTAG -BHQ-3') specific for ORF7 of

North American PRRSV strains were used with the QIAGEN OneStep RT-PCR Kit

(Qiagen). The total reaction volume was 25 µl consisting of 5 µl of 5×OneStep RT-PCR buffer, 1 µl of (0.4 mM) dNTP mix, 1 µl (0.4 µM) of each primer, 0.5 µl (0.2 µM) of probe,

1 µl of RT-PCR enzyme mix, 0.5 µl of 1:10 diluted ROX™ reference dye (Invitrogen), 7 µl of DNase/RNase-free water and 8 µl extracted sample. The reactions were completed in a

7500 Fast Real-Time PCR system (ABI) under the following conditions: 50°C for 30 min,

95°C for 15 min, followed by 40 cycles of 94°C for 15 s, 55°C for 30 s and 60°C for 1 min.

To achieve a standard curve five serial dilutions of NA PRRSV Quant RNA (Applied

Biosystems) were applied.

For the European PRRSV strain (SD 01-08), RNA was quantified using Taqman®

EU PRRSV Reagents (Applied Biosystems). A final volume of 25 µl PCR reaction mixture with 8 µl of PRRSV RNA standard or extracted SD 01-08 RNA was used. The PCR reaction was carried out as previously described (Shen et al., 2010).

2.8. Cytokine assays

IFN-γ ELISA. IFN-γ in the collected supernatant samples was examined using a commercially available sandwich ELISA kit (Swine IFNγ™; Invitrogen). The quantitative value of each sample was calculated by creating a standard curve using the standards available in the ELISA kit. The assay was performed according to the manufacturers’ specifications and as previously described (Opriessnig et al., 2008a).

IL-10 ELISA. IL-10 ELISA plates were coated using commercially available antibody pairs (Swine IL-10 Antibody Pair; Invitrogen) following the manufacturers’ directions, and the assay was performed according to the protocol provided by the manufacturer. The quantitative values were calculated and compared to the standard curve that was created by using the standards available in the ELISA kit.

2.9 Statistical Analysis

Responses were analyzed using repeated measures analysis of variance (ANOVA) models. PCV2 strain, PRRSV, and time were the fixed effects, whereas sample was the subject of repeated measurements. The responses PCV2 mRNA, PRRSV RNA, and PCV2

DNA were log10 transformed before analysis. P-values < 0.05 were considered significant.

When an explanatory variable was tested significant, post-hoc Tukey’s t-test was used for

pairwise comparisons of mean response between the categories of the variable. The data analyses for this project were performed using SAS software, Version 9.2.

3. Results

3.1 Detection and quantification of PCV2 DNA

The groups not infected with PCV2 remained negative throughout the study. There was no significant difference in the amount of log10 PCV2 DNA in the supernatants at any time during the study among groups infected with PCV2 alone or coinfected with PRRSV and PCV2 (Table 3).

3.2 Differentiation of PCV2a and PCV2b DNA

All supernatants obtained from PAMs infected with PCV2 were positive by real-time

PCR for the correct subtype of PCV2 (data not shown).

3.3 Detection and quantification of replicating PCV2 mRNA

At 24 hpi, all supernatants obtained from PCV2 infected PAMs were positive for

PCV2 mRNA and all samples not inoculated with PCV2 were negative. There was a significant difference (p<0.05) between the supernatant obtained from PAMs inoculated with PCV2b and the remaining groups at this time (Table 4). At 48 hpi, all samples not inoculated with PCV2 were still negative, and the majority of the PAMs infected with PCV2 also became negative for PCV2 mRNA. In particular, PAMs infected with chimera

PCV2bORF1-2aORF2 and PCV2b were negative but PCV2a and chimera PCV2aORF1-

2bORF2 were positive. There was no statistical difference in PCV2 mRNA between any

groups at this time point. At 72 hpi and at 96 hpi, all samples were negative for PCV2 mRNA (Table 4).

3.4 Detection and quantification of PRRSV RNA

The supernatants from PAMs not infected with PRRSV remained negative for

PRRSV RNA throughout the course of the study. At 24 hpi, all supernatants from PAMs infected with type 1 PRRSV (SD 01-08) had significantly lower (p<0.05) copy numbers of

PRRSV when compared to the type 2 PRRSV strains (except VR-2332 which was grouped with both type 1 and type 2 PRRSV, Table 5). At no other time point did PRRSV copy numbers vary among groups (Table 5).

3.5 Detection of IFN-γ

Since no statistical differences were noted when viral infection was compared over time, the results for each time point were averaged and statistical analysis was run to determine if any statistical differences were noted between the different infectious groups.

The supernatant obtained from the non-infected negative control group had up-regulated levels of IFN-γ when compared to all other PCV2 treatment groups, and when compared to type 2 PRRSV infection (Table 6). However, IFN-γ levels in groups infected with type 1

PRRSV were comparable to control levels regardless of PCV2 infection status. Also, PAMs concurrently infected with PRRSV JA-142 and chimera PCV2bORF1-2aORF2 had significantly higher (p<0.05) levels of IFN-γ when compared to PAMs infected with PRRSV

JA-142 and any of the other PCV2 strains, but not when compared to the negative control group. All other viral combinations had similar IFN-γ results (Table 6).

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3.6 Detection of IL-10

Since no statistical differences in mean group IL-10 pg/ml concentrations were noted when viral infection was compared over time, for the statistical analysis the results for each group over time were averaged. IL-10 was not upregulated in any of the groups investigated

(data not shown).

4. Discussion

PCV2 and PRRSV coinfection is commonly seen under field conditions and commonly associated with severe disease (Pallarés et al., 2002; Pogranichniy et al., 2002).

Concurrent PRRSV and PCV2 infection under in vivo experimental conditions has yielded mixed results. PRRSV induced upregulation of PCV2 replication in pig coinfection models

(Allan et al., 2000; Rovira et al., 2002) and PCV2-PRRSV coinfection induced more severe pneumonia (Harms et al., 2001). However, the mechanism of PRRSV-PCV2 interaction has yet to be clearly defined. In this study, we attempted to advance knowledge in this area with an in vitro co-infection model to investigate and compare viral replication levels and IL-10 and IFN-γ expression in cells singularly infected with PCV2 or co-infected with PRRSV and

PCV2.

PAMs and other cells of the monocyte/macrophage lineage are the major target cells for PRRSV replication (Chiou et al., 2000). Typically, high amounts of PCV2 can be detected in the cytoplasm of PAMs or other cells of the monocyte/histiocyte lineage (Rosell et al., 1999). When propagated under laboratory conditions, PCV2 was found to require the presence of actively dividing cells (Tischer et al., 1987). The exact role of PAMs in PCV2

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replication is still not clear. It has been shown that PCV2 is located in the cytoplasm of this cell type without being degraded (Gilpin et al., 2003). Recently, nuclear expression of PCV2 proteins was observed in PAMs; however, this was dependent on the pig source used, implying differences in susceptibility between pigs (Meerts et al., 2005a). As PAMs are fully differentiated cells, the authors suggested that a high level of DNA polymerase activity in response to damage of their DNA in macrophages may be utilized by PCV2 for its replication cycle (Meerts et al., 2005a). In this regard, replication of PRRSV in macrophages may be sufficient to increase DNA polymerase to levels that support and enhance PCV2 replication in PAMs (Allan et al., 2000; Harms et al., 2001; Rovira et al., 2002). Research based on PAMs experimentally infected with PCV2 found that PCV2 was present in the cytoplasm, but was not found in the nucleus unless the cells were treated with lipopolysaccharide (Chang et al., 2006b); however, recently conflicting results were obtained by another research group using a similar experimental design (Fernandes et al.,

2007). The detection of spliced cap protein mRNA has previously been shown to discriminate between replicating PCV2 and PCV2 virions (Yu et al., 2005; Yu et al., 2007).

In the current study, we found that PCV2 had limited replication capabilities in PAMs with the peak of presence of the spliced cap protein mRNA by 24 hpi, declining by 48 hpi, and completely abolished by the later observation points. Interestingly, only groups coinfected with PRRSV had detectable PCV2mRNA levels by 48 hpi further supporting the idea that

PRRSV is enhancing PCV2 replication.

Concurrent infection of PCV2 and PRRSV in vivo frequently leads to increased levels of PCV2 DNA, but no change in the amount of PRRSV RNA (Rovira et al., 2002).

Differences in the ability of PRRSV to replicate in PAMs varied among genetic lines of pig

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(Vincent et al., 2005). In the current study, macrophages obtained from two pigs from the same source were utilized and mixed prior to in-vitro usage to account for possible genetic differences. Future studies should utilize macrophages from different pig sources.

It is well known that under cell culture conditions, type 1 PRRSV strains replicate slower and to a lower level than type 2 strains (Lee et al., 2010), but type 2 PRRSV strains when compared to each other tend to grow at similar levels (Kim et al., 2008). Similar results were observed in this study. No differences in amount of PRRSV RNA between type

2 PRRSV strains were noted, but the type 1 strain replicated slower than the type 2 strains regardless of co-infection status. Of note, only one type 1 PRRSV isolate was used and future studies using additional type 1 PRRSV isolates are required to further confirm the present findings.

Cytokines are important regulators of the mammalian immune system. In particular,

IFN-γ is a cytokine that has been shown to interfere with viral replication (Schroder et al.,

2004). As such, it is an important cytokine to evaluate in a viral coinfection model.

Interestingly, PCV2 replication has been found to be enhanced in porcine cell lines before and after treatment with IFN-γ (Meerts et al., 2005b). In the current study, the quantity of

IFN-γ present in the supernatant of singular or co-infected cells was compared. We found that in almost all viral combinations (single or co-infected), IFN-γ levels were significantly reduced when compared to non-infected cells. However, the higher IFN-γ levels in some groups were not associated with enhanced PCV2 replication, implying that PCV2 stimulation likely depends on markedly increased IFN-γ levels.

IL-10, another important cytokine, has been found to decrease IFN-γ expression along with other cytokines (Pestka et al., 2004). A question posed in this study was whether

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or not IL-10 would be increased in co-infected cells. In a recent study, a virulent strain of

PRRSV was unable to induce higher expression of IL-10 (Subramaniam et al., 2011). In a previous in-vivo study, it was found that M. hyopneumoniae infection potentiated PCV2 infection by increasing IFN-γ and IL-10 mRNA expression levels (Zhang et al., 2011). In contrast, our present study did not detect expression of IL-10 in any of the groups.

In this study, a chimera PCV2aORF1-2bORF2 virus and its reciprocal version, chimera PCV2bORF1-2aORF2, were utilized and compared to their parental strains, PCV2a and PCV2b. We hypothesized that there are distinct differences in the interaction between

PCV2a and PCV2b strains and certain PRRSV isolates that are directly influenced by ORF2 and its signature motif. Specifically, we thought that concurrent in vitro infection with

PCV2b or chimera PCV2aORF1-2bORF2 and recent PRRSV isolates would result in higher levels of viral replication and increased levels of pro-inflammatory cytokines compared to

PCV2a or chimera PCV2aORF1-2bORF2 and PRRSV co-infection. However, no ORF2- related differences were detected. Interestingly, in groups infected with ORF1 of PCV2a

(PCV2a; chimera PCV2aORF1-2bORF2), there were significantly higher levels of PCV2 mRNA present at 48 hpi compared to groups inoculated with chimeric viruses containing

ORF1 of PCV2b (PCV2b; chimera PCV2bORF1-2aORF2). Moreover, groups infected with chimeric viruses containing ORF1 of PCV2a (PCV2a; chimera PCV2aORF1-2bORF2) had significantly lower levels of IFN-γ in PAMs coinfected with PRRSV stain JA-142 compared to groups inoculated with chimeric viruses containing ORF1 of PCV2b (PCV2b; chimera

PCV2bORF1-2aORF2) indicating a role of ORF1 in viral replication.

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5. Concluding remarks

The results of this study indicate that in vitro differences in PCV2 or PRRSV replication or IFN-γ and IL-10 production in PAMs inoculated with one of 30 PCV2-

PRRSV combinations were independent of ORF2 origin. Differences in in-vitro replication; however, were attributable to ORF1 of PCV2 and concurrent PRRSV infection which should be further investigated.

Acknowledgements

Funding for this study was provided by the Iowa Healthy Livestock Initiative.

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Vincent, I.E., Carrasco, C.P., Herrmann, B., Meehan, B.M., Allan, G.M., Summerfield, A., McCullough, K.C., 2003. Dendritic cells harbor infectious porcine circovirus type 2

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in the absence of apparent cell modulation or replication of the virus. J. Virol. 77, 13288-13300. Wang, F., Guo, X., Ge, X., Wang, Z., Chen, Y., Cha, Z., Yang, H., 2009. Genetic variation analysis of Chinese strains of porcine circovirus type 2. Virus Res. 145, 151-156. West, K.H., Bystrom, J.M., Wojnarowicz, C., Shantz, N., Jacobson, M., Allan, G.M., Haines, D.M., Clark, E.G., Krakowka, S., McNeilly, F., Konoby, C., Martin, K., Ellis, J.A., 1999. Myocarditis and abortion associated with intrauterine infection of sows with porcine circovirus 2. J. Vet. Diagn. Invest. 11, 530-532. Yu, S., Carpenter, S., Opriessnig, T., Halbur, P.G., Thacker, E., 2005. Development of a reverse transcription-PCR assay to detect porcine circovirus type 2 transcription as a measure of replication. J. Viol. Methods 123, 109-112. Yu, S., Vincent, A., Opriessnig, T., Carpenter, S., Kitikoon, P., Halbur, P.G., Thacker, E., 2007. Quantification of PCV2 capsid transcript in peripheral blood mononuclear cells (PBMCs) in vitro. Vet. Microbiol. 123, 34-42.

