**Atypical Porcine Pestivirus Circulation and Molecular Evolution within an A**ff**ected Swine Herd**

**Alba Folgueiras-González 1,2, Robin van den Braak 1, Bartjan Simmelink 1, Martin Deijs 2, Lia van der Hoek <sup>2</sup> and Ad de Groof 1,\***


Received: 27 May 2020; Accepted: 23 September 2020; Published: 25 September 2020

**Abstract:** Atypical porcine pestivirus (APPV) is a single-stranded RNA virus from the family Flaviviridae, which is linked to congenital tremor (CT) type A-II in newborn piglets. Here, we retrospectively investigated the molecular evolution of APPV on an affected herd between 2013 and 2019. Monitoring was done at regular intervals, and the same genotype of APPV was found during the entire study period, suggesting no introductions from outside the farm. The nucleotide substitutions over time did not show substantial amino acid variation in the structural glycoproteins. Furthermore, the evolution of the virus showed mainly purifying selection, and no positive selection. The limited pressure on the virus to change at immune-dominant regions suggested that the immune pressure at the farm might be low. In conclusion, farms can have circulation of APPV for years, and massive testing and removal of infected animals are not sufficient to clear the virus from affected farms.

**Keywords:** pestivirus; atypical porcine pestivirus (APPV); viral persistence; congenital tremor; swine; asymptomatic; genomic sequence; phylogenetic analysis; purifying selection

#### **1. Introduction**

The genus Pestivirus, belonging to the Flaviviridae family, includes single-stranded, positive-sense RNA viruses of veterinary importance, causing economically relevant diseases in livestock animals, but also affecting wildlife species [1]. While for decades the International Committee on Taxonomy of Viruses (ICTV) only recognized the four so-called classical pestiviruses, namely bovine viral diarrhea virus types 1 and 2 (BVDV-1 and BVDV-2), classical swine fever virus (CSFV) and border disease virus (BDV), the discovery of seven novel atypical pestiviruses in the last few years led to the proposal of a new species-independent classification from Pestivirus A to Pestivirus K. Thus, atypical porcine pestivirus (APPV) is now classified as Pestivirus K [2]. Moreover, new pestivirus genomes in bats [3] and piglets (LINDA virus [4]) have been discovered lately, but not yet classified.

Congenital tremor (CT) in newborn piglets, also known as myoclonia congenita, "shaker pigs" or "dancing pigs", was first reported in 1922 [5]. Its characteristic clinical signs involve tremors of the head and limbs which worsen on stress situations, but are almost gone during sleep. Even though most piglets are clinically healthy at weaning time, the earlier shivering hinders their ability to feed normally from their mother, thus increasing the risk for inadequate colostrum intake, growth retardation or even death by starvation [6]. CT is divided based on its pathology into type A, which displays morphological lesions on the central nervous system, and type B, which does not show any visible morphological lesion [7]. Regarding the distinct causes of disease, type A was further differentiated into five subtypes. Type A-I is associated with one of the classical pestiviruses, CSFV, and characterized by cerebellar hypoplasia in affected pigs [8]. Types A-III and A-IV are both related to different genetic defects only present in the Landrace breed and Saddleback breed, respectively [9,10]. Type-V, also defined by cerebellar hypoplasia, is caused by food poisoning by trichlorfon in pregnant sows [11]. Meanwhile, during the decades, the causing agent of CT type A-II remained unknown, although an infectious agent was suspected [12].

APPV was first characterized in 2015 by Hause et al. [13] through metagenomic sequencing performed on a sample positive for porcine reproductive and respiratory syndrome virus (PRRSV). After its discovery, two independent studies proved APPV's link to congenital tremor in newborn piglets via the inoculation of pregnant sows with infectious animal material [14,15]. However, Koch's postulates have not been fulfilled yet due to the difficulties in establishing an appropriate substrate for in vitro culture and retrieval of pure virus stocks [16]. In 2017, Lamb et al. [4] reported a novel pestivirus, named LINDA virus (lateral-shaking inducing neurodegenerative agent), in an Austrian cohort of CT piglets. Although the clinical manifestations and pathology were similar to the ones of APPV, further phylogenetic analysis showed its proximity to Bungowannah virus, a pestivirus never detected outside Australia.

A research study by Postel et al. [17] analyzed 1460 samples of apparently healthy pigs from different countries. They showed that the prevalence of APPV genotypes was around 10% and up to 60% of the tested animals were seropositive. Kaufmann et al. [18] showed that APPV has been circulating in Switzerland at least since 1986, which was the earliest detection point worldwide. To date, APPV has been reported on four continents and fourteen different countries [19]. Mortality is high in CT-affected litters, from around 10% to as high as 30% in some affected farms [14–16].

APPV is a highly variable enveloped virus which contains a linear, non-segmented, positive-sense RNA strand of around 11–12 kb. A single open-reading frame (ORF) encodes a single polyprotein of 3635 amino acids on average, which is assumed to be cleaved co- and post-translationally into four structural (the core protein C and the envelope glycoproteins Erns, E1 and E2) and eight non-structural proteins (Npro, p7, NS2, NS3, NS4A, NS4B, NS5A and NS5B) [13]. Several novel APPV sequences have been described since the first report in 2015, further dividing the phylogenetic tree and proposing distinct divisions into three, four or five different genotypes and into numerous particular subclades [20]. To date, 58 unique full-genome sequences are available in NCBI GenBank (access date: 1 April 2020). The APPV full-genome sequences have 80–99% identity, showing high variability even within the same country, with difficulty to infer the origin, dissemination and common ancestor of several strains of the virus [19,21].

High variability in RNA viruses, caused by the lack of proofreading repair mechanisms in RNA polymerases, provides them with a selective advantage, promoting rapid adaptation to novel hosts and environments [22]. On the other hand, excessive error rates can also lead to lethal mutagenesis and threaten the viability of the viral populations [23]. Therefore, hypervariable regions that are thought to contribute to immune escape and adaptation, as well as hyperconserved elements that are fundamental for virus replication and maintenance in the population are main research topics for molecular evolution studies [24].

In the current study, we retrospectively investigated the molecular evolution and functional implication of nucleotide and amino acid substitutions in the APPV genome in a farm in the North Brabant province of The Netherlands over a period of seven years. Sampling was done consistently over time in the controlled environment of a closed-herd farm, where the full farming process is carried out within the farm, and the breeding stock is replaced by gilts from the farm itself. Therefore, by eliminating the input of animals from outside, the risk of novel viral introductions is minimized. In 2013, a first CT outbreak with positive cases of APPV was reported in the farm. Two years later, in 2015, a second APPV outbreak occurred in the same farm with clinically affected, trembling piglets that tested positive. Since then, serum samples were taken from CT piglets at the time of the outbreaks, as well as from

apparently healthy pigs as part of regular farm monitoring for APPV and other porcine viruses. Pigs that tested positive were removed from the breeding population to stop the spread of the virus via persistently infected animals. However, several re-emerging peaks of viremic animals were been detected in the farm during the monitoring period, suggesting the continued circulation of APPV in clinically healthy animals.

To assess the possible biological significance of the selective pressures that shape the evolution of APPV within the farm, we looked for evidence of purifying and diversifying selection in the APPV genome. Although viral surface glycoproteins like E2 are well-known targets of positive selection, and variations in these regions can help in immune escape, we also took a full-genome study approach to get a more general, helicopter view and investigate other regions that might have been underestimated until now. The correlation of clinical CT outbreaks with fixation events of non-synonymous, but also synonymous, substitutions can reveal the link to pathogenicity and infectivity of the virus. Here, we reported, to our knowledge, the first longitudinal study on the evolution of the full-length APPV genome within a controlled farm environment. We compared molecular evolution and variability in the genome sequences found on the farm with full-genome sequences from different locations around the world.

#### **2. Materials and Methods**

#### *2.1. Sample Collection*

In 2013, the farm under study increased the population of breeding sows from 280 to 570 sows and relocated to new facilities. Piglets stayed in the farrowing unit after weaning (i.e., litters were not mixed). Piglets were vaccinated with a porcine reproductive and respiratory syndrome virus (PRRSV) live vaccine at 2 weeks of age, and with a combined porcine circovirus type 2 (PCV2) and *Mycoplasma hyopneumoniae* (*M. hyo*) vaccine at 3 weeks of age. PRRSV was regularly diagnosed in the farm and therefore a vaccination strategy in sows and piglets was set up. There were no other pathogens diagnosed in regular screenings.

Serum and fecal samples were obtained from gilts and congenitally trembling piglets during the CT outbreaks in 2013 and 2015. Between January 2016 and April 2016, serum was collected from all gilts selected for breeding (196 animals) at the age of first insemination for APPV monitoring and removal of positive animals.