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Table 1. Oligonucleotide primers used in the construction of chimeric PCV infectious DNA clones. Primer Sequence (5’→3’)

1 CCTCCTTGGATACGTCATAGCTGAAAACGAAAGAAGTG

2 AGCCCGCGGAAATTTCTGACAAACGTTAC

3 CCACTTAACCCTTAAATGAATAATAAAAACCATTA

4 TTTCCGCGGGCTGGCTGAACTTTTGAAAG

5 CTTTCGTTTTCAGCTATGACGTATCCAAGGAGGCG

6 GTTTTTATTATTCATTTAAGGGTTAAGTGGGGGGT

7 CCTCCTTGGATACGTCATATCTGAAAACGAAAGAA

8 CCACTTAAACCCTAATGAATAATAAAAACCATTAC

9 GGTTTTTATTATTCATTAGGGTTTAAGTGGGGGGT

10 CTTTCGTTTTCAGATATGACGTATCCAAGGAGGCG

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Table 2. Experimental design (all groups were done in three replicates).

Group Inoculation PRRSV PCV2 1 PCV2a/a 2 PCV2b/b 3 None PCV2a/b 4 PCV2b/a 5 None 6 PCV2a/a 7 PCV2b/b 8 NC-16845 PCV2a/b 9 PCV2b/a 10 None 11 PCV2a/a 12 PCV2b/b 13 VR-2332 PCV2a/b 14 PCV2b/a 15 None 16 PCV2a/a 17 PCV2b/b 18 MN-184 PCV2a/b 19 PCV2b/a 20 None 21 PCV2a/a 22 PCV2b/b 23 JA-142 PCV2a/b 24 PCV2b/a 25 None 26 PCV2a/a 27 PCV2b/b 28 SD 01-08 PCV2a/b 29 PCV2b/a 30 None

Table 3. Mean group amount of log10 PCV2 DNA ± standard error in supernatants of alveolar macrophages infected with different PCV2 strains (prevalence of PCV2 DNA) at different hours post inoculation (hpi).

There was no interaction with PCV2 and PRRSV strain, thus the data for the different PRRSV groups were combined for each PCV2 strain for a group mean. Different superscripts (a,b) indicate significant (p<0.05) differences in group amount of log10 PCV2 DNA

24 hpi 48 hpi 72 hpi 96 hpi

115 a a a a

No PCV2 0.00±0.0 (0/6) 0.00±0.0 (0/6) 0.00±0.0 (0/6) 0.00±0.0 (0/6)

PCV2a/a 8.08±0.1b (6/6) 7.69±0.1b (6/6) 7.76±0.1b (6/6) 7.92±0.1b (6/6)

PCV2a/b 8.06±0.1b (6/6) 7.72±0.1b (6/6) 7.83±0.1b (6/6) 7.82±0.1b (6/6)

PCV2b/a 8.30±0.2b (6/6) 7.80±0.1b (6/6) 7.95±0.1b (6/6) 7.95±0.1b (6/6) PCV2 strain PCV2

PCV2b/b 7.57±0.1b (6/6) 7.24±0.1b (6/6) 7.28±0.1b (6/6) 7.43±0.1b (6/6)

Table 4. Mean group amount of log10 PCV2 mRNA ± standard error in supernatants of alveolar macrophages infected with different PCV2 strains (prevalence of PCV2 mRNA) at different hours post inoculation (hpi). There was no interaction with PCV2 and PRRSV strain, thus the data for the different

PRRSV groups were combined for each PCV2 strain for a group mean. Different superscripts (a,b,c) indicate significant (p<0.05) differences between groups within a single column.

24 hpi 48 hpi 24 hpi 96 hpi 116

No PCV2 0.00±0.0a (0/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6)

PCV2a/a 4.76±0.1b (6/6) 1.19±0.5a (3/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6)

PCV2a/b 4.80±0.4b (6/6) 0.99±0.4a (3/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6) Strain

PCV2b/a 4.91±0.5b (6/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6) PCV2 PCV2

PCV2b/b 4.14±0.2c (6/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6) 0.00±0.0a (0/6)

Table 5. Mean group amount of log10 PRRSV RNA ± standard error in supernatants of alveolar macrophages infected with different PRRSV strains (prevalence of PRRSV RNA) at different hours post inoculation (hpi). There was no interaction with

PCV2 and PRRSV strain, thus the data for the different PCV2 groups were combined for each PRRSV strain for a group mean. Different superscripts (a,b) indicate significant (p<0.05) differences between groups within a single column.

24 hpi 48 hpi 72 hpi 96 hpi

No PRRSV 0.00±0.0a (0/5) 0.00±0.0a (0/5) 0.00±0.0a (0/5) 0.00±0.0a (0/5)

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Type SD 01-08 6.00±0.5c (5/5) 8.57±0.7b (5/5) 8.14±0.3b (5/5) 7.76±0.3b (5/5)

JA-142 8.38±0.7b (5/5) 8.91±0.3b (5/5) 8.85±0.2b (5/5) 8.17±0.4b (5/5) Strain

b b b b

Type MN-184 8.36±0.4 (5/5) 8.04±0.7 (5/5) 9.13±0.1 (5/5) 8.63±0.1 (5/5) PRRSV PRRSV 2 NC-16845 7.65±0.2b (5/5) 9.13±0.1b (5/5) 8.98±0.2b (5/5) 9.08±0.1b (5/5)

VR2332 7.30±0.1bc (5/5) 8.98±0.2b (5/5) 9.12±0.1b (5/5) 8.73±0.1b (5/5)

Table 6. Mean group IFNG values in pg/ml in supernatants of different infectious combinations. No significant difference was noted between the different times the supernatants were harvested, thus the data from each group over time were all combined for the group mean. Different letter superscripts (a,b) indicate significant (p<0.05) differences between groups within a single column and different number superscripts (1,2,3) indicate significant (p<0.05) differences between groups within a single row. PRRSV Strain

Type 1 Type 2 No PRRSV

SD 01-08 JA-142 MN-184 NC-16845 VR-2332

No PCV2 5.32±5.5a(1) 1.91±3.5a(1,2) -2.95±1.7a(2,3) -4.08±2.7a(2,3) -5.86±1.4a(3) -3.71±3.3a(2,3) 118

PCV2a/a 1.35±4.6a,b(1,3) 5.17±1.7a(1) -6.29±0.9a(2) -1.34±1.5a(1,2) -0.72±3.0a(1,2) -4.04±4.4a(2,3)

PCV2a/b -4.45±2.3b(1) 2.76±3.1a(1) -0.01±3.4a,b(1) -3.01±2.3a(1) -3.89±3.5a(1) -3.62±1.9a(1)

PCV2b/a -5.56±1.6b(1) 3.92±2.7a(2) 4.33±0.7b(2) -4.00±2.6a(1) -4.25±4.2a(1) 2.47±2.3a(2) PCV2 Strain PCV2

PCV2b/b -1.28±0.8a,b(1,2) 5.94±4.6a(2) -1.59±3.7a,b(1) -3.02±1.9a(1) -0.12±3.6a(1,2) -4.62±2.6a(1)

50% decrease compared to negative control

100% decrease compared to negative control

150% decrease compared to negative control

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Fig 1. Open reading frame (ORF) 1 and ORF2 composition of the four PCV2 isolates used in this study.

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CHAPTER 5. PRRSV INFLUENCES INFECTION DYNAMICS OF PCV2

SUBTYPES PCV2A AND PCV2B BY PROLONGING PCV2 VIREMIA AND

SHEDDING

A paper accepted by

Veterinary Microbiology

A. Sinha, H.G. Shen, S. Schalk, N.M. Beach, Y.W. Huang,

X. J. Meng, P.G. Halbur, and T. Opriessnig

ABSTRACT

The objective of this study was to determine the effect of porcine reproductive and respiratory syndrome virus (PRRSV) infection on porcine circovirus type 2 (PCV2) subtypes a (PCV2a) or b (PCV2b) viremia and shedding characteristics in oral, nasal and fecal samples in experimentally infected pigs. Twenty-three, 2- to 6-week-old pigs were randomly divided into five groups: Negative control (n=3), PCV2a-I (n=5), PCV2a-PRRSV-

CoI (n=5), PCV2b-I (n=5), and PCV2b-PRRSV-CoI (n=5). Blood, oral, nasal and fecal swabs were collected in regular intervals from day post inoculation (dpi) 0 until dpi 70 and tested by quantitative real-time PCR for the presence and amount of PCV2 DNA and by

ELISA for the presence of PCV2-specific antibodies. The results indicate that there was a significantly (P < 0.05) higher load of PCV2a and PCV2b DNA in serum, oral swabs, nasal swabs and fecal swabs and a higher prevalence of detectable PCV2 antigen in tissues of pigs concurrently infected with PCV2 and PRRSV compared to pigs singularly infected with

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PCV2 further confirming that PRRSV enhances replication of PCV2. Moreover, PRRSV infection significantly prolonged the presence of PCV2 DNA in serum and increased the amount of PCV2 DNA in oral and nasal secretions and fecal excretions in the later stages of infection between dpi 28 and 70. Shedding patterns were similar between groups infected with PCV2a and PCV2b, indicating that there was no subtype-specific interaction with the

PRRSV isolate used in this study. The results from this study highlight the interaction between PRRSV and PCV2 and the importance of controlling PRRSV infection in order to reduce PCV2 virus loads in pig populations.

1. Introduction

Porcine circovirus type 2 (PCV2) is responsible for considerable economic loss in the swine industry worldwide (Segalés et al., 2005a). Porcine circovirus associated disease

(PCVAD) can manifest as enteric, respiratory, reproductive and systemic disease

(Opriessnig et al., 2007). Lymphoid depletion is the hallmark lesion of PCVAD (Segalés et al., 2004) and is thought to induce immunosuppression or immune-modulation in the host

(Darwich et al., 2003) that often leads to secondary infections with other viral or bacterial pathogens (Pallarés et al., 2002; Kim et al., 2003; Dorr et al., 2007).

PCV2 is a single-stranded, non-enveloped, DNA virus (Tischer et al., 1982;

Crowther et al., 2003) that has been divided into several subtypes of which PCV2a and

PCV2b have been documented worldwide (Segalés et al., 2008). The less prevalent subtypes include PCV2c that has been reported only in Denmark in archived samples (Dupont et al.,

2008) and PCV2d and PCV2e that have been reported in China (Wang et al., 2009).

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Several groups have characterized the shedding patterns of PCV2 from pigs experimentally infected with PCV2 and found that PCV2 is shed in nasal and oral secretions as well as in urine and feces (Magar et al., 2000; Bolin et al., 2001; Shibata et al., 2003;

Segalés et al., 2005b; Caprioli et al., 2006). Horizontal transmission of PCV2 via oral-nasal and oral-fecal routes has been suggested as the main mechanism of infection (Segalés et al.,

2005b; Dupont et al., 2009) which was further confirmed by a recent experimental study

(Patterson et al., 2011b). In a retrospective study involving 146 different swine case submissions to a veterinary diagnostic laboratory in Spain, serum, nasal, tonsillar, tracheobronchial, urinary and fecal swab samples were tested for the presence and quantity of PCV2 (Segalés et al., 2005b). The authors confirmed that shedding of PCV2 occurs via the oronasal, urine and fecal routes and that the PCV2 DNA load was correlated with the clinical status of pigs (Segalés et al., 2005b). Recently, the PCV2 shedding patterns in U.S. pigs naturally (Patterson et al., 2011a) or experimentally (Patterson et al., 2011b) infected with PCV2b were characterized and results indicated that PCV2b was shed for at least 181 days in naturally infected pigs and 69 days in experimentally-infected pigs.

Vertical transmission associated with PCV2 resulting in reproductive failure in field cases has been well documented (O'Connor et al., 2001; Ladekjær-Mikkelsen et al., 2001).

The PCV2 shedding patterns in boar semen have been investigated in both naturally

(McIntosh et al., 2006b) and experimentally (Larochelle et al., 2000; Madson et al., 2008) infected boars. It was found that boars shed PCV2 intermittently (Larochelle et al., 2000) or continuously (Madson et al., 2008). Furthermore, in one experimental study it was concluded that PCV2-spiked semen administered via artificial insemination was able to cause reproductive failure in sows (Madson et al., 2009a). Investigators have also been able

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to prove that sows inoculated with PCV2 during pregnancy shed PCV2 in colostrum and milk during lactation (Ha et al., 2009).

Co-infections increase the severity of the PCVAD caused by PCV2 (Allan et al.,

1999, 2000; Kennedy et al., 2000; Harms et al., 2001; Rovira et al., 2002; Opriessnig et al.,

2004). Porcine reproductive and respiratory syndrome virus (PRRSV) infection has been shown to enhance the severity of clinical PCVAD under field and experimental conditions

(Harms et al., 2001; Rovira et al., 2002; Dorr et al., 2007). Since both PCV2 and PRRSV infect the monocytic cell lineage (Duan et al., 1997; Chang et al., 2006), PRRSV has been suggested to be associated with enhanced PCV2 replication (Allan et al., 2000) resulting in increased amounts of PCV2 DNA present in the sera of PCV2-PRRSV co-infected groups

(Rovira et al., 2002). Moreover, PRRSV was found to increase Interleukin (IL)-10 expression which in turn suppresses T cell response (Charerntantanakul et al., 2006); hence, this may allow PCV2 to circumvent the host immune response.

Co-infection of PCV2 and PRRSV in pigs is frequently observed in swine herds under field conditions; however, to our knowledge, the effect of PRRSV on PCV2 shedding patterns under controlled experimental conditions has not been investigated to date. The objectives of this study were to determine if co-infection of piglets with PRRSV and PCV2 has any affect on PCV2 shedding characteristics and if there are differences between PCV2 subtypes (PCV2a and PCV2b) in co-infected pigs.