To monitor the presence of APPV in serum over time, five litters with CT born from gilts during the 2015/2016 outbreak were repeatedly sampled until the age of 18 weeks. Moreover, 6 gilts (4 APPV-positive at 5 and 18 weeks of age, and 2 negative) and 2 boars (1 APPV-positive at 5 and 18 weeks of age, and 1 negative), were followed for APPV presence in serum and fecal shed until the age of 10 months. Two APPV-positive gilts were co-housed with one negative gilt, and the boars were housed next to each other with direct contact.

From April 2016 to 2019, clinically healthy 10-week-old gilts in the sow breeding line maintained on the farm were regularly monitored for the presence of APPV in serum. Samples from animals with congenital tremor symptoms were also taken for analysis. In total, 1498 samples were taken during this period.

Blood was collected using the Vacuette 5/8 mL Sep Clot Activator (Greiner-Bio One, Kremsmünster, Austria) and serum was obtained by centrifugation for 10 min at 3200× *g* at 4 ◦C. Fecal samples were collected using Sigma Virocult swabs and vials (MWE, Corsham, UK), vortexed, transferred to 1.5 mL Eppendorf tubes and centrifuged for 10 min at 10,000× *g* at 4 ◦C.

RNA was extracted from 200 μL samples by the automated MagNA Pure 96 system (Roche Applied Science, Manheim, Germany) using the protocol 'Viral NA plasma external lysis SV3.1 .

#### *2.2. Quantitative Reverse Transcription*

A universal, quantitative, reverse transcription PCR (qRT-PCR) was used to quantitatively detect APPV in samples using primers in the 5 -untranslated region (UTR) of the genome (APPV-PAN2-F3-B: CGYGCCCAAAGAGAAATCGG and APPV-PAN2-R3-B: CCGGCACTCTATCAAGCAGT) [14]. One-step qPCR reactions were performed in a final volume of 50 μL containing 1 μL SuperScript III RT/Platinum Taq Mix (Thermo Fisher Scientific, Waltham, MA, USA), 25 μL 2× SYBR Green Reaction Mix (Thermo Fisher Scientific, Waltham, MA, USA), 17 μL water, 1 μL forward primer (10 μM), 1 μL reverse primer (10 μM) and 5 μL of the RNA isolate. Thermocycling was performed in the CFX96 Touch real-time PCR detection system (Bio-Rad Laboratories, Hercules, CA, USA) starting with an RT reaction for 3 min at 55 ◦C, a pre-denaturation step for 5 min at 95 ◦C and 40 cycles of 15 s at 95 ◦C (denaturation) and 30 s at 60 ◦C (annealing and elongation). The specificity of SYBR Green qPCR was validated by melting curve analysis between 65 ◦C and 95 ◦C with an increasing gradient of 0.5 ◦C per 5 s. Results were analyzed with the CFX Manager software (Bio-Rad Laboratories, Hercules, CA, USA).

To obtain a standard curve for the quantitative analysis, a recombinant bacterial plasmid containing the 5 -UTR PCR target was made (GenScript, Piscataway, NJ, USA). The copy number of the recombinant plasmid was calculated, and eight dilution series (108–101 copies/μL) were included in duplicate in RT-qPCR to calculate the number of virus copies per μL.

#### *2.3. Viral Genome Amplification and Sanger Sequencing*

A starting sequence of 1073 bp obtained from a previous study using Illumina Sequencing (Amsterdam UMC, Amsterdam, The Netherlands) was used as reference [14]. This short read was mapped to the 58 full-genome sequences available in NCBI GenBank (access date: 1 April 2020). The three most similar full-genome sequences (GenBank accession numbers: KY624591.1, MH885413.1 and KX778724.1) were aligned by CLUSTALW using Geneious Prime v2019.0.4 (http://www.geneious.com) (Geneious, Auckland, New Zealand), and the consensus sequence was extracted for primer design.

The cDNA from the extracted RNA of the clinical samples (Section 2.1) was synthesized using the QuantiTect reverse transcription kit (Qiagen, Hilden, Germany) following the manufacturer's manual. The entire viral genome from the 2013 sample was amplified using an initial series of twelve overlapping PCRs. Gaps and faulty PCRs were solved following a genome walking strategy (amplification and sequencing primers available in Supplementary Table S1). PCR reactions were performed in a final volume of 25 μL containing 12.5 μL 2× Phusion High-Fidelity PCR Master Mix with HF Buffer (New England BioLabs, Ipswich, MA, USA), 0.75 μL DMSO 100%, 6.75 μL water, 1.25 μL FW 10 μM primer, 1.25 μL REV 10 μM primer and 2.5 μL of template cDNA. Thermocycling was performed in the CFX96 Touch real-time PCR detection system (Bio-Rad Laboratories, Hercules, CA, USA) using an initial denaturation step at 98 ◦C for 5 min, 40 cycles of 30 s at 98 ◦C (denaturation), 30 s at the annealing temperature optimal for the primer set and 1 min and 30 s at 72 ◦C (elongation), followed by 7 min at 72 ◦C. The PCR products were stored at 4 ◦C until further analysis. Five microliters of the PCR products were analyzed by agarose gel electrophoresis (1.4% *w*/*v* agarose) to check the fragment size and the specificity of the amplification. PCR products were purified using the QiaQuick PCR purification kit (Qiagen, Hilden, Germany) following the manufacturer's manual.

Sequencing PCR reactions were performed using the BigDye Terminator v3.1 cycle sequencing kit (Applied Biosystems, Carlsbad, CA, USA) in a final volume of 20 μL containing 4 μL BigDye Terminator ready reaction mix, 3 μL 2× Phusion High-Fidelity PCR Master Mix with HF Buffer (New England BioLabs, Ipswich, MA, USA), 0.5 μL DMSO 100%, 2.5 μL WFI, 2.5 μL FW or REV 10 μM primer and 7.5 μL of the purified PCR product. Thermocycling was performed in 30 cycles of 10 s at 95 ◦C (denaturation), 10 s at the annealing temperature optimal for the primer set and 2 min at 60 ◦C (elongation). The PCR products were stored at 4 ◦C for further analysis. The cycle sequencing products were purified using the DyeEx 2.0 Spin kit (Qiagen, Hilden, Germany) following the manufacturer's manual. Capillary electrophoresis was performed using the 3500 Genetic Analyzer (Applied Biosystems, Carlsbad, CA, USA) and data were analyzed using the Sequencher 5.4.6 (Gene Codes, Ann Arbor, MI, USA) and Geneious Prime v2019.0.4 (Biomatters Ltd., Auckland, New Zealand) software.

Based on the obtained sequence, Sanger sequencing was performed following the same methodology on the other five samples (years 2015, 2016, 2017, 2018 and 2019) by GenScript (Piscataway, NJ, USA).

#### *2.4. Determination of E2 Sequences*

The amplification of E2-coding sequences was performed using a two-step RT-PCR protocol as described in Section 2.3. The amplification and sequencing PCR reactions of 857 bp and 953 bp fragments were performed using two primer pairs flanking the E2-coding region (E2-F1: 5 -TGGTGCCTATTGTTGTCAGG-3 , E2-R1: 5 -AGTTCTTCCTTGACGGCTAG-3 , E2-F2: 5 -GCCCTGGTGAACATAGTCAC-3 and E2-R2: 5 -TCCTTGACGGCTAGCATTATG-3 ). Capillary electrophoresis was performed using the 3500 Genetic Analyzer (Applied Biosystems, Carlsbad, CA, USA) and trimmed sequences (702 bp) were analyzed using Geneious Prime v2019.0.4 (Biomatters Ltd., Auckland, New Zealand).

#### *2.5. Submission of Sequences*

The sequences of the APPV genomes were deposited in GenBank under the accession numbers MT512531–MT512537. The partial E2-coding sequences were deposited in GenBank under the accession numbers MW011356–MW011406.

#### *2.6. Nucleotide and Amino Acid Analysis of Variants*

The obtained partial E2-coding sequences were aligned using the CLUSTALW translation alignment in Geneious Prime v2019.0.4 (Biomatters Ltd., Auckland, New Zealand) with BLOSUM cost matrix, a gap open cost of 5 and a gap extended cost of 4.

The fifty-eight full-genome APPV sequences from NCBI GenBank database (access date: 1 April 2020) were aligned along with the sequences obtained in the current study [25]. Multiple sequence alignment was performed using the CLUSTALW translation alignment in Geneious Prime v2019.0.4 (Biomatters Ltd., Auckland, New Zealand) with BLOSUM cost matrix, a gap open cost of 10 and a gap extended cost of 6.66. For the farm dataset, we called single nucleotide substitutions on the aligned genomes using Geneious Prime v2019.0.4 and differentiated them into synonymous and non-synonymous. Ambiguous nucleotide and amino acid calls, which are considered by the software as variants, were treated as missing data.

The prediction of *O*-glycosylation and *N*-glycosylation motifs in the Erns, E1 and E2 glycoproteins were performed with DictyOGlyc1.1, NetOGlyc4.0 and NetNGlyc1.0 prediction algorithms via the webserver from the Technical University of Denmark Department of Bio and Health Informatics (DTU Bioinformatics, http://www.cbs.dtu.dk/services/) [26,27].

#### *2.7. Phylogenetic Analysis and Estimation of Evolutionary Rates within the Farm*

The Recombination Detection Program version 4 (RDP4) was used to screen for recombination events on the multiple sequence alignment of the six APPV genomes found in the farm, using RDP, GENECONV, BootScan, Maxchi, Chimaera, Siscan and 3Seq methods. Recombination was considered when *p*-value was <0.0001 and the recombinant score was >0.6 [28].

The full-genome APPV sequences available in NCBI and the six APPV in-farm sequences obtained in this study were aligned using the CLUSTALW translation alignment in Geneious Prime v2019.0.4 (Biomatters Ltd., Auckland, New Zealand) with BLOSUM cost matrix, a gap open cost of 10 and a gap extended cost of 6.66. The phylogenetic tree was created with the MEGA X software using the neighbor-joining method with the Kimura two-parameter substitution model [29,30]. Complete deletion was done in case of gaps or missing data. The analysis was performed for 500 bootstrap replicates. An estimation of the evolutionary rates was calculated for the obtained E2-coding sequences, as well as for the six full-genome sequences using the TempEst v1.5.3 software [31].

#### *2.8. Selection Pressure Analysis*

Selection pressure analyses were done for the multiple sequence alignment of the six farm sequences as well as for the multiple sequence alignment including the NCBI full-genome APPV sequences. Selection pressure analyses were performed on the full ORF, using mixed-effects model of evolution (MEME) and fixed-effects likelihood (FEL). All these algorithms were implemented on the HyPhy (Hypothesis Testing using Phylogenies) open-source software package and can be accessed through the Datamonkey webserver (https://www.datamonkey.org/) [32].

MEME uses a mixed-effect maximum likelihood approach. The algorithm estimates a synonymous α parameter (dS) and a two-category mixture of non-synonymous (dN) β parameters for each site. MEME infers two ω (dN/dS) classes and uses a likelihood ratio test to compare between the models and check for episodic diversifying selection. A significance threshold of *p* < 0.1 was used on the analysis [33].

The fixed-effects likelihood (FEL) algorithm was used in order to detect negatively selected sites on the alignment. FEL uses a maximum likelihood approach to infer non-synonymous (dN) and synonymous (dS) substitution rates, also assuming a constant selection pressure for each site on the alignment. All branches were tested for selection. A model with synonymous rate variation, where the dS parameter in the codon model is allowed to vary across sites, was used for the analysis. A significance threshold of *p* < 0.1 was used on the analysis [24].

#### **3. Results**

#### *3.1. 2013–2016: Congenital Tremor Outbreaks, Follow Up Studies and Eradication Strategy Design*

In 2013, the farm increased the internal population of breeding sows, aimed to breed gilts for the replacement of production sows (sows breeding line) from 280 to 570 sows and, at the same time, relocated to new facilities with state-of-the-art climate control and housing conditions. Piglets stayed in the farrowing unit after weaning, thus litters were not mixed. In the same year, a first large-scale CT outbreak was reported on the farm and a second large-scale outbreak occurred at the end of 2015/early 2016. Both outbreaks were related with positive cases of APPV and various other clinical observations (e.g., reduced vitality and mortality of the piglets, and increased return to estrus percentage). In the time between 2013 and 2015, no major abnormalities were observed, although incidentally few piglets with tremors were seen, but with no impact on production.

During the 2015–2016 outbreak, CT prevalence varied from 5% in litters from fourth parity sows, up to 55% in litters from first and second parity sows. In the latter case, the mortality in the farrowing unit ranged from 25% in litters from second parity sows to 69% in the ones from first parity sows. The related effects on the production data are shown in Supplementary Table S2.

After the second large-scale outbreak, it was decided to monitor all gilts selected for breeding for the presence of APPV, both purebred line gilts and production gilts. Serum of those animals was analyzed at the age of first insemination with the aim to remove positive gilts from the breeding population. A universal, quantitative, reverse transcription PCR (qRT-PCR) was used to quantitatively detect APPV on pig serum samples. This strategy was applied between January and April 2016, during which time 196 gilts were tested and 15% of them tested positive for APPV (Supplementary Table S3).

During parallel monitoring of pigs born with CT and positive for APPV in serum, we observed that a significant percentage of pigs turned PCR-negative for APPV in serum around the age of 18 weeks. The piglets born with CT were considered as persistent carriers of the virus and, at the time of weaning, the virus was still present in the piglets that still showed recognizable, but less severe, tremors. A follow-up of these piglets showed that the percentage of APPV-positive PCR scores in

serum was reduced to 45% at 18 weeks of age, with viral copy numbers in serum also being greatly reduced (Supplementary Table S4).

To gain further insight into the dynamics of APPV viremia, 6 gilts (4 APPV-positive at 5 and 18 weeks of age, and 2 negative), and 2 boars (1 APPV-positive at 5 and 18 weeks of age, and 1 negative), were followed for APPV presence in serum until the age of 10 months. Two APPV-positive gilts were co-housed with one negative gilt, in separate cages, and the boars were housed next to each other with direct contact. No APPV was detected in any of the 8 animals at the age of 24 weeks and at any time point thereafter, with the exception of 1 boar testing PCR-positive at the age of 32 weeks (Supplementary Table S5). Fecal shed from the same animals was also monitored until the age of 10 months via qPCR. Results showed, in the age range between 24 and 44 weeks, intermittent shedding in the feces of the persistently infected (PI) gilts and temporary presence in the feces of horizontally infected gilts (Supplementary Table S6).

#### *3.2. 2016–2019: Re-Emergence of Congenital Tremor and Monthly Quantitative Detection of APPV*

Since April 2016, when regular sampling from clinically healthy 10-week-old gilts in the sow breeding line maintained on the farm started, serum samples were received monthly for APPV monitoring purposes. Until December 2019, a total of 1505 serum samples from 10–16-week-old pigs were analyzed.

Figure 1 shows the percentage of APPV-positive animals among the tested set for each month since the screening started in April 2016 until the last analyzed data from December 2019. In this period, two peaks of CT symptoms in newborn piglets were detected in the farm in April 2016 and May 2017 (blue columns in Figure 1). In order to diminish horizontal transmission of the virus in the sow breeding population, infected APPV pigs detected during the monitoring were removed from the population as these were likely persistently infected.

**Figure 1.** Percentage of atypical porcine pestivirus (APPV)-positive pigs detected during monthly screening in the farm by RT-qPCR analysis. The *x*-axis shows the sampling date from April 2016 to December 2019. The percentage of positive samples is shown in the *y*-axis. Blue-colored columns show that positive PCR results were accompanied by clinical CT symptoms on those specific months—i.e., April 2016 and May 2017. The number of PCR-positive animals in relation to the total number tested each specific month is shown above the columns.

#### *3.3. Characterization of APPV Sequences*

Forty-eight E2-coding sequences were determined from the APPV-positive samples collected during monthly monitoring in the farm at different time points between 2016 and 2019, along with three sequences obtained previously in 2013 and 2015 during the large-scale CT outbreaks (Supplementary Table S7). The E2-coding sequences showed a maximum pairwise genetic distance of only 0.71%. The low genetic diversity observed within the farm supported the hypothesis that only one viral strain was circulating in the herd with no viral introductions from the outside.

The amino acid sequences were also compared according to the collection date (Supplementary Figure S1). Samples obtained from different animals at the same time point showed a completely identical genome, with the exception of the highly mutated codon D752, varying to either Gly, Ser or Asn residues (Section 3.5). Besides, a more thorough approach was taken for the samples obtained from affected animals at the time of the CT outbreaks, with 8 out of 12 sequences obtained in April 2016 and 8 out of 9 in May 2017. No characteristic differences were seen at those time points that could lead to an explanation of the distinct symptoms in relation to the nucleotide or amino acid sequences.

#### *3.4. Phylogenetic Analysis and Estimation of Evolutionary Rates of APPV within the Farm*

A phylogenetic analysis of the full-length APPV sequences available worldwide and obtained from the NCBI database and six APPV sequences obtained in the present study (Section 3.5) confirmed the division of APPV into three different clades, two of them containing only sequences from China (Figure 2). APPV sequences from the current study clustered in one subtree together with two German sequences (NC\_030653 and KU041639, from 2015), one Austrian sequence (KX778724, from 2016), two Chinese sequences (MH885413, from 2018, and KY624591, from 2016) and one South Korean sequence (MF979135, from 2016). The other full-APPV genome from The Netherlands available in NCBI was clustered within the same clade, but into a different subtree, together with eight sequences from Switzerland and two from the US (Figure 2).