2. Materials and methods

2.1. Animals and housing

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Twenty-three, 2- to 6-weeks-old, cross-bred, specific-pathogen-free (SPF) pigs were purchased from a herd known to be free of PCV2 and PRRSV as determined by routine serology. The pigs were housed in a BSL-2 facility at Iowa State University. Pigs were kept in groups of 3-5 animals in five separate 2 × 2.5 m rooms on a concrete floor, equipped with one nipple drinker and a self feeder. The pigs were fed a diet free of animal proteins

(excluding whey) and antibiotics (Nature’s Made, Heartland Co-op, Cambridge, IA, USA).

2.2. Experimental design

The experimental protocol was approved by the Iowa State University Institutional

Animal Care and Use Committee (IACUC Number 10-07-6441-S). The experimental design is summarized in Table 1. After arrival (0 day post inoculation, dpi), 10/23 pigs were inoculated with PCV2a, 10/23 pigs were inoculated with PCV2b, and 3/23 remained non- infected as control pigs. In addition, 5/10 PCV2a and 5/10 PCV2b pigs were also inoculated with PRRSV on the same day as PCV2 inoculation. Blood, oral, fecal and nasal samples were collected at regular intervals and tested for presence of PCV2 DNA and anti-PCV2 antibodies. On each sample collection day, two different teams collected samples from

PCV2a (PCV2a-I, PCV2a-PRRSV-CoI) and PCV2b infected pigs (PCV2b-I, PCV2b-

PRRSV-CoI) to minimize risk of room cross-contamination. Necropsy was performed on all pigs at 70 dpi.

2.3. Clinical evaluation

All pigs were checked daily for respiratory signs, lethargy, inappetence and other signs of disease such as lameness.

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2.4. Inoculation

Inoculation was done intranasally by holding the pig upright and slowly dripping 1.5 ml of the inoculum into each nostril for a total of 3 ml of PCV2a or PCV2b per pig. PRRSV inoculation was done in a similar way. Three separate inoculation teams were used for

PCV2a, PCV2b, and PRRSV inoculation to avoid cross-contamination between rooms.

PRRSV inoculation was done approximately 30 min before PCV2a and 30 min after PCV2b inoculation. Specifically, 3 ml of a PCV2a strain 40895 (Fenaux et al., 2000; Opriessnig et al., 2006) was used as the PCV2a inoculum at a dose of 104.5 50% tissue culture infective dose (TCID50) per ml and administered to each of the 10 pigs in the PCV2a group. Similarly,

3 ml of a PCV2b strain NC-16845 (Opriessnig et al., 2008b) at an infectious dose of 104.5

TCID50 per ml was administered to each of the 10 pigs in the PCV2b group. In addition, 3 ml of a PRRSV strain ATCC VR2385 (Meng et al., 1994; Halbur et al., 1995b) at an

5.0 infectious dose of 10 TCID50 per ml was given to 5/10 PCV2a and 5/10 PCV2b inoculated pigs, respectively.

2.5. Sample collection

Blood, oral, fecal and nasal swabs were collected on dpi 0, 1, 3, 5, 7, 9, 11, 13 and

14. Afterwards, samples were collected in 7-day-intervals until the termination of the study at dpi 70 resulting in a total of 17 sample collection points. Oral, nasal and fecal swabs were collected using individually packaged sterile swabs (Fisherbrand applicators, Fisher

Scientific, Inc.) which were stored in 5 ml polystyrene round bottom tube (Falcon®, BD

Biosciences, San Jose, CA, USA) containing 1 ml of sterile saline solution (Fisher Scientific

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Inc.). Blood was collected in 8.5 ml serum separator tubes (BD vacutainer®, BD

Biosciences), centrifuged at 2,000 x g for 10 min at 4o C and the serum was stored in 5 ml polystyrene round bottom tubes (Falcon®, BD Biosciences) at -80o C until further use.

2.6. PCV2 DNA and PRRSV RNA extraction

PCV2 DNA and PRRSV RNA extraction were performed with an automated extraction machine (King Fisher® Flex 96 Ambion®; Thermo Scientific, Pittsburg, PA,

USA) and the MagMAXTM-96 viral isolation kit (Ambion ®, Austin, TX, USA) according to the instructions of the manufacturer.

2.7. PCV2 DNA detection and quantification

PCV2 DNA detection was conducted to verify the presence of PCV2 in all serum, oral, fecal and nasal samples and DNA quantification was done by a quantitative real-time

PCR using the same primers and probe as previously described (Opriessnig et al., 2003).

Differentiation of PCV2a and PCV2b DNA was achieved by a multiplex real-time PCR using primers and probes designed for the signature motif region of ORF2 capable of distinguishing between PCV2a and PCV2b (Opriessnig et al., 2010). The following modifications were used for both real-time PCR assays: A TaqMan® Fast Universal PCR

Master Mix (Applied Biosystems, Foster City, CA, USA) was used, the total PCR reaction volume including 2.5 μl of DNA extract was 25 μl, and the thermal cycle conditions were 95

°C for 2 min, followed by 40 cycles of 95 °C for 10 s and 60 °C for 1 min. Samples were considered negative if no signal was observed during the 40 amplification cycles. The detection limit of PCV2 quantitative real-time PCR assay was 1×103 copies/ml.

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2.8. PCV2-IgG antibody detection

All serum samples were tested using the SERELISA® PCV2 Ab Mono Blocking kit

(Synbiotics Europe, Lyon, France) according to the manufacturers’ instructions. Results were expressed as sample to negative corrected (SNc) ratio. Samples were considered negative for the presence of anti-PCV2 antibodies if the SNc ratio was greater than 0.50 and they were considered positive if the calculated SNc ratio was less than or equal to 0.50.

2.9. PRRSV RNA detection

Quantitative PRRSV real time RT-PCR was conducted on RNA extracts from all serum samples obtained from PRRSV infected pigs. In brief, a forward primer (NA15096F;

5'- TAGTGAGCGGCAATTGTGTCTG-3'), a reverse primer (NA15229R; 5'-

GGCGCACAGTATGATGCGT-3') and a probe (NA15173P; 5'- FAM-

CAGGGAGGATAAGTTACACTGTGGAGTTTAG -BHQ-3') specific for the ORF7 of type 2 PRRSV strains were designed according to the GenBank accession number

EU880438. The total reaction volume was 25 µl consisting of 5 µl of 5× OneStep RT-PCR buffer (Qiagen, Valencia, CA, USA), 1 µl of (0.4 mM) dNTP mix, 1 µl (0.4 µM) of each primer, 0.5 µl (0.2 µM) of probe, 1 µl of RT-PCR enzyme mix, 0.5 µl of 1:10 diluted

ROX™ reference dye (Invitrogen Life Technologies, Baltimore, Maryland, USA), 7 µl of

DNase/RNase-free water and 8 µl extracted sample. The reactions were completed in a 7500

Fast Real-Time PCR system (Applied Biosystems) under the following conditions: 50 °C for

30 min, 95 °C for 15 min, followed by 40 cycles of 95 °C for 15 s, and 60 °C for 1 min. To produce a standard curve, five serial dilutions of TaqMan® NA PRRSV RNA Control

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(Applied Biosystems) were included in each PCR run. Samples were considered negative if no signal was observed during the 40 amplification cycles.

2.10. PRRSV- IgG antibody detection

Serum samples collected at dpi 0, 35 and 70 were tested for the presence of antibodies to PRRSV (HerdChek* PRRS X3, PRRS virus antibody test kit, IDEXX

Laboratories Inc., Westbrook, Massachusetts, USA). According to the manufacturers’ instructions, a sample-to-positive (S/P) ratio of equal to or greater than 0.4 was considered positive.

2.11. Necropsy

At 70 dpi, the pigs were euthanized humanely using an intravenously administered overdose of pentobarbital (Vortech Pharmaceuticals, Dearborne, MI, USA) and necropsy was performed. Lung sections were scored for macroscopic lesions ranging from 0% to

100% (Halbur et al., 1995b). Lymph node size was scored from 0 (normal) to 3 (enlarged four times its normal size) (Opriessnig et al., 2004). Sections of organs including lymph nodes, spleen, thymus, tonsil, lung, heart, kidney, liver, colon and ileum were collected in

10% neutral buffered formalin and routinely processed for histological examination.

2.12. Histopathology

Microscopic lesions were evaluated by a veterinary pathologist (TO) blinded to the treatment groups. The lung sections were scored for the presence and severity of interstitial pneumonia ranging from 0 (normal) to 6 (severe diffuse) (Halbur et al., 1995b). Lymphoid

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tissues (lymph nodes, tonsils and spleen) were evaluated for the presence of lymphoid depletion ranging from 0 (normal) to 3 (severe) and histiocytic inflammation and replacement of follicles ranging from 0 (normal) to 3 (severe) (Opriessnig et al., 2004).

Sections of formalin fixed lung, heart, kidney, liver, colon and ileum were evaluated for the presence of lymphohistiocytic inflammation.

2.13. Immunohistochemistry

Immunohistochemistry (IHC) for detection of PCV2-specific antigen was performed on formalin-fixed and paraffin-embedded sections of lymph nodes (tracheobronchial, mesenteric, mediastinal, superficial inguinal, and external iliac), tonsil and spleen using a rabbit polyclonal antiserum (Sorden et al., 1999). Scores ranged from 0 (no signal) to 3

(more than 50% of the lymphoid follicles contain cells with PCV2-antigen staining)

(Opriessnig et al., 2004). Immunohistochemistry for detection of PRRSV-specific antigen was performed on lung sections as previously described (Halbur et al., 1995a).

2.14. Statistical analysis

Statistical analysis of the data was performed using the JMP® software version 8.0.1

(SAS Institute, Cary, NC). Summary statistics were calculated for all groups to assess the overall quality of the data including normality. One-way analysis of variance (ANOVA) was used to evaluate the differences among treatment groups. If differences in group means

(positive and negative animals within a group were included) were observed then Tukey-

Kramer test was used for each pair-wise comparison. A P-value of less than 0.05 was set as a statistically significant level throughout this study. Real-time PCR results (copies per ml)

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were log10 transformed prior to statistical analysis; negative samples were treated as 1 copy per ml (log10 = 0). A non-parametric ANOVA (Kruskal-Wallis) was used for non-normally distributed data or when group variances were dissimilar, pair-wise comparisons were done using Wilcoxon rank sum test. A chi-square distribution was conducted to determine differences in prevalence.

3. Results

3.1. Clinical disease

Pigs co-infected with PRRSV and PCV2 developed mild respiratory signs starting at dpi 5 (sneezing and increased respiratory rates) which resolved by dpi 14. One PCV2a-I pig developed severe lameness due to bacterial arthritis and had to be euthanized at 63 dpi.

Wasting was not observed in any of the pigs.

3.2. PCV2-IgG antibody detection

At dpi 0, all pigs were negative for anti-PCV2 antibodies and negative control pigs remained seronegative for PCV2 throughout the study (data not shown). The group mean

SNc ratio for all PCV2 infected pigs is summarized in Fig. 1. In the PCV2-infected pigs, seroconversion was first observed at dpi 9 (PCV2b-PRRSV-CoI) or at dpi 13 (PCV2a-I,

PCV2a-PRRSV-CoI, PCV2b-I). All pigs co-infected with PRRSV had seroconverted to

PCV2 by dpi 21, all PCV2a-I pigs had seroconverted by dpi 35 and all PCV2b-I pigs had seroconverted by dpi 42. Group mean SNc ratios among PCV2-infected groups were significantly (P < 0.05) different on dpi 7 (PCV2a-I pigs had a higher SNc ratio compared to

PCV2a-PRRSV-CoI pigs), dpi 21 (PCV2a-I and PCV2b-I pigs had higher SNc ratios

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compared to PCV2a-PRRSV-CoI and PCV2b-PRRSV-CoI pigs) and dpi 28 (PCV2a-I and

PCV2b-I pigs had higher SNc ratios compared to PCV2a-PRRSV-CoI and PCV2b-PRRSV-

CoI pigs) (Fig. 1).

3.3. PCV2 DNA detection in serum samples

The prevalence of PCV2 DNA-positive pigs in sera is summarized in Table 2 and the mean group amounts of log10 PCV2 DNA in groups infected with PCV2 are summarized in

Fig. 2. All pigs were negative for PCV2 DNA at dpi 0, and all negative control pigs remained negative for PCV2 DNA through termination of the study at dpi 70. All PCV2

PCR-positive pigs were confirmed to be infected only with the PCV2 subtype they were inoculated with as determined by PCV2a/b differential real-time PCR, indicating that cross- contamination between rooms had not occurred (data not shown). PCV2 viremia was first detected on dpi 5 in PCV2a-I and PCV2b-PRRSV-CoI groups and by dpi 11, all pigs in all

PCV2-infected treatment groups were viremic. All pigs co-infected with PRRSV (PCV2a-

PRRSV-CoI, PCV2b-PRRSV-CoI) remained PCV2 viremic until dpi 70, whereas singularly

PCV2-infected pigs had intermittent viremia starting at dpi 35 (Table 2). PCV2b-I pigs had significantly (P < 0.05) less amounts of PCV2 DNA compared to PCV2a-PRRSV-CoI at dpi

11, 13, 14, 21 and 28. From dpi 42 to 70, PCV2a-PRRSV-CoI and PCV2b-PRRSV-CoI had significantly (P < 0.05) higher amounts of PCV2 DNA compared to PCV2a-I and PCV2b-I.

Concurrent PRRSV infection (regardless of PCV2 subtype) significantly increased the amount of PCV2 DNA in serum on dpi 13, 14, 21, 35, 42, 49, 56, 63, and 70. In addition, pigs infected with PCV2a (regardless of PRRSV co-infection status) had significantly (P <

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0.05) higher amounts of PCV2 DNA compared to pigs infected with PCV2b on dpi 11 and

28 (Fig. 2).

3.4. PCV2 shedding in oral, nasal, and fecal secretions and excretions

3.4.1. Oral swabs

PCV2 DNA was not detected in any of the oral swabs obtained from the negative controls (data not shown). PCV2 DNA was detected in oral secretions of PCV2-inoculated groups from dpi 1 onwards (Tables 2 and 3). There was a significant (P < 0.05) effect of

PRRSV status (regardless of PCV2 subtype) on the amount of PCV2 DNA present in oral secretions on dpi 11, 13, 21, 28, 35, 56, 63, and 70 on which days the mean group log10

PCV2 DNA amount was higher in pigs concurrently infected with PCV2 and PRRSV compared to pigs singularly infected with PCV2 (Fig. 3). In addition, PCV2a was shed in significantly (P < 0.05) higher amounts compared to PCV2b on dpi 9, 12, 13, 14 and 49

(data now shown).

3.4.2. Nasal swabs

PCV2 DNA was not detected in any of the nasal swabs obtained from the negative controls (data not shown). PCV2 DNA was detected in the majority of the nasal swabs on dpi 1 (Tables 2 and 3). At dpi 3 through dpi 11, there was a decreased prevalence of PCV2

DNA-positive pigs; however, between dpi 13 and 70 the majority of PCV2-inoculated pigs were positive for PCV2 DNA in nasal swabs without differences in prevalence among treatment groups (Table 2). PRRSV-infected pigs (regardless of PCV2 subtype) had a trend of having higher amounts of PCV2 DNA in nasal swabs compared to singularly PCV2- infected pigs and the difference was significant (P < 0.05) on dpi 13, 14, 21, 35, 42 and 70

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(Fig. 4). In addition, PCV2a (regardless of PRRSV status) was shed in significantly (P <

0.05) higher amounts compared to PCV2b on dpi 9 and 13, whereas on dpi 49 the amount of

PCV2b DNA in nasal swabs was significantly (P < 0.05) higher compared to PCV2a DNA

(data not shown).

3.4.3. Fecal swabs

PCV2 DNA was not detected in any of the fecal swabs obtained from the negative controls (data not shown). The prevalence of PCV2 DNA-positive fecal swabs in PCV2- infected pigs is summarized in Table 2, and was similarly high in all groups during the duration of the study. On dpi 5, 21, 28, 35, 49 and 56, pigs concurrently infected with

PRRSV and PCV2, regardless of PCV2 subtype, had significantly (P < 0.05) higher amounts of PCV2 DNA in feces (Table 3 and Fig. 5). On dpi 9, 13 and 35, pigs infected with PCV2a (PCV2a-I, PCV2a-CoI) had significantly (P < 0.05) higher amounts of PCV2

DNA in fecal samples compared to pigs infected with PCV2b.

3.5. PRRSV- IgG antibody detection

All pigs were negative for anti-PRRSV specific antibodies on dpi 0. All pigs in the

PRRSV-infected groups (PCV2a-PRRSV-CoI, PCV2b-PRRSV-CoI) seroconverted by dpi

35 and remained positive throughout the study without differences in mean group S/P ratios.

All pigs in the other groups remained PRRSV seronegative through dpi 70 (data not shown).

3.6. PRRSV RNA detection in serum

PRRSV RNA was not detected in any serum samples collected on dpi 0. Pigs that were not inoculated with PRRSV (Negative controls, PCV2a-I, PCV2b-I) remained negative

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for PRRSV RNA in serum throughout the study. In the PRRSV-infected groups, PRRSV was detected in 4/5 PCV2a-PRRSV-CoI pigs and in 3/5 PCV2b-PRRSV-CoI pigs on dpi 1.

All 10 PRRSV-infected pigs were PCR positive on dpi 3, 5, 7, 9, 13 and 14. Thereafter,

PRRSV was detected in individual pigs in both groups until dpi 56. All PCV2a-PRRSV-CoI and PCV2b-PRRSV-CoI pigs were PRRSV negative on serum on dpi 63 and 70. There were no significant differences in the amounts of PRRSV RNA between the two PRRSV-infected groups; however, the magnitude of the PRRSV viremia appeared bicyclical in both groups with peaks at dpi 5 and at dpi 21 after which the amount of PRRSV steadily decreased in all pigs (data not shown).

3.7. Macroscopic lesions

At 70 dpi, there were mild lung lesions present in individual PCV2-PRRSV co- infected pigs characterized by 4-17% of the lung surface affected by mottled tan consolidation. In addition, some of the lungs failed to collapse. Lymph nodes appeared normal to mildly enlarged in most infected pigs.

3.8. Microscopic lesions

PCV2-associated lymphoid lesions were essentially resolved in all pigs and lymphoid tissues were microscopically normal. A portion of the pigs in the co-infected groups had mild-to-severe lymphohistsiocytic interstitial nephritis (4/5 PCV2a-PRRSV-CoI pigs and 1/5 PCV2b-PRRSV-CoI pig) and individual pigs in all groups had mild focal interstitial pneumonia characterized by accumulation of low numbers of macrophages and lymphocytes in alveolar septa.

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3.9. Presence of PRRSV and PCV2 antigen in tissues

PRRSV antigen was not detected in any of the lung sections. PCV2 antigen was detected in low to moderate amounts (score range from 1 to 2) in the majority of lymphoid tissues tested of all ten PCV2-PRRSV-CoI pigs regardless of PCV2 subtype whereas PCV2 antigen (low amount in one follicle in the tonsil) was detected in 1/5 PCV2b infected pigs and in 0/5 PCV2a infected pigs.

4. Discussion

Both horizontal (Segalés et al., 2005b; Dupont et al., 2009) and vertical (O'Connor et al., 2001; Ladekjær-Mikkelsen et al., 2001) routes of PCV2 transmission have been documented; however, it is known that PCV2 replication dynamics are highly influenced by certain co-factors (Opriessnig et al., 2007). To the authors’ knowledge, no study to date has investigated the shedding characteristics of PCV2 over time in pigs concurrently infected with PRRSV under experimental conditions. In this study, we also compared the shedding characteristics of the two primary PCV2 subtypes, PCV2a and PCV2b, side by side in the growing pig model.

Several research groups have investigated the length of PCV2 viremia under both controlled experimental and natural settings (Bolin et al., 2001; Shibata et al., 2003; Caprioli et al., 2006; McIntosh et al., 2006a; Opriessnig et al., 2010). The PCV2 shedding pattern of experimentally PCV2-infected cesarean-derived, colostrum-deprived (CDCD) pigs was monitored for 70 days using oropharyngeal, nasal and fecal swabs and it was found that

PCV2 was shed consistently for the 70-day observation period (Shibata et al., 2003). In

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another controlled experiment, pigs inoculated with PCV2a had persistent viremia for 140 days despite the presence of anti-PCV2 antibodies (Opriessnig et al., 2010).

A recent controlled study demonstrated that PCV2b shedding in an experimental setting lasted for 69 days (Patterson et al., 2011b). The same study also demonstrated that, although the amount of PCV2 DNA detected in nasal swabs, oral swabs and fecal swabs was not different at dpi 16 and 19, a higher amount was detected in nasal swabs at dpi 69

(Patterson et al., 2011b). In the present study, similar results were obtained with PCV2 DNA being shed in oral and nasal secretions and feces for an extended period of time with peak shedding between dpi 14 to dpi 35, which was similar for both PCV2 subtypes used in this study. In singular PCV2-infected pigs, low quantities of PCV2a or PCV2b DNA were detectable in sera through dpi 63-70.

In this study, we also compared the shedding patterns of PCV2a and PCV2b side by side in growing pigs, which to our knowledge, has not been reported to date. When PCV2b spread rapidly through North America during 2005 and 2006 (Cheung et al., 2007; Gagnon et al., 2007), it is believed that this may have been due to higher levels of shedding of the

PCV2b subtype which enhanced its ability to transmit from pig to pig and contaminate facilities and transport vehicles. In a previous experimental study, the shedding pattern of these two subtypes with emphasis on semen shedding was investigated in mature boars

(Madson et al., 2008). In that study PCV2a and PCV2b DNA were detected in the serum as early as dpi 2 and in semen between dpi 4 and dpi 5 with detectable semen virus shedding until dpi 60 for PCV2a and dpi 90 for PCV2b; however, no difference was detected between

PCV2 subtypes. This is similar to what we found in the present study using the growing pig

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model, indicating that both subtypes have similar shedding patterns in pigs infected with

PCV2 alone and with concurrent PRRSV infection.

It has been suggested that PRRSV-induced enhancement of PCV2 replication occurs via initiation of the cell cycle in cells co-infected with both viruses as monocytes and macrophages are common targets for both PRRSV and PCV2 (Allan et al., 2000). Recently, it was demonstrated that PCV2 replication can be enhanced by IFNγ in vitro (Meerts et al.,

2005). As a type II interferon, INFγ is critical to inhibit viral replication and is increased in pigs infected with PRRSV. Finally, under in vitro conditions it was demonstrated that increased pro-inflammatory response due to PRRSV and a reduced PRRSV-associated cytophatic effect on alveolar macrophages enhance PCV2 replication (Chang et al., 2005).

In this study, a PRRSV control group was not included as it has been previously shown that

PCV2 does not affect PRRSV replication or lesions (Opriessnig et al., 2008a; Rovira et al.,

2002). The amount of PRRSV was not different between pigs concurrently infected with

PCV2a or PCV2b; however, interpretation of a possible effect of PCV2 on PRRSV replication is impossible in the present study due to the lack of a singular-PRRSV-infected control group. PRRSV viremia was observed in individual pigs up to dpi 56; however, comparison of the PCV2 levels in coinfected pigs that were PRRSV viremic by dpi 56 with their coinfected non-PRRSV viremic group mates at the same dpi revealed no differences and there was no association between prolonged presence of PRRSV infection and presence and amount of PCV2. Although most experiments involving PRRSV are terminated at 3-6 weeks (Halbur et al., 1995b; Halbur et al.,1996; Opriessnig et al., 2008a; Rovira et al., 2002) and only few studies on chronic PRRSV infection are available in the literature, similar

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observations (i.e. the presence of PRRSV viremic pigs by dpi 56) have been reported

(Molina et al., 2008).

Under in vivo conditions, it has been determined that PRRSV increases PCV2 DNA load in the sera of co-infected pigs (Rovira et al., 2002; Opriessnig et al., 2008a) and PCV2 antigen in tissues (Allan et al., 2000), which result in more severe PCV2-associated lesions

(Harms et al., 2002) and a higher likelihood of developing clinical manifest disease. Clinical manifest PCVAD was not observed in any of the pigs which is in contrast to other experiments (Harms et al., 2002). Explanations for this difference may be related to the type of pigs utilized (conventional) or the particular PRRSV strain used (ATCC VR2385).

Similar to previous studies, we found a significantly higher load of PCV2 DNA in the sera of PCV2 and PRRSV co-infected pigs compared to singularly infected PCV2 pigs; however, in contrast to previous studies which were terminated between 17 dpi and 30 dpi (Allan et al., 2000; Rovira et al., 2002; Harms et al., 2002; Opriessnig et al., 2008a), the pigs in the current study were kept until dpi 70. Interestingly, although the PCV2 DNA load in sera was significantly higher in co-infected pigs earlier on (dpi 13, 14, 21), we also observed the maintenance of a high level of PCV2 DNA between dpi 35 to 79 with mean log10 PCV2

DNA quantities late in the infection course essentially similar or even higher when compared to peak viremia measured in singular infected pigs between dpi 11 and dpi 14

(Fig. 2).

Infectivity of the PCV2 positive samples was not determined. Currently available cell culture systems for PCV2 isolation and propagation often are not very efficient

(Opriessnig et al., 2007) and usage of a pig bioassay as previously utilized for determining the viability of PCV2 in boar semen (Madson et al., 2009b) was not an option due to the

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large numbers of samples to be tested. However, at 70 dpi all coinfected pigs still had low- to-moderate amounts of PCV2 antigen in the majority of the lymphoid tissues indicating presence of residual PCV2 whereas only 1/10 singular PCV2-infected pigs had low amounts of PCV2 antigen (staining in one follicle in the tonsil). As it has been shown that a minimum viral load of 108 PCV2 genomes per 500 ng DNA is required for visible IHC staining

(Brungborg et al., 2004) this further highlights the potentiating effect of PRRSV on the chronic stages of PCV2 infection.

Several experimental studies have compared the presence of PCV2 DNA in serum, nasal, oral and fecal excretions and have concluded that PCV2 DNA is present in all the samples in essentially equivalent amounts (Chung et al., 2005; McIntosh et al., 2006a; Grau-

Roma et al., 2009; Patterson et al., 2011b). In the current study, we determined the amounts of PCV2 shed in different secretions and excretions and compared PCV2 subtypes and evaluated the effect of concurrent PRRSV infection. To our knowledge, with the exception of analysis of fecal swabs collected 21 dpi from pigs concurrently infected with PCV2a and

PRRSV (Opriessnig et al., 2008a), PCV2 shedding patterns so far have only been investigated in pigs singularly infected with PCV2 under experimental conditions. The amounts of PCV2 DNA in oral swabs (Fig. 3), nasal swabs (Fig. 4), and fecal swabs (Fig. 5) were an average of 1-3 logs higher in co-infected pigs compared to singularly PCV2- infected pigs, indicating that pigs subclinically co-infected with PRRSV and PCV2 shed much higher amounts of PCV2 for longer periods of time thereby increasing the risk of transmission to other pigs and increased amounts of PCV2 circulating within the herd and the environment. From a practical perspective, it is important to monitor the PRRSV status of herds so that appropriate intervention strategies (i.e. PCV2 vaccines) can be implemented

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to control PCVAD in herds when PRRSV is introduced or becomes active. Similarly, proactive efforts to keep PRRSV out of herds will result in less risk of PCVAD becoming a problem.