**Figure 2.** Phylogenetic analysis of the complete coding region alignment of atypical porcine pestivirus (APPV) sequences worldwide obtained from NCBI database and six APPV sequences retrieved in the present longitudinal study on the Dutch swine farm between 2013 and 2019. The neighbor-joining method with the Kimura two-parameter substitution model was used. Gaps and missing data were

subjected to complete deletion. Bootstrap values are provided at the root of the clusters. The scale bar is a measure of the proportion of divergence. Run for 500 bootstrap replicates. Clade I shown as expanded tree. Clades II and III, containing only sequences from China, are collapsed and shown in green and purple in the lower part of the tree. APPV branches obtained in the current study are shown in pink.

#### *3.5. Genome and Protein Variations of APPV within the Farm*

The samples with the highest numbers of APPV genome copies were selected from each of the years, and six full-length APPV genomic sequences from samples collected in the farm on 2013 and each year between 2015 and 2019, were determined. The 2013 and 2015 APPV genome sequences were obtained from serum samples from CT-affected piglets at the time of the large-scale outbreaks. The length of the complete APPV in-farm coding sequence (excluding the 5 -UTR region) was 10,908 nt, in line with the previously reported length of the coding region [14]. Multiple sequence alignment was performed on these six sequences using the CLUSTALW translation alignment in Geneious Prime v2019.0.4. The sequences from the different years showed a high similarity at the nucleotide level, between 99.74% and 99.90%. The number of nucleotide differences ranged from only 10 nucleotide substitutions between 2017 and 2019 genomes to 26 nucleotide substitutions between 2013 and 2017 genomes. On the amino acid level, the differences ranged from only 1 amino acid substitution between 2017 and 2019 to 7 amino acid substitutions between 2013 and 2017 and between 2016 and 2017 (Table 1).


**Table 1.** Distance matrix from the multiple sequence alignment of six in-farm APPV sequences.

The nucleotide substitutions found in the alignment were classified regarding their protein location and differentiating synonymous substitutions and non-synonymous substitutions. All substitutions are shown in Supplementary Table S8. Forty-five nucleotide substitutions were found along the APPV genome during the monitoring years in the farm from which thirty-four positions corresponded to synonymous substitutions. Eleven non-synonymous substitutions occurred on the APPV genome in the farm between 2013 and 2019. Among those, one amino acid change, V563A, was in the structural glycoprotein gene E1, while two amino acid changes, I726V and D752G, were in the E2 structural glycoprotein encoding region. The other eight non-synonymous substitutions occurred in the genes of non-structural proteins: two in Npro (S48P and H152L), one in NS2 (R1122K), three in NS5A (K2437R, G2550E and A2824T) and two in NS5B (D2979N and N3166T) (Supplementary Table S8).

Some amino acid changes in the strains have not been described before. The E1 non-synonymous substitution V563A only occurred within the farm, while the genotype of this site worldwide was always conserved as a Val residue. The synonymous substitution found on the same coding gene, V567, was also exclusive, when compared with other sequences in Clade I. Interestingly, site T608 in the in-farm sequences possessed a unique genotype compared with the consensus I608, and the V608 variant present as well in Clades II and III. Moreover, site I790 also showed a unique genotype, found only in one sequence from Switzerland from 2011 (accession number MN099165), while a Val residue is encoded on that position in the sequences worldwide. The glycosylation status of the pestivirus glycoproteins plays an important role in virulence. In this line, the substitution on E2 glycoprotein D752 to a Gly, Ser or Asn residue (Supplementary Table S8, Supplementary Figure S1) modified the 752NDT754 N-glycosylation motif predicted by NetNGlyc1.0 server (http://www.cbs.dtu.dk/services/). In the non-structural proteins, a unique motif was found in the N-terminal protease. The site 126KPAPASR132 was unique within the APPV sequences retrieved from the current farm study, including up to four modifications on its amino acid chain, in comparison with other geographically distinct APPV full genome sequences.

An estimation of the evolutionary rates within the farm was calculated for the full ORF of the APPV genome, as well as for the E2 glycoprotein-the coding sequence using TempEst v1.5.3 software [31]. An evolutionary rate of 3.224 <sup>×</sup> 10−<sup>4</sup> substitutions/site/year (correlation coefficient <sup>=</sup> 0.9009; *R*<sup>2</sup> = 0.8117) was estimated for the full ORF of APPV over a period of six years of evolution, from 2013 to 2019, within the studied farm. The envelope glycoprotein E2, considered in all pestiviruses as the main antigen eliciting immune response, had an estimated evolutionary rate of 9.347 <sup>×</sup> 10−<sup>4</sup> substitutions/site/year (correlation coefficient = 0.7706; *R*<sup>2</sup> = 0.5938). Both evolutionary rates were in line with the ones reported in other pestivirus species. The evolution of the CSFV E2 gene was estimated between 1.73 <sup>×</sup> <sup>10</sup>−<sup>3</sup> and 5.76 <sup>×</sup> 10−<sup>4</sup> substitutions/site/year [34,35], while the full genome estimation was 1.03 <sup>×</sup> 10−<sup>4</sup> substitutions/site/year [36]. For BVDV, evolutionary rates of 1.40 <sup>×</sup> 10−<sup>4</sup> substitutions/site/year for the full genome and 1.26 <sup>×</sup> <sup>10</sup>−<sup>3</sup> substitutions/site/year for the E1-E2 coding region were reported [37]. A recombination analysis was done using the RDP4 software in order to find breakpoints within the multiple sequence alignment that might have undergone recombination, and to screen them for evidence of phylogenetic incongruence previous to phylogenetic tree building [28]. No recombination events were found at the farm level.

#### *3.6. Selection Patterns within a Farm: Six-Year Evolution Study on an APPV in-Farm Variant*

Negative, also called purifying, selection consists of the evolutionary pressure hindering the fixation of non-beneficial or deleterious protein variations in the population. On the other hand, positive or diversifying selection is the evolutionary pressure that promotes the fixation of beneficial protein variations in the population. This last one can be considered pervasive, when it is constant through time, or episodic, when it is sporadic and affecting only some lineages. Using the algorithms implemented in the HyPhy software packages via the Datamonkey webserver [32], we were able to identify a number of codons potentially subjected to selection pressures on our APPV genome between 2013 and 2019.

No positively selected codons, neither episodic nor pervasive, were found by any of the methods. Thus, positive selection cannot be considered as an evolution pattern within the farm during the six years of the study. On the other hand, FEL (fixed-effects likelihood) found evidence of pervasive negative/purifying selection at 9 sites among the 3635 codons, with a *p*-value threshold of 0.1. The nine codons subjected to purifying selection were found to be in the N-terminal protease (codon 132), the Erns glycoprotein (codon 418), the NS2 protein (codons 1054 and 1198), the NS3 protein (codon 1814), the NS5A protein (codons 2676 and 2848) and the NS5B protein (codons 2905 and 3243). FEL algorithm indicated, under the global MG94xREV model, a total non-synonymous/synonymous rate ratio for the in-farm alignment equal to 0.172, indicating the prevalence of negative selection within the APPV genome sequence.

#### **4. Discussion**

In 2013, a first large-scale CT outbreak was reported in a farm in The Netherlands, with a second large-scale outbreak at the end of 2015/early 2016, both related to APPV infections. After the second outbreak, all gilts selected for breeding were monitored by serum PCR screening at the age of first insemination for the presence of APPV with the aim of removing infected animals from the breeding population. However, there were some practical limitations to this approach: (1) The removal of a high percentage of the selected gilts at the age of first insemination severely disrupted the normal farm sow replacement strategy. (2) Serological tests were not available at the time and thus, the difference between persistent carriers and gilts with horizontal infections could not be established. (3) Parallel monitoring of persistently infected pigs from the second outbreak revealed that, in contrast to what it may have been expected based on experience with other pestiviruses, APPV disappeared from the serum in adult pigs. The data published by Schwartz et al. [16] showed stable presence of APPV in serum until 14 weeks of age, but our observations on a larger number of animals showed that a significant percentage of pigs turned PCR-negative for APPV around the age of 18 weeks.

Fecal shed was also monitored to potentially identify persistently APPV-infected pigs. In the age range between 24 and 44 weeks, monitoring APPV infection is difficult due to the absence of the virus in serum and intermittent shedding in the feces of PI gilts. Horizontally infected gilts show temporary presence in feces and serum. Therefore, a gilt-monitoring strategy was technically complicated and could lead to false-negative test results and consequently, the maintenance of persistently infected animals in the breeding population. It is of note that APPV-positive fecal shed has not proven to be infectious.