5. Conclusion

A higher quantity of PCV2a and PCV2b DNA was found in serum, oral swabs, nasal swabs and fecal swabs in pigs concurrently infected with PCV2 and PRRSV compared to pigs singularly infected with PCV2, further confirming that PRRSV enhances replication of

PCV2. Moreover, PRRSV infection significantly prolonged the presence of PCV2 DNA in serum and several body secretions, thus emphasizing the importance of PRRSV control in order to reduce the PCV2 shedding and transmission in the pig population.

Conflict of interest

None of the authors of this paper has a financial or personal relationship with other people or organizations that could inappropriately influence or bias the content of the paper.

Acknowledgements.

We thank Dr. Joao C. Gomes-Neto, Matthew Umphress, Joseph Thomas and Brett

Kroeze for assistance with the animal work.

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Table 1. Experimental design. Number Inoculationa Group designation of pigs PCV2b PRRSVb Negative controls 3 - - PCV2a-I 5 PCV2a - PCV2a-PRRSV-CoI 5 PCV2a PRRSV PCV2b-I 5 PCV2b - PCV2b-PRRSV-CoI 5 PCV2b PRRSV aPorcine circovirus type 2 (PCV2) and Porcine reproductive and respiratory syndrome virus (PRRSV) inoculation was done on 0 day post inoculation (dpi). bThe following virus isolates were used for inoculation: PCV2a strain 40895, PCV2b strain NC16845 and PRRSV strain VR2385.

Table 2.

Prevalence of porcine circovirus type 2 (PCV2) DNA shed in different sample types (serum samples, oral swabs, nasal swabs, fecal swabs) in pigs experimentally infected with PCV2a, PCV2b, or concurrently infected with porcine reproductive and respiratory syndrome virus (PRRSV) at different days post inoculation (dpi).

Sample type Group 1 3 5 7 9 11 13 14 21 28 35 42 49 56 63 70 PCV2a-I 0/5 0/5 1/5 3/5 4/5 5/5 5/5 5/5 5/5 5/5 3/5 1/5 2/5 0/5 1/5 1/4a PCV2a-PRRSV-CoI 0/5 0/5 0/5 1/5 4/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 Serum samples PCV2b-I 0/5 0/5 0/5 2/5 4/5 5/5 5/5 5/5 4/5 5/5 4/5 2/5 1/5 2/5 0/5 2/5 PCV2b-PRRSV-CoI 0/5 0/5 1/5 2/5 4/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 4/5 5/5

149 PCV2a-I 5/5 1/5 2/5 1/5 5/5 4/5 5/5 5/5 5/5 3/5 2/5 3/5 2/5 4/5 5/5 3/4a

PCV2a-PRRSV-CoI 5/5 5/5 3/5 0/5 4/5 5/5 5/5 5/5 5/5 5/5 4/5 2/5 5/5 4/5 5/5 4/5 Oral swabs PCV2b-I 5/5 3/5 4/5 0/5 2/5 1/5 5/5 3/5 5/5 3/5 1/5 1/5 1/5 0/5 0/5 0/5 PCV2b-PRRSV-CoI 2/5 3/5 2/5 1/5 1/5 4/5 5/5 5/5 5/5 5/5 5/5 4/5 2/5 5/5 5/5 5/5 PCV2a-I 4/5 5/5 2/5 2/5 5/5 2/5 5/5 5/5 5/5 5/5 2/5 4/5 4/5 3/5 3/5 1/4a PCV2a-PRRSV-CoI 4/5 3/5 3/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 4/5 Nasal swabs PCV2b-I 5/5 3/5 4/5 1/5 2/5 1/5 5/5 5/5 4/5 4/5 4/5 3/5 5/5 5/5 4/5 4/5 PCV2b-PRRSV-CoI 5/5 5/5 1/5 2/5 4/5 2/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 4/5 5/5 5/5 PCV2a-I 4/5 4/5 5/5 2/5 5/5 5/5 5/5 4/5 5/5 5/5 5/5 4/5 5/5 5/5 4/5 2/4a PCV2a-PRRSV-CoI 2/5 4/5 5/5 4/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 5/5 Fecal swabs PCV2b-I 3/5 3/5 4/5 1/5 3/5 4/5 5/5 5/5 5/5 5/5 4/5 3/5 5/5 2/5 4/5 5/5 PCV2b-PRRSV-CoI 1/5 5/5 5/5 2/5 4/5 4/5 5/5 5/5 5/5 5/5 5/5 2/5 5/5 5/5 4/5 4/5 aOne animal was euthanized and necropsied on dpi 63.

Table 3.

Mean group amount of log10 porcine circovirus type 2 (PCV2) DNA shed in different sample types (oral swabs, nasal swabs, fecal

swabs) in pigs experimentally infected with PCV2a, PCV2b, or concurrently infected with porcine reproductive and respiratory

syndrome virus (PRRSV) at different days post inoculation (dpi).

Sample Group 7 14 21 28 35 42 49 56 63 70 type

PCV2a-I 0.9±0.9 6.1±0.5A 5.8±0.4A,B 3.1±1.3 1.5±0.9A 2.6±1.1 1.7±1.1A,B 3.3±0.8A 4.0±0.1A 2.7±1.1A,B

A A A,B A A A A Oral PCV2a-PRRSV-CoI 0.0 6.4±0.2 7.1±0.4 4.8±0.3 3.6±0.9 2.1±1.3 5.0±1.3 3.7±1.0 4.5±0.3 3.9±1.0 swabs PCV2b-I 0.0 3.0±1.3B 4.5±0.3B 2.6±1.1 0.8±0.8A 0.9±0.9 0.8±0.8B 0.0B 0.0B 0.0B

PCV2b-PRRSV-CoI 0.9±0.9 6.0±0.4A 6.8±0.3A 5.3±0.2 5.6±0.4B 4.0±1.1 1.6±1.0A.B 5.1±0.2A 5.7±0.3C 5.3±0.1A

150 PCV2a-I 2.2±1.4A,B 6.2±0.3 6.2±0.4A,B 5.1±0.4A,B 2.0±1.2A 4.5±1.2A,B 3.7±0.9A 2.7±1.1 3.3±1.4 0.9±0.9A

A A A A,B A A,B A,B Nasal PCV2a-PRRSV-CoI 5.6±0.2 7.5±0.7 7.9±0.4 6.1±0.5 5.3±0.6 6.7±0.4 5.5±0.5 5.5±0.5 5.0±0.3 3.9±1.0 swabs PCV2b-I 1.0±1.0B 6.0±0.2 4.3±1.1B 3.3±0.8B 4.7±1.3A,B 2.6±1.1B 5.7±0.3A,B 5.1±0.3 4.5±1.2 3.5±0.9A,B

PCV2b-PRRSV-CoI 2.1±1.3A,B 6.6±0.3 7.9±0.3A 6.4±0.4A 6.6±0.9B 5.1±0.3A,B 6.4±0.4B 4.2±1.1 5.5±0.2 5.3±0.2B

7.6±0.6A,B 5.9±0.3A,B 6.7±0.5A,B 3.9±1.1 4.9±0.3 4.7±0.2A,B 3.9±1.0 1.7±1.0A PCV2a-I 2.5±1.5 6.3±1.6

9.4±0.5A 7.4±0.6A 7.6±0.3A 5.5±0.2 5.7±0.4 5.6±0.6A 5.8±0.4 5.1±0.3B PCV2a-PRRSV-CoI 4.3±1.1 7.5±0.9 Fecal swabs 6.3±0.5B 4.8±0.4B 4.0±1.1B 3.8±1.6 4.3±0.3 2.2±1.3B 3.5±0.9 4.2±0.2A.B PCV2b-I 0.9±0.9 6.3±0.3

8.2±0.7A,B 6.7±0.3A 6.5±0.5A,B 2.0±1.2 5.5±0.5 5.0±0.6A,B 4.6±1.2 4.0±1.0A.B PCV2b-PRRSV-CoI 2.0±1.2 6.9±0.5

A,B,C Different superscripts ( ) within a column indicate significant (P<0.05) differences in amount of log10 PCV2 DNA for a sample

type.

151

Fig. 1. Porcine circovirus type 2 mean group sample to negative corrected (SNc) ratios in serum of pigs infected with PCV2a, PCV2b or concurrently infected with porcine reproductive and respiratory syndrome virus (PRRSV). Samples were considered to be positive for anti-PCV2 antibody if the SNc ratio was equal or less than 0.50.

152

Fig. 2. Mean group amount (based on PCR positive and negative pigs within a group) of log10 porcine circovirus type 2 (PCV2) DNA in serum of pigs infected with PCV2a, PCV2b or concurrently infected with porcine reproductive and respiratory syndrome virus

(PRRSV). A sample with no threshold cycle (CT) value during the 40 amplification cycles was considered negative. The detection limit of PCV2 quantitative real-time PCR assay was 1×103 copies/ml.

PCV2-I PCV2-PRRSV-CoI 8 * 7 * 10/10 10/10 10/10 6 * 10/10 * 9/10 10/10 *10/10 * 10/10 5 * 10/10 * 9/10 9/10 8/10 9/10 8/10 4 7/10 * *

7/10 3 7/10 6/10 6/10 6/10 5/10 5/10

7/10 5/10 153 2 4/10 4/10 4/10

Mean group log10 PCV2 DNA in oral swabs oral in DNA PCV2log10group Mean 3/9 3/10 1 3/10 1/10

0 1 3 5 7 9 11 13 14 21 28 35 42 49 56 63 70 Days post inoculation

Fig. 3. Mean group amount (based on PCR positive and negative pigs within a group) of log10 porcine circovirus type 2 (PCV2) DNA in oral swabs of pigs singularly infected with PCV2 (PCV2-I, combined PCV2a-I and PCV2b-I pigs, n=10) or concurrently infected with porcine reproductive and respiratory syndrome virus (PRRSV) (PCV2-PRRSV-CoI, combined PCV2a-PRRSV-CoI and PCV2b-PRRSV-CoI pigs, n=10). A sample with no threshold cycle (CT) value during the 40 amplification cycles was considered negative. Prevalence of PCV2 DNA positive pigs is indicated next to each data point. Asterisks indicate significant (P < 0.05) differences between groups. The detection limit of PCV2 quantitative real- time PCR assay was 1×103 copies/ml

PCV2-I PCV2-PRRSV-CoI 9

8 * 10/10 * 7 * 10/10 10/10 * 10/10 10/10 * 6 10/10 10/10 10/10 10/10 9/10 10/10 * 5 9/10 9/10 9/10 9/10 9/10 8/10 4 7/10 9/10 8/10 7/10 7/10 8/10 7/10 7/10 6/10 3 6/10 5/9 2 4/10

154

3/10 3/10 Mean group log10 PCV2 DNA in nasal swabs nasal in DNA PCV2log10group Mean

0 1 3 5 7 9 11 13 14 21 28 35 42 49 56 63 70 Days post inoculation

Fig.4. Mean group amount (based on PCR positive and negative pigs within a group) of log10 porcine circovirus type 2 (PCV2) DNA in nasal swabs of pigs singularly infected with PCV2 (PCV2-I, combined PCV2a-I and PCV2b-I pigs, n=10) or concurrently infected with porcine reproductive and respiratory syndrome virus (PRRSV) (PCV2-PRRSV-CoI, combined PCV2a-PRRSV-CoI and PCV2b-

PRRSV-CoI pigs, n=10). A sample with no threshold cycle (CT) value during the 40 amplification cycles was considered negative. Prevalence of PCV2 DNA positive pigs is indicated next to each data point. Asterisks indicate significant (P <

0.05) differences between groups. The detection limit of PCV2 quantitative real-time PCR assay was 1×103 copies/ml.

PCV2-I PCV2-PRRSV-CoI 10 * 9 10/10

8 10/10 * * 7 10/10 10/10 10/10 10/10 6 * 9/10 10/10 * * 10/10 10/10 10/10 9/10 9/10 9/10 5 9/10 9/10 9/10 8/10 10/10 9/10 4 7/10 8/10 7/10 7/10 7/10 3 6/10 7/9

155 Mean group log10 PCV2 DNA in fecal swabs fecal in DNA PCV2log10group Mean 3/10 3/10

0 1 3 5 7 9 11 13 14 21 28 35 42 49 56 63 70 Days post inoculation

Fig. 5. Mean group amount (based on PCR positive and negative pigs within a group) of log10 porcine circovirus type 2 (PCV2) DNA in fecal swabs of pigs singularly infected with PCV2 (PCV2-I, combined PCV2a-I and PCV2b-I pigs, n=10) or concurrently infected with porcine reproductive and respiratory syndrome virus (PRRSV) (PCV2-PRRSV-CoI, combined PCV2a-PRRSV-CoI and PCV2b-PRRSV-CoI pigs, n=10). A sample with no threshold cycle (CT) value during the 40 amplification cycles was considered negative. Prevalence of PCV2 DNA positive pigs is indicated next to each data point. Asterisks indicate significant (P < 0.05) differences between groups. The detection limit of PCV2 quantitative real-time PCR assay was 1×103 copies/ml.

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CHAPTER 6: PRRSV INFECTION AT THE TIME OF PCV2 VACCINATION HAS

NO IMPACT ON VACCINE EFFICACY

A paper published in

Clinical Vaccine and Immunology 17:1940-1945, 2010

A. Sinha, H.G. Shen, S. Schalk, N.M. Beach, Y.W. Huang,

P.G. Halbur, X.J. Meng, T. Opriessnig

ABSTRACT

Several porcine circovirus type 2 (PCV2) vaccines are now commercially available and have been shown to be effective at decreasing the occurrence of porcine circovirus associated disease (PCVAD). Many herds are coinfected with PCV2 and porcine reproductive and respiratory syndrome virus (PRRSV). Some producers and veterinarians are concerned that if pigs are vaccinated for PCV2 at or near the time they are typically infected with PRRSV, the efficacy of the PCV2 vaccine will be compromised. The impact of PRRSV on PCV2 vaccination is unclear and has not been investigated under controlled conditions. The objective of the present study was to determine whether the presence of PRRSV viremia has an effect on the efficacy of commercial PCV2 vaccinations. Three-week-old PCV2-negative conventional pigs with passively derived anti-PCV2 antibodies were either vaccinated with one of three commercial PCV2 vaccines or left non-vaccinated. A portion of the pigs were infected with PRRSV 1 week prior to PCV2 vaccination. To determine vaccine efficacy, a

PCV2 challenge was conducted at 8 weeks of age. PCV2 vaccination regardless of PRRSV

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infection status at the time of vaccination, was similarly effective in inducing an anti-PCV2

IgG response in the presence of maternally derived immunity and in protecting the pigs from

PCV2 challenge, as determined by reduction in the level of PCV2 viremia and a reduction in the prevalence and amount of PCV2 antigen in lymphoid tissues compared to nonvaccinated pigs. The results indicate that acute PRRSV infection at the time of PCV2 vaccination has no adverse effect on PCV2 vaccine efficacy.

INTRODUCTION

Porcine circovirus type 2 (PCV2) is a single-stranded, circular, nonenveloped DNA virus with an icosahedral symmetry (6, 44). It has an ambisense genome of approximately

1.8 kb and belongs to the family of Circoviridae which are known to be very host-specific and often associated with subclinical infections (36, 45). PCV2 associated disease

(PCVAD), first recognized in 1991 as postweaning multisystemic wasting syndrome

(PMWS) (15), has spread throughout the global swine population. Besides the systemic presentation of PCV2 infection, other disease manifestations of PCVAD include respiratory disease, enteritis in grow-finisher pigs, reproductive failure, and porcine dermatitis and nephropathy syndrome (PDNS) (27). Lymphoid depletion and granulomatous inflammation in several organ systems are the hallmark lesions of PCVAD (41).

Pigs concurrently infected with PCV2 and other pathogens have a greater potential of developing clinical PCVAD (27). In a retrospective study in Korea, 76.2% (80/105) of pigs suffering from porcine respiratory disease complex (PRDC) were found to be co- infected with PCV2 and various other pathogens including porcine reproductive and respiratory syndrome virus (PRRSV), swine influenza virus (SIV) and porcine parvovirus

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(PPV) (18). In another investigation in the United States, PRRSV was present in 51.9% of

484 cases of systemic PCVAD and was the most frequent pathogen (34). In a cross-sectional study performed on 583 conventionally reared pigs of different ages, coinfections of PCV2 with PRRSV, SIV, and Mycoplasma hyopneumoniae had the greatest effect on the development of diseases in the early to late nursery phases (8). In a case-control study of pigs with and without PCVAD, pigs with concurrent PCV2 and PRRSV infection had a higher odds ratio of developing PCVAD (35).

PRRSV is a single-stranded, positive-sense, enveloped RNA virus that belongs to the family Arteriviridae, genus Arterivirus (4). PRRSV primarily infects the monocyte/macrophage cell line (11) via CD 163 scavenger receptor and heparin sulfate and sialoadhesin receptors (10, 46) which are also involved in the primary defense of the innate immune system (9, 21). It causes reproductive failure in pregnant sows and respiratory disease in pigs of all ages, and is also associated with neonatal diarrhea (1, 3, 23, 37). Under experimental conditions, with concurrent PRRSV and PCV2 infection in 1-to-2-day-old colostrum-deprived (CD) pigs (2), 5-week-old PCV2 seropositive pigs (38), or 3-week-old cesarean-derived colostrum-deprived (CD/CD) pigs (16), more severe clinical disease and lesions as well as enhanced replication and distribution of PCV2 were observed compared to the conditions of pigs inoculated with PCV2 alone. These results suggest that immune modulation by PRRSV may be a key factor in the development of PCVAD.

PRRSV can be transmitted transplacentally resulting in infected neonates, and

PRRSV viremia may last up to 154 days with virus being detectable in tissues for up to 202 days (20). PRRSV seroconversion and viremia typically occur during the nursery phase of production. It is a common practice for veterinarians and producers to adjust the timing of

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administration (delay or move earlier) of common vaccinations such as M. hyopneumoniae

(9) to avoid vaccine failure associated with PRRSV-induced immunosuppression.

At present, no information exists on the efficacy of PCV2 vaccination in PRRSV viremic pigs. The objective of this study was to determine the effect of PRRSV infection at the time of PCV2 vaccination on PCV2 vaccine efficacy. Three currently available commercial PCV2 vaccines were compared in the conventional pig model using pigs with passively-acquired anti-PCV2 antibodies with or without experimental PRRSV infection.

MATERIALS AND METHODS

Animals and housing. Ninety-nine, specific pathogen free (SPF) pigs were weaned at 2 weeks of age from high-health sows free of PRRSV and SIV. PCV2 vaccination was not used in the breeding herd. The dams of the piglets used in this study were not PCV2 viremic

(negative for PCV2 DNA on serum evaluation) but were positive for PCV2 antibody, antibody as determined by PCV2 enzyme linked immunosorbent assay (ELISA) approximately 1 week before farrowing. Upon arrival at the research facility at Iowa State

University, the pigs were housed in four separate rooms: the non-infected negative control group and the group infected with PRRSV alone were housed in separate rooms, and all other pigs regardless of treatment group were randomly assigned to one of two rooms, which each contained four pens. The pens were 2 m × 2.3 m in size, and each one was equipped with one nipple drinker and a self feeder that contained a complete feed ration free of animal proteins (excluding whey) and antibiotics (Nature’s Made, Heartland Coop, Iowa, USA).

Experimental Design. The 99 pigs were randomly divided into 10 groups of 9 to 11 pigs in each group after arrival at the Iowa State University research facility. The

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experimental design is summarized in Table 1. At 2 weeks of age, 51 out of 99 pigs were inoculated with PRRSV. At 3 weeks of age, portions of the PRRSV-infected (n=29) and non-infected pigs (n=29) were vaccinated with one of three commercially available vaccines, as described in Table 1. At 5 weeks of age, 2 groups (n=20) were each given a booster dose of vaccines. At 10 weeks of age, 79 pigs were challenged with PCV2b. All pigs were necropsied at 13 weeks of age. Blood samples were collected from each group on the day of arrival at the facility and weekly thereafter until necropsy. The serum samples collected on the day of PCV2 challenge and at days post challenge (dpc) 7, 14, and 21 were tested for the presence and amount of PRRSV RNA by reverse transcription (RT) real-time

PCR and for PCV2 DNA by quantitative real-time PCR. The experimental protocol was approved by the Iowa State University Institutional Animal Care and Use Committee

(IACUC; approval number: 8-08-6618-S).

PRRSV inoculation. At two weeks of age, 51 pigs (in the PRRSV-Fort Dodge

Animal Health (FDAH)-PCV2, PRRSV-Boehringer Ingelheim Vetmedica Inc.(BIVI)-

PCV2, PRRSV-Intervet-PCV2, PRRSV, and PRRSV-PCV2 groups) were inoculated with

5 approximately 10 50% tissue culture infective doses (TCID50) of PRRSV isolate VR2385 as described (40). Each pig received 2 ml of the PRRSV inoculum intranasally by slowly dripping 1 ml into each nostril.

Vaccination. Three commercial vaccines were used in this experimental study. At 3 weeks of age, pigs in the PRRSV-FDAH-PCV2 and FDAH-PCV2 groups were vaccinated intramuscularly with 2 ml of Suvaxyn® PCV2 vaccine (serial number 1861220A; Fort

Dodge Animal Health, Inc., Fort Dodge, IA). Similarly, pigs in the PRRSV-BIVI-PCV2 and

BIVI-PCV2 groups were vaccinated intramuscularly with 1 ml of Ingelvac CircoFLEX

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vaccine (serial number 309-136; Boehringer Ingelheim Vetmedica Inc.) at 3 weeks of age.

Pigs in the PRRSV-Intervet-PCV2 and Intervet-PCV2 groups were vaccinated intramuscularly using 2 ml of Circumven vaccine (serial number 02137920; Intervet

Schering-Plough Animal Health;) at 3 and 5 weeks of age.

PCV2 challenge. At ten weeks of age, pigs in groups PRRSV-FDAH-PCV2,

PRRSV-BIVI-PCV2, PRRSV-Intervet-PCV2, FDAH-PCV2, BIVI-PCV2, Intervet-PCV2,

PRRSV-PCV2 and PCV2 were challenged with PCV2b. PCV2b isolate NC16845 (31) was

4.5 used at a dose of 10 TCID50 per ml. Each pig received 2 ml intranasally and 2 ml intramuscularly.

Clinical evaluation. All the pigs were weighed on the day of arrival, the day of vaccination (3 and 5 weeks of age), the day of challenge (10 weeks of age; 0 dpc), and the day of necropsy (21 dpc); and the average daily weight gain (ADWG) was calculated. Other cliniocal parameters were not evaluated.

Anti-PCV2-IgG-antibodies. All serum samples collected prior to PCV2 challenge

(2, 3, 4, 5, 6, 7, 8, 9 and 10 weeks of age) and at dpc 7, 14, and 21 were tested using the

SERELISA® PCV2 Ab Mono Blocking kit (Synbiotics Europe, Lyon, France) according to the manufacturers’ instructions. Results were expressed as a sample to negative corrected

(SNc) ratio. Samples were considered positive for the presence of anti-PCV2 antibodies if the calculated SNc ratio was less than or equal to 0.50, and they were considered negative if the SNc ratio was greater than 0.50.

PCV2 DNA detection and quantification. The DNA from serum samples collected at dpc 7, 14, and 21 was extracted using the QIAamp DNA minikit (Qiagen, Valencia, CA).

The DNA was tested for the presence and amount of PCV2 DNA using a quantitative real-

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time PCR as described previously (33).

Anti-PRRSV-IgG-antibodies. Serum samples collected from pigs at 2 weeks of age and at dpc 0 and 21 were tested for the presence of antibodies to PRRSV (HerdChek X

PRRS virus antibody test kit, IDEXX Laboratories Inc., Westbrook, MA). A sample-to- positive (S/P) ratio of equal to or greater than 0.4 was considered a positive result.

PRRSV RNA detection and quantification. RNA extraction from serum samples collected at the day of PCV2 vaccination, 7 days after PCV2 vaccination, and dpc 0, 7, 14, and 21 from all the PRRSV-inoculated groups was performed using a QIAamp Viral RNA minikit (Qiagen). Detection and quantification of PRRSV genomic RNA from the RNA extracts was achieved by real time RT-PCR as described previously (40).

Necropsy. All pigs were humanely euthanized by pentobarbital overdose (Fatal-plus;

Vortex Pharmaceutical, LTD; Dearborn, MI) and necropsied at 13 weeks of age (21 dpc).

The total amount of lung surfaces affected by macroscopic lesions (ranging from 0 to 100%)

(14) and size of lymph nodes with scores ranging from 0 (normal size) to 3 (four times the normal size) (32) were estimated in a blinded fashion. Sections of liver, spleen, colon, kidney, lymph nodes (mesenteric, superficial inguinal, tracheobronchial, and mediastinal), ileum, tonsil and thymus were collected at necropsy; fixed in 10% neutral-buffered formalin and processed routinely for histological examination.

Histopathology. Microscopic lesions were evaluated by a veterinary pathologist

(T.O.) blinded to the treatment groups. Sections of heart, liver, kidney, ileum and colon were evaluated for the presence of lymphohistiocytic inflammation and were scored from 0

(none) to 3 (severe). Lung sections were scored for the presence and severity of interstitial pneumonia, with scores ranging from 0 (normal) to 6 (severe, diffuse) (14). Lymphoid

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tissues including lymph nodes, tonsil and spleen, were evaluated for the presence of lymphoid depletion, with scores ranging from 0 (normal) to 3 (severe) and histiocytic inflammation and replacements of follicles, with scores ranging from 0 (normal) to 3

(severe) (32).

IHC. Immunohistochemistry (IHC) staining for the detection of PCV2 antigen was conducted on selected formalin-fixed and paraffin-embedded sections of lymph nodes

(mesenteric, superficial inguinal, tracheobronchial, and mediastinal), spleen, tonsil, and thymus using a rabbit polyclonal antiserum (42). PCV2 antigen scoring was done by a pathologist blinded to the treatment status of the animals, with scores ranging from 0 (no signal) to 3 (abundant amount of PCV2 antigen) as described previously (32).

Overall lymphoid lesions score. The overall microscopic lymphoid lesion score, which accounts for lymphoid depletion, histiocytic inflammation, and amount of PCV2 antigen, was calculated as previously described (32).

Statistical Analysis. Statistical analysis of the data was performed using the JMP software version 8.0.1 (SAS Institute, Cary, NC). Summary statistics were calculated for all groups to assess the overall quality of the data including normality. Continuous data were analyzed using the one-way analysis of variance (ANOVA). A P value of less than 0.05 was set as a statistically significant level throughout this study. If an ANOVA was significant, pairwise testing using Tukey's adjustment was performed to assess specific group differences. Real-time PCR results (number of copies per ml of serum) were log10 transformed prior to statistical analysis. Percent reduction for PCR data was measured as follows: 100 − [(100 × mean log10 genomic copies/ml in the vaccinated group) / (mean log10 genomic copies/ml in positive control animals)]. Nonrepeated measures of necropsy and

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histopathology data were assessed using a nonparametric Kruskal-Wallis one-way ANOVA.

If this nonparametric ANOVA test was significant (P < 0.05), then pairwise Wilcoxon tests were used to assess differences between groups. In order to determine differences in prevalence, a chi square test was used.