Based on these considerations, we reasoned that the early removal of PI animals in the purebred gilt breeding population, by performing qPCR analysis on serum samples before the age of 16 weeks, was the best strategy to control APPV at the farm level. Hence, a monthly monitoring program for APPV infections in 10-week-old selected breeding gilts of the sow breeding line was set up in the "closed" CT-affected farm with the purpose of reducing the risk of horizontal APPV infections to pregnant gilts and subsequent vertical infection of newborn piglets. It was assumed that these persistent carriers of APPV, often born without showing clinical symptoms after vertical transmission of the virus and with subsequent use in the farm sow replacement breeding program, contributed significantly to the risk of infections of pregnant gilts and low-parity sows in group housing. The results of our analysis showed that even with stringent removal of APPV PCR-positive, assumed persistently infected replacement gilts, the virus cannot be completely eliminated from the farm. The genotype remained unchanged throughout the years, as demonstrated by the low genetic diversity observed in the partial E2-coding sequences from the samples obtained during monthly monitoring, making it unlikely that new introductions via sperm were involved in the continuing APPV infections in the farm. Re-emerging peaks of viremic animals, often without symptoms of the disease but incidentally with the birth of CT piglets during these six years within this closed-herd farm, even with infected breeding animals removed from the population, showed that the virus was difficult to control let alone eliminate. Besides, the E2-coding sequences obtained from the majority of affected animals at the time of the CT outbreaks did not show any characteristic difference at the nucleotide nor amino acid level when compared to the asymptomatic carriers that could lead to an explanation for the clinical manifestation.

The hypervariable regions, which contribute to virus escape from the immune system, as well as the conserved elements involved in crucial virus replication processes and maintenance of the viral population, are obvious targets for drug and vaccine design [24]. Moreover, it can be hypothesized that certain substitutions in the viral coding sequence may be involved in the cyclic peaks of viremia and/or correlate with the pathogenicity of the virus. In the present study, forty-five nucleotide substitutions were found along the APPV genome during the monitoring years in the farm, eleven of them causing an amino acid change. Among those, two occurred on the E2 protein-coding gene, usually recognized as the main antigen able to elicit neutralizing antibodies in infected animals [38,39]. Recent studies on subunit E2 vaccines from APPV have shown strong immune responses in mice [40]. Moreover, E1–E2 heterodimers have been considered in other pestivirus species as key for viral infectivity [38]. Although the results obtained after the analysis of the APPV sequences on the followed farm did not point at any positively selected sites, two of the non-synonymous substitutions found on the heterodimer-coding genes were almost unique to the farm under study. More interestingly, the substitution D752G in E2 glycoprotein, which showed high variability throughout the fifty-one partial E2 sequences analyzed, modified the predicted 752NDT754 *N*-glycosylation motif. Generally, E2 glycosylation sites are highly conserved due to their primary role

in viral entry and infection and their removal has shown viral attenuation in other pestivirus species [41]. Further research needs to be done to decipher the functionality of this highly variable motif.

The unique motif 126KPAPASR132 was found in the N-terminal protease in all APPV sequences retrieved from the farm. This motif included up to four modifications on its amino acid chain, in comparison with other geographically distinct APPV full-genome sequences. Although the exact functionality of this site is not known, Npro plays an important role in viral evasion of the innate immune response in other pestivirus species. Npro activity decreases the levels of the interferon regulatory factor 3 (IRF3) via proteasomal degradation, inhibiting its downstream signaling and thus suppressing type-I interferon responses in infected animals [42]. Previous research studies have related the functionality of this protein with the enhancement of co-infections with bovine respiratory syncytial virus (BRSV) in cattle [43]. APPV in persistently infected animals is commonly found together with other viruses and, it may therefore intensify those secondary infections. More importantly, experimental Erns and Npro mutations in the BVDV genome have failed to induce persistent infections in cattle [44].

The other 34 substitutions found along the APPV genome during the six years evolution in the farm, but not leading to amino acid changes, are not less important as the codon usage bias has been related to virus translation efficiency [45], RNA structures critical for replication and packing [46] and enhanced virulence [47]. Future studies on the codon usage and effects on RNA structure using reverse genetics may therefore shed light on the actual effect of synonymous substitutions. We are aware of the fact that, with only six full-genome sequences included in the current study, our results were based on a limited amount of sequence data. However, given the dynamics of the virus at the farm, we had maximized our analysis possibilities. Pigs with high APPV loads in serum, which are needed for a successful sequencing strategy, were only occasionally observed and, in addition, pigs from the same litter were expected to have the same viral genotype, as demonstrated by the partial E2 sequences obtained from a large number of samples, and further sequence analysis would not add meaningful data. Moreover, the limited genome variation shown within the farm suggested that more sequences obtained during the same years would not radically change the observations and conclusions drawn in the present study. Nonetheless, we acknowledge that an increase in the number of APPV full-length sequences available, especially within longitudinal studies, will be key for the scientific community to further understand the evolutionary dynamics and genomic features of this pestivirus.

With regard to the evolutionary pressures of APPV, the present study showed a general genome-wide purifying pressure, especially strong on the non-structural proteins. These results indicated the importance for the virus to maintain the functionality of the non-structural viral proteins, avoiding the fixation of detrimental amino acid substitutions that might hinder the virus ability to evade the host immune system and cause persistent infections. In Flaviviridae, non-structural proteins have been determined as key players in viral escape from the host immune system. Hepatitis C virus (HCV) NS3 protein inhibits tumor necrosis factor alpha (TFN-α) stimulated NF-κB activation to evade the host innate immunity [48]; NS4A proteins of dengue virus 1 (DENV-1) inhibit the interferon- β (IFN-β) signaling pathway mediated by the retinoic acid-inducible gene I (RIG-I) and TANK-binding kinase-1 (TBK1) proteins [49]; and NS5 protein of several flaviviruses is also involved in the inhibition of IFN signaling by degrading the signal transducer and activator of transcription 1 (STAT1) or STAT2 [50,51]. Even though the signaling pathways and target sites of the majority of the non-structural proteins remain unknown for several virus species, these examples acknowledge their potential function in the host innate immune evasion. The strong purifying selection we found could be explained by a lack of pressure from the host immune system, as the persistently infected animals may potentially be immunotolerant, although experimental evidence is needed to ultimately confirm this hypothesis. Regular use of ELISA systems, to monitor whether immunity is raised to the virus, would bring light to this issue [39].

Monitoring studies in APPV-infected farms during the last years showed that APPV caused persistent infections in piglets when they were infected in-utero, occasionally developing congenital tremors, while horizontal infections within the herd were transient and without visible clinical signs [39]. As aforementioned, this is similar to BVDV infections in cattle, which only causes persistent infections in the calves when the virus is transmitted in-utero before 120 days of gestation. While some persistently infected calves are born with congenital malformations, others are clinically healthy with BVDV infections going unnoticed until the onset of clinical signs in the new parities [52]. A widely accepted hypothesis, supported by several research studies, is that non-cytopathic BVDV strains fail to induce type-I interferon responses in the infected fetus developing persistent infections and immunotolerant calves [52,53]. These persistently infected animals shed the virus continuously during their lifetimes, provoking transient horizontal infections within the herds, but more importantly infecting pregnant cows leading to the birth of persistently infected calves [54].

Our study results of APPV circulation on the affected sow farm showed similarities with the ways that BVDV persists within the cattle populations. Persistently APPV-infected animals remained undetected in a population that was not continuously monitored by PCR because they did not always show congenital tremors. Although our control strategy did not result in full elimination of the virus, at least it resulted in no further large-scale outbreaks since early 2016. Our approach was based on the hypothesis that persistently infected gilts are the main contributors to the spread of the virus and thus, removing those indirectly leads to the prevention of infections of pregnant sows. Nevertheless, the combination with a direct approach based on the elimination of infected pregnant animals could further improve the eradication strategy, as experimental studies have shown that virus infection during early stages of gestation leads to viremia and transplacental transfer.

Research on APPV vaccines is still limited. High strain heterogenicity and persistent infections have been a major bottleneck in the case of BVDV vaccination [55], and for APPV the same problems will be encountered when vaccines are designed and tested. Importantly, effective controlling strategies in closed farm situations should start with intensive testing of all the animals in the herd, followed by removal of positive carriers from the population. Testing should not only be done via RT-qPCR in serum, but also in fecal swabs, as it has been observed that the virus disappears from blood while it can still (intermittently) be present in feces [14]. The strategy presented in this study was not adequate to fully eliminate the virus; APPV kept circulating in the sow population after removal of persistent carriers in the sow breeding herd, suggesting that all gilts should be monitored. In addition, serological data could reveal if the virus is still circulating within the herd via horizontal infections. Vaccination of seronegative gilts and sows is an option that can be considered if removal of carriers alone does not sufficiently control APPV infections.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/1999-4915/12/10/1080/s1, Table S1: List of amplification and sequencing primers used in the study; Table S2: Details on the production data of the studied farm during the 2015–2016 CT outbreak; Table S3: Serum APPV monitoring of gilts selected for breeding at the age of first insemination between January and April in 2016; Table S4: Follow-up of five litters with CT born from gilts for the presence of APPV in serum over time (2015/2016 outbreak); Table S5: Follow-up of boars and gilts until the age of 10 months for the presence of APPV in serum (2015/2016 outbreak); Table S6: Follow-up of boars and gilts until the age of 10 months for the presence of APPV in fecal shed (2015/2016 outbreak); Table S7: APPV E2 sequencing sample list and corresponding APPV viral loads; Table S8: Nucleotide substitutions in the APPV in-farm multiple sequence alignment; Figure S1: Multiple sequence alignment of the partial APPV E2 amino acid sequences chronologically ordered.

**Author Contributions:** Conceptualization, A.F.-G., A.d.G. and L.v.d.H.; methodology, A.F.-G., L.v.d.H., M.D., R.v.d.B. and B.S.; software, A.F.-G., R.v.d.B. and B.S.; data analysis, A.F.-G.; writing—original draft preparation, A.F.-G.; writing—review and editing, A.d.G., L.v.d.H. and R.v.d.B.; supervision, A.d.G. and L.v.d.H. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received funding from the European Union's Horizon 2020 research and innovation program, under the Marie Skłodowska-Curie Actions grant agreement no. 721367 (HONOURs).

**Acknowledgments:** The authors wish to acknowledge the input of the veterinarian Jolanda Rooijendijk, from DAC de Peelhorst, and the farmers who sent in samples of diseased animals. We thank Ignacio Postigo-Hidalgo for useful discussions on bioinformatics.

**Conflicts of Interest:** The authors of this manuscript have the following competing interests: A.F.G., R.v.d.B., B.S. and A.d.G. are employed at MSD Animal Health, a commercial company. M.D., A.d.G. and L.v.d.H. are inventors on a patent application on APPV.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **The Outcome of Porcine Foetal Infection with Bungowannah Virus Is Dependent on the Stage of Gestation at Which Infection Occurs. Part 1: Serology and Virology**

#### **Deborah S. Finlaison \* and Peter D. Kirkland**

Virology Laboratory, Elizabeth Macarthur Agricultural Institute, New South Wales Department of Primary Industries, Menangle, NSW 2568, Australia; peter.kirkland@dpi.nsw.gov.au

**\*** Correspondence: deborah.finlaison@dpi.nsw.gov.au

Received: 29 May 2020; Accepted: 24 June 2020; Published: 26 June 2020

**Abstract:** Bungowannah virus is a novel porcine pestivirus identified in a disease outbreak in Australia in 2003. The aim of this study was to determine the outcome of infection of the pregnant pig with this virus. Twenty-four pregnant pigs were infected at days 35, 55, 75 or 90 of gestation. Blood, tonsillar and rectal swabs were collected from each pig at birth and then weekly until euthanasia or death. Tissues were sampled at necropsy. Viral load was measured by real-time reverse-transcription polymerase chain reaction (qRT-PCR) and antibody levels in serum by peroxidase-linked immunoassay. Bungowannah virus was detected in the serum and excretions of all infected pigs at birth regardless of the stage of gestation at which infection occurred. Persistent infections occurred following infection prior to the development of foetal immunocompetence. Unexpectedly some animals infected at day 55 of gestation later cleared the virus and seroconverted. Viraemia and viral shedding resolved quickest following infection in late gestation.

**Keywords:** Bungowannah virus; foetus; pestivirus; porcine; real-time PCR; serology; virology

#### **1. Introduction**

Bungowannah virus is a novel pestivirus identified from an outbreak of disease in a piggery in New South Wales, Australia, in June 2003 [1]. It is genetically distinct from the other recognised pestiviruses of pigs, classical swine fever virus (CSFV) and atypical porcine pestivirus (APPV) [2,3] with its closest genetic relationship to the recently identified Linda virus [4]. The disease was referred to as the porcine myocarditis syndrome, or PMC, because histological changes in affected animals consist almost exclusively of a multifocal non-suppurative myocarditis, with myonecrosis in some cases. The outbreak initially presented as sudden death in 2- to 3-week-old weaning age pigs, but soon after the onset there was a marked increase in the birth of stillborn foetuses and a slight increase in the occurrence of mummified pigs. Cumulative losses in some weeks exceeded 50% of pigs born, and it is estimated that as many as 50,000 pigs died in the initial outbreak. Due to the reproductive effects and disease occurring almost exclusively in the first 2–3 weeks of life it was presumed to be predominantly the consequence of in utero infection. This hypothesis was supported by the detection of elevated serum IgG levels in up to 50% of stillborn pigs and by the absence of disease in pigs soon after weaning or in sows farrowing affected litters [1].

The pestiviruses are well recognised reproductive pathogens where the outcome of infection is dependent on a number of factors including the pathogenicity of the infecting strain, the stage of gestation that infection occurs in relation to organogenesis and development of immune competence, where infection prior to foetal immunocompetence may result in a persistent infection due to

immunotolerance [5–14]. Persistently infected (PI) animals remain serologically negative and demonstrate cell-mediated unresponsiveness to the infecting strain, shed virus throughout their lives, and are usually epidemiologically more important in ongoing virus transmission than acute, transiently infected animals [5,13–22]. It has been shown experimentally that pigs infected post-natally with Bungowannah virus develop transient infections that resolve over a 10-day period and transmit the virus inefficiently [23]. PI animals are usually the reservoirs of pestiviruses in nature but no PI pigs surviving past 6–8 weeks old have been identified in the affected piggery. Therefore, to better understand how virus transmission is maintained in an infected population it is important to know if long-term infections can occur and to clarify the significance and sources of viral shedding following in utero infections.

This study examined the virological and serological characteristics of in utero infection of the porcine foetus with Bungowannah virus at different stages of gestation. The primary objectives were to determine:


Infection of the porcine foetus with Bungowannah virus was successful, and the virological and serological characteristics following in utero infection at different stages of gestation are reported. The clinical signs and gross pathology are described in an accompanying manuscript [24].

#### **2. Materials and Methods**

#### *2.1. Londitudinal Study Design*

Twenty-four pregnant pigs (22 gilts and two parity-1 sows) with known joining dates were obtained from a piggery known to be free of Bungowannah virus, and sampled and shown to be seronegative on the day of inoculation. Pregnancy was confirmed by ultrasound examination prior to selection for the study. Gilts from this piggery were routinely vaccinated against parvovirus, leptospirosis, erysipelas and *E. coli* with commercial vaccines at selection and again 4 weeks later. Both gilts and sows were also vaccinated against leptospirosis, erysipelas and E. coli at 13 weeks of pregnancy. The animals were moved to facilities at the Elizabeth Macarthur Agricultural Institute 3–5 days before they were due to be infected.

The pigs were challenged intranasally at approximately day 35 (34–36), 55 (55–58), 75 (72–76) or 90 (90–92) of gestation (n = 6 per group referred to as D35, D55, D75 and D90) [24]. These time-points were selected as they were similar to those used in a previous study where foetuses were directly inoculated [25] and they span the gestational age at which the pig foetus is considered to become immunocompetent (70 days). Due to animal accommodation availability and the logistics of undertaking a study of this size, the four treatment groups were challenged and managed separately. The four batch farrowings occurred over a 5-month period. To facilitate intranasal challenge, the pregnant pigs were sedated approximately 30 minutes prior with Azaperone (40 mg/mL—up to 2 mL/20 kg).

All pregnancies were allowed to proceed to 113 days of gestation when all pigs were induced to farrow on day 114 to optimise collection of blood from pigs prior to suckling. Farrowing was induced with 500 μg cloprostenol given intramuscularly on the morning of day 113 and, if required, followed 24 h later with Oxytocin (10 i.u.) intramuscularly.

Pigs from infected litters were weaned when 21–25 days old. As the presence of Bungowannah virus could still be detected at weaning in several challenge groups, as many pigs as the secure containment facilities could hold were weaned and retained until at least 5 to 8 weeks of age (D35, n = 11; D55, n = 22; D75, n = 23; D90, n = 20). Selected pigs from D35 and D55 were kept in the study for a longer period of time to allow ongoing monitoring of virological and serological parameters and clinical signs [24]. Throughout the study, pigs that were moribund, not feeding or did not appear to be viable were euthanased by an intravenous overdose of pentobarbitone sodium.