RESULTS

ADWG. There were no significant (P > 0.05) differences in initial body weights among the groups of pigs at arrival in the research facility. After PRRSV infection, between

2 and 3 weeks of age the ADWG in PRRSV-infected pigs was significantly (P < 0.05) lower than that in non-infected pigs (0.25 ± 0.01 kg versus 0.28 ± 0.01 kg). Similarly, from 3 to 10 weeks of age, the ADWG in PRRSV-infected pigs was significantly (P < 0.0001) lower than that in non-infected pigs (0.66 ± 0.01 kg versus 0.77 ± 0.01 kg). PCV2 vaccination had no effect on ADGW as there was no significant (P > 0.05) difference between the vaccinated and non-vaccinated groups from 3 to 10 weeks of age. The ADWG for the individual groups from the time of PCV2 challenge until necropsy (10 to 13 weeks of age) is summarized in

Table 2. There was no significant (P > 0.05) difference between PRRSV-infected and noninfected pigs or between vaccinated and nonvaccinated pigs. However, the ADWG in

PCV2-infected pigs was significantly (P < 0.05) lower than that in non-infected pigs (1.00 ±

0.02 kg versus 1.08 ± 0.05 kg).

Anti-PCV2-IgG antibody levels. At arrival, 95/99 pigs had passively acquired

PCV2 antibodies, with group mean SNc ratios between 0.00 and 0.10 (Fig. 1), and no significant differences were observed among different treatment groups. The group mean

PCV2 ELISA SNc ratios in vaccinated pigs were significantly (P < 0.0001) lower than those

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in non-vaccinated pigs between 5 weeks of age (vaccinated pigs, 0.22 ± 0.04; non- vaccinated pigs, 0.39 ± 0.04) and 10 weeks of age (vaccinated pigs, 0.06±0.02; non- vaccinated pigs, 0.51 ± 0.04). Significantly (P < 0.05) higher SNc ratios (lower levels of antibody) were observed in the BIVI groups than in the FDAH and Intervet groups during the growing period; however, this was independent of PRRSV status. At the day of PCV2 challenge at 10 weeks of age, 98% (58/59) of the vaccinated pigs had detectable anti-PCV2 antibodies whereas only 42.5% (17/40) of the non-vaccinated pigs were seropositive. After

PCV2 challenge (dpc 7, 14, and 21), vaccinated groups had significantly (P < 0.001) lower

SNc ratios than non-vaccinated challenge pigs, which were independent of the product used or PRRSV infection status at the time of vaccination and at 14 and 21 dpc.

Prevalence and amount of PCV2 DNA in serum. Pigs in the negative-control group and in the PRRSV group remained negative for PCV2 DNA throughout the study period. After PCV2 challenge, PCV2 DNA was detected in the majority of the non- vaccinated pigs at 7 (70%), 14 (100%) and 21 (100%) dpc. Among the vaccinated pigs,

PCV2 viremia was limited to a few individual pigs (Table 3). Vaccination significantly (P <

0.05) reduced the amount of PCV2 compared to that in non-vaccinated pigs (Table 3). The mean reductions of PCV2 viremia in the serum at 21 dpc was 88.6% for FDAH-PCV2,

88.8% for BIVI-PCV2, 90.2% for Intervet-PCV2, 89.4% for PRRSV-FDAH-PCV2, 66.8% for PRRSV-BIVI-PCV2, and 100% for PRRSV-Intervet-PCV2.

Anti-PRRSV-IgG antibody levels. All pigs were negative for anti-PRRSV antibodies at the start of the study at 2 weeks of age (data not shown). All pigs that were not experimentally infected with PRRSV remained seronegative for PRRSV until the termination of the study at 13 weeks of age. At 10 weeks of age (dpc 0), the mean PRRSV

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ELISA S/P ratio values were 2.30 ± 0.11 for PRRSV infected , non-PCV2 vaccinated pigs

(n=21) and 2.17 ± 0.09 for PRRSV-infected, PCV2-vaccinated pigs (n=30) with no significant (P > 0.05) difference existing between groups. Similarly, there was no significant

(P > 0.05) difference in mean group PRRSV S/P ratios on dpc 21 between PCV2-vaccinated

(1.78 ± 0.11) and non-vaccinated (2.08 ± 0.13) pigs or between PCV2 challenged (1.87 ±

0.10) and nonchallenged (2.04±0.17) pigs.

Prevalence and amount of PRRSV RNA in serum. At the time of PCV2 vaccination (7 days after PRRSV inoculation) and 7 days after PCV2 vaccination (14 days after PRRSV inoculation), 100% of the pigs in the PRRSV challenged groups were positive for PRRSV RNA in serum, with PRRSV RNA loads ranging from 6.16 to 6.66 log10 copies/ml (Table 4). From 0 to 21 dpc, low levels of PRRSV RNA were detected in a few (1 to 3) pigs in all PRRSV-infected groups (Table 4). No significant (P > 0.05) differences were observed for PRRSV prevalence or RNA loads among the different treatment groups.

Macroscopic, microscopic lesions and PCV2 antigen in lesions. Macroscopic lung lesions were characterized by failure of the lungs to collapse and by focal-to-diffuse, mottled tan areas of pneumonia. Vaccinated, PCV2-challenged pigs (n=59) had significantly (P <

0.05) lower mean lung scores than non-vaccinated-PCV2 infected pigs (n=20) (1.97 ± 0.32 versus 3.95 ± 1.00). Moreover, PRRSV-negative PCV2-challenged pigs (n = 39) had significantly (P < 0.05) lower mean lung scores than PRRSV-positive PCV2-challenged pigs (n = 40) (1.46 ± 0.28 versus 3.45 ± 0.62). There was no difference in the severity of macroscopic lung lesions among vaccinated groups (P > 0.05). In the PRRSV-PCV2 group,

9/10 pigs had lymph nodes of 2 to 3 times the normal size. In the remaining groups individual pigs (1 to 2 per group) had mildly enlarged lymph nodes.

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Microscopic lesions were characterized by mild to severe lymphoid depletion and mild to moderate histiocytic replacement in lymphoid tissues. Among vaccinated pigs,

83.3% (49/59) had normal lymphoid tissues and 16.9% (10/59) had mild lymphoid lesions

(overall lymphoid scores 1 or 2). Non-vaccinated and PCV2 infected pigs however, had the following lesion distribution: 10% (2/20) had normal lymphoid tissues, 45% (9/20) had mild lymphoid lesions (scores 1, 2, or 3), 40% (8/20) had moderate lymphoid lesions (scores 4, 5, or 6), and 5% (1/20) had severe PCV2 associated lesions (score, 7). There was no difference in lesion severity between PRRSV-infected and noninfected pigs.

PCV2 antigen was detected in 8/10 pigs in the PRRSV-PCV2 group, 7/10 pigs in the

PCV2 group, and in 0/59 pigs in the vaccinated groups; the differences were statistically significant (P < 0.05).

DISCUSSION

Various commercially available killed PCV2 vaccines are being used worldwide, and their efficacies have been demonstrated under both experimental conditions (13, 25, 29, 30,

40) and field conditions (5, 12, 17, 19, 39). PCV2 vaccination is recommended for usage in weaned pigs at about 3 weeks of age or older. However, at this age many pigs derived from

PRRSV positive breeding herds are infected with PRRSV, which is thought to negatively impact PCV2 vaccine efficacy. In the present study we evaluated PCV2 vaccine efficacy in

PRRSV-positive conventional pigs by experimental inoculation with a known virulent isolate of PRRSV (14,24, 28). It is well known that PRRSV strains in virulence (28) and the results of this study may be different with other PRRSV challenge isolates.

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Increased PCV2 replication and clinical PCVAD were observed in pigs experimentally coinfected with PRRSV and PCV2 (2, 16, 38). Although PRRSV-infected pigs had significantly increased macroscopic lung lesions in the present study, significant differences in the severity of microscopic lesions or amount of PCV2 antigen were not observed in PCV2-challenged pigs. A possible explanation for the discrepancy in observations is the difference in timing of pathogen administration: in the current study, the pigs were infected with PRRSV at 2 weeks of age, followed by PCV2 challenge at 10 weeks of age, whereas previously, coinfections were conducted at the same time (16) or within an interval of a few days (2, 38) and were done in younger pigs (1 to 35 days of age).

Although PCV2 vaccines are in general very effective, cases of apparent vaccine failures continue to occur. Vaccine failures under field conditions may be due to many reasons, including failure to follow the proper administration protocols and to use the recommended dose of the vaccines (30). The field strains might be different from the vaccine strains, and the breed of the pigs can also influence the outcome of PCVAD (22,

30).

The presence of high levels of passively derived anti-PCV2 antibodies at the time of vaccination has also been associated with decreased vaccine efficacy and vaccine failures.

Experimental proof of this is lacking to date, as available research results indicate no adverse effect of maternally derived immunity on PCV2 vaccine efficacy (13, 29, 40). Similarly, the results of the current study indicate that PCV2 vaccination in the face of high levels of passively derived anti-PCV2 immunity was effective in inducing a humoral immune response. At 10 weeks of age, a significant higher prevalence of seropositive pigs was found in the vaccinated group than in the nonvaccinated control group.

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Several studies have investigated the effect of PRRSV infection on vaccine efficacy but provided conflicting results. PRRSV infection at the time of classical swine fever virus vaccination significantly inhibited the host immune response and resulted in vaccination failure after subsequent exposure to classical swine fever virus (7, 43). For pseudorabies virus vaccination, although PRRSV infection affected the duration of the T-lymphocyte response, it did not inhibit the efficacy of the vaccine, which was capable of inducing protective immunity (7). In the present study, our data revealed that all the commercial vaccines used in this study were able to induce a protective immunity against PCV2 in the face of PRRSV infection at the time of vaccination. The difference in the experimental results might be due to different timings of infection with PRRSV, vaccination, and challenge. In the present study, pigs were challenged 5 weeks (two dose administrations) to 7 weeks (one dose administration) after vaccination, in contrast to a study where challenge occurred 3 weeks after initial vaccination (43).

In a previous study (26), we found that PCV2 infection had adverse effects on the efficacy of a modified live virus (MLV) PRRSV vaccine. The different effects of PRRSV and PCV2 infection on the efficacy of vaccines may be due to the different mechanisms that these two viruses use to induce immunosuppression (replication in macrophages versus lymphoid depletion), which are still not completely understood, and the types of vaccine

(attenuated live versus inactivated) tested.

All PCV2 vaccines used, regardless of PRRSV infection status at the time of vaccination, were able to suppress PCV2 replication after PCV2 challenge, as demonstrated by significantly reduced levels of PCV2 viremia and significantly reduced amounts of PCV2 antigen in lymphoid tissues. The results of our study demonstrated that by 21 dpc there was

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no difference in the prevalence of PCV2 DNA in vaccinated groups regardless of PRRSV infection status at the time of vaccination or the vaccine product used. We conclude that prior PRRSV infection does not have an adverse effect on commercial PCV2 vaccination in conventional growing pigs and that pigs can be effectively immunized against PCV2 in the face of maternal antibodies.

ACKNOWLEDGEMENT

We thank Matthew Umphress for assistance with the animal work. Funding for this study was provided by the National Pork Board CheckOff Dollars.

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TABLE 1. Experimental design Group designation No. PRRSV infection PCV2 vaccine amt (ml) Challenge of (2 wk) 3 wk 5 wk (10 wk) pigs

PRRSV-FDAH-PCV2 10 PRRSV 2 PCV2

FDAH-PCV2 10 2 PCV2

PRRSV-BIVI-PCV2 10 PRRSV 1 PCV2

BIVI-PCV2 9 1 PCV2

PRRSV-Intervet-PCV2 10 PRRSV 2 2 PCV2

Intervet-PCV2 10 2 2 PCV2

PRRSV 11 PRRSV

PCV2 10 PCV2

PRRSV-PCV2 10 PRRSV PCV2

Negative Controls 9

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TABLE 2. ADWG from 0 to 21 dpc

Group ADWG ± SE (Kg)a

PRRSV-FDAH-PCV2………………………………………………………………………………… 1.02 ± 0.03A,B,C

FDAH-PCV2…………………………………………………………………………………………… 0.96 ± 0.03B,C

PRRSV-BIVI-PCV2………………………………………………………………………………….. 1.12 ± 0.04A,B

BIVI-PCV2……………………………………………………………………………………………… 1.03 ± 0.03A,B,C

PRRSV-Intervet-PCV2……………………………………………………………………………. 0.99 ± 0.05B,C

Intervet-PCV2……………………………………………………………………………………….. 1.00 ± 0.05A,B,C

PRRSV…………………………………………………………………………………………………… 0.98 ± 0.06B,C

PCV2…………………………………………………………………………………………………….. 0.91 ± 0.05C

PRRSV-PCV2………………………………………………………………………………………….. 0.98 ± 0.06B,C

Negative Controls…………………………………………………………………………………. 1.21 ± 0.06A a Different superscripts (A, B, and C) within the same column represent significant differences (P < 0.05) between groups.

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TABLE 3. PCV2 DNA-positive pigs among all pigs I each group at different days after PCV2 challenge

No. of pigs PCV2 DNA positive /no. of pigs in group on the a Group following days after PCV2 challenge :

7 14 21

PRRSV-FDAH-PCV2 1/10 (0.42±0.4)A 0/10 (0.00±0.0)A 0/10 (0.00±0.0)A

FDAH-PCV2 1/10 (0.38±0.4)A 0/10 (0.00±0.0)A 0/10 (0.00±0.0)A

PRRSV-BIVI-PCV2 2/10 (0.89±0.6)A 3/10 (1.56±0.8)B 2/10 (0.81±0.5)A

BIVI-PCV2 4/9 (1.98±0.8)AB 2/9 (0.92±0.6)BA 1/9 (0.41±0.4)A

PRRSV-Intervet-PCV2 0/10 (0.00±0.0)A 0/10 (0.00±0.0)A 0/10 (0.00±0.0)A

Intervet-PCV2 0/10 (0.00±0.0)A 0/10 (0.00±0.0)A 1/10 (0.36±0.4)A

PRRSV 0/11 (0.00±0.0)A 0/11 (0.00±0.0)A 0/11 (0.00±0.0)A

PCV2 7/10 (3.67±0.8)B 10/10 (6.69±0.4)C 10/10 (5.96±0.3)B

PRRSV-PCV2 7/10 (3.94±0.9)B 10/10 (7.22±0.2)C 10/10 (7.23±0.3)B

Negative Controls 0/9 (0.00±0.0)A 0/9 (0.00±0.0)A,B 0/9 (0.00±0.0)A

aDifferent superscripts (A, B, and C) within same column represent significant differences

(p<0.05) in mean amount of log10 PCV2 DNA between groups. Values in parentheses

represent the mean log10 number of PCV2 DNA copies ± standard error for the group.

TABLE 4. PRRSV RNA-positive pigs among all pigs in each group for pigs experimentally inoculated with PRRSV 7 days prior to PCV2 vaccination and 8 weeks prior to PCV2 challenge

Group No. of pigs PRRSV RNA positive /no. of pigs in group on the indicated days aftera:

PCV2 vaccination PCV2 challenge

0 7 0 17 14 21

10/10 (6.16±0.3) 10/10 (6.66±0.3) 0/10 (0.00±0.0) 1/10 2/10 0/10 PRRSV-FDAH-PCV2 (0.43±0.4) (0.96±0.6) (0.00±0.0)

10/10 (6.35±0.1) 10/10 (6.40±0.3) 0/10 (0.00±0.0) 1/10 2/10 2/10 PRRSV-BIVI-PCV2 (0.32±0.3) (0.66±0.4) (0.67±0.5)

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10/10 (6.29±0.2) 10/10 (6.33±0.2) 2/10 (0.63±0.4) 1/10 3/10 2/10 PRRSV-Intervet-PCV2 (0.34±0.3) (1.07±0.6) (0.00±0.0)

11/11 (6.48±0.2) 11/11 (6.59±0.4) 0/11 (0.00±0.0) 1/11 1/11 0/11 PRRSV (0.32±0.3) (0.37±0.4) (0.00±0.0)

10/10 (6.44±0.3) 10/10 (6.29±0.4) 2/10 (0.69±0.5) 1/10 2/10 2/10 PRRSV-PCV2 (0.28±0.3) (0.83±0.6) (0.79±0.5)

a Values in parentheses represent the mean log10 PRRSV RNA copies ± standard error of the group.

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FIG. 1. Mean anti-PCV2 IgG antibody responses in the different treatment groups following PCV2 challenge. PRRSV inoculation, the first vaccination (Vac), challenge, and necropsy were performed at 2, 3, 10, and 13 weeks of age, respectively. Samples with an SNc ratio of 0.5 or greater are considered negative. Error bars indicate standard errors.

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CHAPTER 7: GENERAL CONCLUSIONS

Porcine circovirus type 2 (PCV2), the primary causative agent of porcine circovirus associated disease (PCVAD), was first associated with wasting in Canadian pigs in 1991

(Harding and Clark, 1997). PCV2, which is ubiquitously distributed in the swine populations worldwide, causes great economic loss in swine producing countries (Segalés et al., 2005a) and mainly affects pigs between the ages of 4-10 weeks after the decline of anti-PCV2 passive antibodies(Opriessnig et al., 2004). PCVAD is a multisystemic disease and can manifest as post-weaning multisystemic wasting syndrome (PMWS), respiratory disease, enteritis, reproductive failure, and porcine dermatitis and nephropathy syndrome (Opriessnig et al., 2007). PCVAD is also a multi-factorial disease, and co-infections generally lead to increased and aggravated clinical PCVAD (Ellis et al., 2004; Pallarés et al., 2002).

Concurrent infection of porcine reproductive and respiratory syndrome virus (PRRSV) and

PCV2 is commonly associated with more severe clinical signs in affected farms (Pesch et al., 2003; Pogranichniy et al., 2002; Wellenberg et al., 2004).

Increased PCV2 viremia can be commonly seen in pigs coinfected with PRRSV and

PCV2 (Rovira et al., 2002) and is thought to be due to PRRSV enhancement of PCV2 replication (Allan et al., 2000). The North American spread of clinical PCVAD in 2005, which was characterized by increased morbidity and mortality in growing pigs, was associated with PCV2b, a newly recognized subtype. As PRRSV is also common in North

America, this has raised concerns about specific PCV2-PRRSV interactions and increased transmission of PCV2b in PRRSV positive flows. It is unknown whether PRRSV increases

PCV2 infection via enhancement of apoptosis in the host, increased shedding of PCV2,

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immunomodulating of the host response system, decreasing the efficacy of PCV2 vaccines, or any combination thereof, but many of these mechanisms are poorly understood and are important to study.

The present thesis has been devoted to characterize the role of PRRSV in PCV2 subtype a or b infection in co-infected pigs including: apoptosis, immune modulation, PCV2 shedding, and PCV2 vaccine efficacy. The information created by this work is important for swine veterinarians and producers developing and implementing PCV2 prevention and control strategies, veterinary diagnosticians investigating PCVAD cases of co-infections, and researchers studying immunological responses in PCV2 co-infections.

Hepatic failure is the major cause of death in gnotobiotic pigs affected by PCVAD; however, it is not known what factors mitigate the hepatic failure. Many researchers have attempted to establish the role of PCV2 in producing lymphoid depletion and hepatic failure via apoptosis, but have been unsuccessful (Resendes et al., 2004; Resendes et al., 2010;

Rosell et al., 2000; Shibahara et al., 2000). The main aim of our first study was to determine the role of PCV2 in causing apoptosis in gnotobiotic piglets. After separating 48 gnotobiotic piglets into three different groups and infecting them with PCV2a or PCV2b individually, the groups were further divided into clinically affected and non-affected piglets, according to displayed clinical signs. It was determined in this study that clinically affected piglets had a significantly higher amount of apoptotic cells compared to the clinically unaffected pigs and controls. We also established that a difference in the apoptotic rate in hepatic tissue between PCV2a and PCV2b was not present, and that both the cleaved Caspase-3 (CCasp3) immunohistochemistry (IHC) and TUNEL stain were equally reliable in the detection of apoptotic cells in hepatic tissue. The findings in the gnotobiotic pig model emphasizes that

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hepatic failure can be caused by the apoptosis of hepatic cells due to infection by PCV2. In a follow-up study, we further investigated if PRRSV has any effect on the PCV2 induction of apoptosis in lymphoid tissues in PRRSV- PCV2 co-infected pigs. Forty-four conventional pigs were divided into three groups and either remained uninfected or were co-infected with

PRRSV and PCV2 subtype a or b. We were not able to detect any difference between the groups including the control pigs, likely associated with the health status of the pigs, which did not display clinical signs.

PRRSV commonly replicates in porcine alveolar macrophages (PAMs) (Bautista et al., 1993; Mengeling et al., 1996), and since macrophages and dendritic cells are unable to degrade PCV2 (Gilpin et al., 2003; Vincent et al., 2003), there are concerns that this combination might lead to macrophage alterations. Such alterations might include uncontrolled release of cytokines, formation of oxygen radicals, nitric oxides, and proteases which have been previously reported with processes in PCV2 infection (Chang et al., 2006) which can directly alter the immune system. PCV2a and PCV2b differ in a signature motif located between nucleotides 1472 and 1486 of open reading frame 2 (ORF2), which corresponds to a six amino acid difference in the capsid protein (Cheung et al., 2007).

Previously, utilizing chimeric DNA clones between PCV2a and PCV2b isolates (i.e.

PCV2a/b and PCV2b/a) under both in-vitro and in-vivo conditions, it was found that PCV2 a/b and PCV2b/a clones had decreased virulence when compared to PCV2a/a and PCV2b/b respectively (Allemandou et al., 2011). In our next study, we designed an in vitro experiment where we included several PCV2-PRRSV combinations to establish a PCV2-

PRRSV in-vitro co-infection model and to further investigate the significance of ORF2 and the PCV2 signature motif in PCV2-PRRSV interaction. In-vitro differences in PCV2 or

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PRRSV replication or IFN-γ and IL-10 production in pulmonary alveolar macrophages inoculated with one of 30 PCV2-PRRSV combinations were small and independent of

PCV2 ORF2 origin, though PCV2a-ORF1 dependent in-vitro enhancement of PCV2 replication was observed.

Horizontal transmission of PCV2 via oral-nasal and oral-fecal routes have been considered to be the main mechanisms of PCV2 infection within and across pig herds

(Dupont et al., 2009; Segalés et al., 2005b). Numerous experimental studies and field observations have concluded that concurrent infection of pigs with PCV2 and various other pathogens increases the severity of PCVAD in affected pigs (Allan et al., 1999; Kennedy et al., 2000; Pallarés et al., 2002). Since PRRSV causes enhanced PCV2 replication (Allan et al., 2000; Allan et al., 2007), it is of concern that PRRSV might be able to induce elevated shedding of PCV2 in PRRSV-PCV2 co-infected pigs, which might lead to rapid transmission of PCV2 among herds. Since the influence of PRRSV on PCV2 transmission in growing pig is still unknown, we designed our next experiment to determine if PRRSV has any effect on the PCV2 shedding in PRRSV-PCV2 co-infected piglets. Our study established that PCV2 shedding in the co-infected groups were 1-3 logs higher than singularly infected groups when quantified by real-time PCR (Opriessnig et al., 2003), and shed for a longer period of time. This suggests that in field situations, co-infection with

PRRSV might enhance rapid transmission of PCV2 in herds. The present experiment also determined that there is no difference between PCV2a and PCV2b in the shedding of PCV2

DNA between singular or co-infected pigs. Our growing pig model provides evidence that

PRRSV-PCV2 co-infection in farms can be the source for rapid dissemination and infection

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in the herd, and emphasizes that monitoring PRRSV herd status and proper PCV2 vaccine strategies could help control PCVAD in herds.

PCV2 usually affects pigs in the early to late nursery phase (Dorr et al., 2007), and

PRRSV can be present in the newborns due to infected sows, therefore immune suppression in piglets might exist, leading to PCV2 vaccine failure as witnessed with classical swine fever vaccine failure (Suradhat et al., 2006). Mycoplasma hyopneumoniae vaccination times are usually adjusted to avoid vaccine failure due to the possibility of PRRSV immune- suppression (Drew, 2000), though the impact of PRRSV infection at the time of PCV2 vaccination is not yet known, and was the primary aim of our last study. After inoculating a portion of conventional 2-week-old piglets with PRRSV, commercially available PCV2 vaccines were administered at the anticipated peak of PRRSV viremia seven days later. The groups were challenged with PCV2b at 10 weeks of age. Our study established, for the first time, that PCV2 vaccines are very efficacious, even when administered to pigs with concurrent PRRSV viremia, as determined by significantly reduced viremia and PCV2 antigen in lymphoid tissues of all vaccinated groups. The results of this study indicate that

PRRSV viremia at the time of PCV2 vaccination does not affect PCV2 vaccine efficacy.

The research described in this dissertation has contributed to the collection of knowledge on the effect of PRRSV infection on PCV2 in co-infected growing pigs. Future work needs to be conducted to determine if apoptosis occurs in PRRSV-PCV2 co-infected clinically affected pigs to better understand the effect of apoptosis in pigs affected with

PCVAD. Further studies should investigate if PCV2 vaccines are equally efficacious when pigs are co-infected with PRRSV strains other than the VR2385 strain used in our studies, and if PCV2 vaccines are able to produce protective immunity in pigs simultaneously

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exposed to multiple strains of PRRSV, which is quite common in the field situations. The effect of PRRSV on PCV2 shedding and transmission should also be further investigated by utilizing different strains of PRRSV and also with dual infection of PCV2a and PCV2b to see if any recombination might occur to predispose the pigs to further elevated disease symptoms. More work needs to be undertaken to convert our in vitro study of cytokine release into an in vivo study to observe the cytokine release and cytokine mRNA expressions in the host when it is co-infected with different combinations of PRRSV and

PCV2 strains.

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ACKNOWLEDGMENTS

I would like to sincerely thank my committee members Dr. Tanja Oppriessing, Dr.

Cathy Miller, Dr. Rodney Butch Baker, Dr. Bradley Blitvich and Dr. Brett Sponseller for their direction, time, and assistance. I would like to give a special thanks to Dr. Tanja

Opriessnig for guidance, support, mentoring and teaching throughout this process. I would also like to thank Dr. Cathy Miller for teaching me and helping me from the very beginning.

I want to thank Joseph Brodie and others in the Veterinary Diagnostic Laboratory for aiding in histopathological work in my projects. I would also like to acknowledge and thank

Dr. Huigang Shen for assisting with PCR assays, Dr. Joao Carlos-Neto for helping me understand more about pathological lesions and Shayleen Schalk for assisting with project work and editing of my papers. I would like to thank the Laboratory Animal Resources staff who contributed in caring for the animals.

In addition, I would like to extend special thanks to Dr. Adam Bogdanove, Dr. Lisa

Nolan and Dr. Subhashinie Kariawasam for believing in me and helping me out whenever necessary.

I would specially like to thank my son Rahul for being patient and encouraging me during my exams and projects and eating food outside without complaint when the going got tough. I would like to thank my mother Mitra Sinha and father late Arun K Sinha for giving me the background education and help in every way possible to succeed in this long journey.

I want to thank my brother and his family to be there for me. I am grateful to you all.