Pigs from the three litters that did not become infected with Bungowannah virus (on the basis that the virus could not be detected in serum or body fluid for any of the pigs in the litter in a real-time reverse-transcription polymerase chain reaction (qRT-PCR) assay and negative precolostral serology results) were kept as control pigs until they were 14 to 21 days old, at which time they were euthanased.

Individual pig identifications (IDs) have been retained in the text where relevant to facilitate comparison between virological, serological, clinical signs, gross [24] and histopathology findings described in related manuscripts. The first number relates to the litter ID and the second the animal ID within the litter (generally in order of birth) e.g., 8-01 indicates the first piglet to be born in litter 8.

The animal studies were approved by the Animal Ethics Committee of the Elizabeth Macarthur Agricultural Institute, AEC Reference No. M09/02 (6 March 2009).

#### *2.2. Inoculum*

The pregnant pigs were challenged with approximately 5.3–5.8 log10 TCID50 of Bungowannah virus in 5 mL of phosphate buffered gelatin saline (PBGS; pH 7.3, 2.5 mL per nostril), with the exception of Litter 11 where the sow only received approximately 3.8 log10 TCID50 of virus. This inoculum was prepared as previously described [23] and confirmed to be free of porcine parvovirus, porcine circovirus type 2 and other pestiviruses by PCR [26,27]. Testing for porcine reproductive and respiratory syndrome virus was not performed because Australia is free of this virus.

#### *2.3. Sample Collection—Pregnant Animals*

A vaginal swab was collected from each sow on the day of farrowing. In addition, a piece of placenta was collected and swabbed. Vaginal swabs were then collected every 2–3 days until 14 days post-farrowing and then once to twice weekly until 21–28 days.

All swabs collected during the course of the study were placed in 2 mL of PBGS and stored at 4 ◦C prior to testing.

#### *2.4. Sample Collection—Piglets*

A clotted blood sample, and oropharyngeal and rectal swabs were collected from pigs shortly after birth and where possible before they suckled. While this was achieved in most instances, some animals farrowed earlier than 114 days or commenced farrowing earlier than 24 h after receiving the cloprostenol injection and had suckled. For pigs that were stillborn, body cavity fluid (in preferential order of pericardial>pleural>abdominal fluid) was collected rather than serum.

Thereafter, oropharyngeal and rectal swabs and clotted blood samples were collected weekly up to 8 weeks of age. After this time-point, any remaining pigs were sampled every 10–14 days up to 3 months of age. Three animals from the D55 group that were seronegative at birth were followed for an extended period (10-1 to 189 days, and 8-01 and 8-05 to 329 days of age). Urine was collected opportunistically from weaned pigs.

All pigs were subjected to a detailed necropsy and a wide range of samples were collected for virology, serology and histopathology. Samples for qRT-PCR were collected by firmly rubbing a swab across the freshly cut surface of a section of heart, lung, thymus, spleen, small intestine, inguinal lymph node and brain, and from the surface of the tonsil from pigs of infected litters; from pigs of uninfected litters, swabs were collected directly from the surface of the tonsil and from a cut section of lung and spleen.

#### *2.5. Transmission Study Design*

Two 5-and-a-half week-old pigs were obtained from the same piggery as the pregnant animals. After an acclimatisation period of 2 days in isolation, the two naïve pigs were housed for 28 days in the same room and pen with two pigs from D55 (8-01 and 10-01) considered PI with Bungowannah virus. The PI pigs were selected for the study based on low pre-suckle IgG levels (448 and 135 μg/mL), and high quantities of Bungowannah virus RNA in serum (6.9 and 5.5 log10 copies/mL serum) and on oropharyngeal swabs (6.1 and 6.8 log10 copies/swab) at 6 weeks of age compared with cohorts. Retrospective testing demonstrated both were seronegative for Bungowannah virus antibodies at birth, when tested in the peroxidase-linked immunoassay.

#### *2.6. Sample Collection—Transmission Study*

Serum, oropharyngeal and rectal swabs were collected from the introduced pigs on day 0, and then from the introduced and PI pigs on days 1, 8 (serum) or 9 (oropharyngeal and rectal swabs), 15, 22 and 28 days post-introduction.

#### *2.7. RNA Extraction and Real-Time Reverse-Transcription Polymerase Chain Reaction (qRT-PCR)*

Total nucleic acid was extracted from 50 uL of sera and PBGS containing swabs using the MagMax™-96 Viral RNA Isolation Kit (Ambion, Austin, Texas Cat. No. 1836). A Kingfisher®96 magnetic particle handling system (Thermo Fisher Scientific, Waltham, MA, USA) was used for the extraction process and the purified nucleic acid was eluted in 50 μL of buffer.

qRT-PCR was used for the detection and quantification of Bungowannah virus RNA. Five microlitres of nucleic acid was amplified using the AgPath-ID™ One-Step RT-PCR kit (Applied Biosystems, Foster City, CA, USA, Cat No. 4387424) with primers and probe as previously published [28]. The reaction was run in an Applied Biosystems 7500 Fast or 7900HT Real-Time PCR System (Foster City, CA, USA) both run in standard mode. A 10-fold dilution series of Bungowannah virus RNA standards ranging from 107–102 RNA copies/5 μL [25] was included in each assay and the quantity of Bungowannah virus RNA in a sample determined from the standard curve. For quantification purposes 2.3 and 2.6 log10 copies/mL were considered the sensitivity of the assay for serum and swabs/tissues respectively although the accuracy of quantification was considered likely to be variable below 4.3 or 4.6 log10 copies/mL. Additionally, means were calculated against a baseline of 2.3 or 2.6 log10 copies/mL for negative samples depending on the sample type.

#### *2.8. Serology*

A peroxidase-linked immunoassay was used for the detection of Bungowannah virus antibodies [28]. Where an animal recorded an antibody titre ≥5120, a titre of 5120 (12.3 log2) was used for calculation of means; where no antibody was detected, a titre of 8 (3 log2) was assigned for calculation of means.

#### *2.9. Analysis of Results*

To facilitate analysis of data, Bungowannah qRT-PCR results for serum, oropharyngeal and rectal swabs for each challenge time-point were grouped into time intervals based on the age of the pigs (0–1, 2–9, 10–17, 18–24, 25–31, 32–38, 39–45, 46–52, 53–60, 61–70 and 71–80 days). The same process was applied for serology results although the first time intervals were 0 and 1–9 days. Pigs that where known to have suckled prior to Day 0 sampling were classified as having been sampled on day 1 for the purpose of this study. Where an animal had been sampled twice within one of these intervals, the mean result for that animal was used when calculating the time-point mean. In addition, the results for animals in the D35 and D55 challenge groups were subdivided based on the presence or absence of antibodies to Bungowannah virus in samples collected prior to suckling.

#### **3. Results**

#### *3.1. Longitudinal Study*

#### 3.1.1. Pregnant Pigs

All pregnant pigs became infected with Bungowannah virus (as confirmed by seroconversion), with transplacental infection detected for 20 of the 23 (87%) litters. Transplacental infection did not occur for one pregnant pig in each of the D35, D55 and D75 challenge groups [24]. One pig from D55 was found to be not pregnant. For each sow that produced an infected litter, virus was detected in high quantities on vaginal swabs (mean 7.2 log10 copies/swab; range 5.8–8.2 log10 copies/swab) on the day of farrowing, decreasing rapidly by 5–7 days post-farrowing (mean 3.3 log10 copies/swab; range <2.6–5.4 log10 copies/swab) (Figure 1). By day 12–14 post-farrowing the mean amount of virus detected was 2.9 log10 copies/swab (range <2.6–4.3 log10 copies/swab). Viral RNA was not detected on any vaginal swabs collected after 23 days post-farrowing (n = 15). The mean quantity of Bungowannah virus RNA detected on placental swabs at day 0 (7.0 log10 copies/swab; range 5.7–8.1 log10 copies/swab) from the infected litters was comparable to that detected on vaginal swabs. Bungowannah virus RNA was not detected on vaginal or placental swabs collected from sows farrowing uninfected litters.

**Figure 1.** Box and whisker plot summarising the quantity of Bungowannah virus detected in vaginal swabs between 0–29 days post-farrowing following in utero infection (x = mean; — = median; the whiskers extend up from the top of the box to the largest data element that is less than or equal to 1.5 times the interquartile range (IQR) and down from the bottom of the box to the smallest data element that is larger than 1.5 times the IQR. Values outside this range are considered outliers and are represented by dots).

#### 3.1.2. Piglets—qRT-PCR

Bungowannah virus RNA was detected in the serum, body cavity fluid or internal tissues of 225/226 pigs in the 20 infected litters at birth regardless of the stage of gestation that the dam had been infected. It was concluded that one pig (11-15) had not become infected in utero based on the absence of viraemia at birth and because it became seronegative at 9 weeks old indicating the absence of a humoral immune response to infection. This pig was born after it was believed farrowing had concluded and had an antibody titre of ≥10,240 when sampled the next day, presumably due to ingestion of colostrum. No viraemia was detected during the 10 weeks it was followed in the study.

All pigs from three litters (n = 42) were uninfected at birth (based on the absence of viraemia and seronegative serology at birth). The pigs in these three litters all developed high antibody titres after ingestion of colostrum. They did not become infected with Bungowannah virus (based on the absence of a viraemia) despite being in the same room as pigs from infected litters where no direct contact was possible. Occasionally low quantities of Bungowannah virus were detected on oropharyngeal (n = 4; mean 3.5 log10 copies/swab; range 3.3–3.9 log10 copies/swab) and rectal swabs (n = 7; mean 3.2 log10 copies/swab; range 2.7–4.9 log10 copies/swab), presumably due to environmental contamination as a result of sharing a room with an infected litter. Viral RNA was never detected on oropharyngeal swabs from the animal in which 4.9 log10 copies were detected on the rectal swab, or on tissue samples at necropsy. Additionally, all tissue samples collected from these pigs were negative for Bungowannah virus RNA.

Results for the D35 and D55 groups have been subdivided based on whether Bungowannah virus antibodies were detected at birth (Ab +ve) or not (Ab −ve). Samples were collected from all weaned/surviving pigs in the D35, D75 and D90 groups until 75, 56 and 35 days of age respectively. In some cases on the day of euthanasia a direct tonsillar swab was collected rather than an oropharyngeal swab. This result is captured under the results for tissues. In the D55 group, samples were collected through until 120 days in the Ab +ve group (n = 4) and to 329 days in the Ab −ve group (n = 2; 8-01 and 8-05). The number of animals from which serum or body fluid was collected at each time-point for Bungowannah virus PCR is summarised in Table S1.


**Figure 2.** Box and whisker plot summarising the quantity of Bungowannah virus detected in serum between 0 to 75 days of age following in utero infection at one of four stages of gestation (x = mean; — = median; the whiskers extend up from the top of the box to the largest data element that is less than or equal to 1.5 times the interquartile range (IQR) and down from the bottom of the box to the smallest data element that is larger than 1.5 times the IQR. Values outside this range are considered outliers and are represented by dots).

**Figure 3.** Box and whisker plot summarising the antibody titre against Bungowannah virus in serum as measured by peroxidase-linked immune assay between 0 to 75 days of age following in utero infection at one of four stages of gestation (x = mean; — = median; the whiskers extend up from the top of the box to the largest data element that is less than or equal to 1.5 times the interquartile range (IQR) and down from the bottom of the box to the smallest data element that is larger than 1.5 times the IQR. Values outside this range are considered outliers and are represented by dots).

**Figure 4.** Box and whisker plot summarising the quantity of Bungowannah virus detected on oropharyngeal swabs between 0 to 75 days of age following in utero infection at one of four stages of gestation (x = mean; — = median; the whiskers extend up from the top of the box to the largest data element that is less than or equal to 1.5 times the interquartile range (IQR) and down from the bottom of the box to the smallest data element that is larger than 1.5 times the IQR. Values outside this range are considered outliers and are represented by dots).

**Figure 5.** Box and whisker plot summarising the quantity of Bungowannah virus detected on rectal

swabs between 0 to 75 days of age following in utero infection at one of four stages of gestation (x = mean; — = median; the whiskers extend up from the top of the box to the largest data element that is less than or equal to 1.5 times the interquartile range (IQR) and down from the bottom of the box to the smallest data element that is larger than 1.5 times the IQR. Values outside this range are considered outliers and are represented by dots).

#### 3.1.3. Piglets—Serology

The antibody titres of surviving pigs in each of the challenge groups between age 0 to 75 days are presented in Figure 3. Pigs that were known to have fed and were seropositive were excluded from the day 0 antibody titre calculations. The results can be broadly divided into four groups:


#### *3.2. Transmission Study*

Bungowannah viral RNA was detected on oropharyngeal swabs collected from the introduced pigs at 24 h after their entry to the room/pen containing the presumptively PI animals. Peak viraemia and viral shedding were detected at 8 (serum) or 9 (oropharyngeal and rectal swabs) days post-introduction. Both introduced pigs seroconverted between days 8 and 15 with a titre of 5120 recorded at day 15. Figure 7 illustrates the differences in levels of viraemia and shedding observed between the PI and transiently infected pigs in this study. The differences in viral load detected on oropharyngeal swabs were >2 log10 copies per swab higher for the PI pigs compared to the transiently infected in-contact pigs at peak shedding on day 9. This difference was more than 10-fold higher for the rectal swabs. At the time of peak viraemia in the transiently infected pigs the viral load was equal to that of pig 10-01 but still >2 log10 copies/mL less than that detected in 8-01.

**Figure 6.** Quantity of Bungowannah virus detected in serum, oropharyngeal and rectal swabs, and antibody titre against Bungowannah virus detected in: (**A**) 10-01 from birth to 189 days of age; (**B**) 8-01 (D55 Ab −ve) from birth to 329 days of age; (**C**) 8-05 (D55 Ab −ve) from birth to 329 days of age.




**Table 2.** Quantity of Bungowannah virus RNA detected in selected tissues and fluids from pigs 8-01 and 8-05 (D55 Ab −ve) at 11 months of age.

**Figure 7.** Comparison of the quantity of virus detected in serum (**A**), oropharyngeal swabs (**B**) and rectal swabs (**C**) for persistently infected (PI) animals (8-01 and 10-01) and transiently infected in-contact animals (W1 and W2).

#### **4. Discussion**

This study demonstrates that the porcine foetus can become infected with Bungowannah virus following intranasal challenge of the dam and characterises the virological and serological responses following in utero infection of the porcine foetus at different stages of gestation. Persistent infections, as described for other pestiviruses, were observed, as was a chronic infection state where animals that had been presumed to be PI animals later seroconverted and cleared the viral infection. Transmission of Bungowannah virus infection from chronically infected pigs to naïve pigs was readily achieved, with evidence of virus transfer within 24 h.

Preliminary investigations to identify the causative agent of the porcine myocarditis syndrome utilised a direct foetal inoculation route [25]. In the current study, the goal was to determine the outcomes using a natural route of infection at different stages of gestation. Transplacental infection was successfully achieved following intra-nasal exposure of the dam in 87% (20/23) of the litters. In addition, this outcome was not affected by the stage of gestation of the sow at the time of challenge.

Regardless of the stage of gestation that infection occurred, Bungowannah virus was detected in the serum, body fluid and excretions of infected pigs at birth and this was unrelated to the presence of precolostral Bungowannah virus-specific antibody in these animals. Generally, there was minimal difference in virus shedding between the pigs in the different challenge groups at birth, although the mean and median were lowest for the D90 group. The highest viral loads in serum or body fluid were recorded in the D35 (Ab −ve) and D55 (Ab −ve) groups. Over the course of the study it was noted that viral shedding reduced more quickly the later in gestation that the dam was challenged for those pigs which were seropositive at birth or in the D90 group. For those pigs seronegative at birth (D35 and D55 groups), viral shedding remained elevated throughout the course of the study. The exception was those animals that seroconverted after 6–7 weeks (10-01) and between 18 and 28 weeks (8-01 and 8-05) after which viral shedding gradually decreased, although the reduction varied between pigs. In contrast, the viral load in serum declined for all challenge groups after the ingestion of colostrum, but, as maternal antibodies waned in those animals that were seronegative at birth (D35 and D55), the viral load in serum increased as has been described for CSFV [13–15]. Despite animals 8-01, 8-05 and 10-01 seroconverting, it still took approximately 3 months for the viraemia to clear after this event.

It is interesting to compare our results with experimental studies of classical swine fever virus in the porcine foetus which were conducted 40–50 years ago with the less sensitive technologies of virus isolation and antigen detection by immunofluorescence [13,14,21,29]. While test sensitivity may affect the direct comparison of the proportion of animals infected with CSFV at birth compared to Bungowannah virus, the findings are similar. Virus could be detected in a proportion of all pigs born following infection with CSFV both pre- and post-immunocompetence as we have observed. Van Oirschot [13] also described congenitally infected pigs that cleared their CSFV infection by 2 weeks of age. The proportion of pigs that recovered increased the later in gestation that infection took place and was also associated with a low viral load in serum at birth. While precolostral CSFV neutralising antibodies were only detected in one animal (sow infected at 90 days of gestation), these virology findings are similar to those observed for congenital Bungowannah virus infections.

The pig foetus is able to mount a humoral immune response to Bungowannah virus following in utero infection. The highest antibody titres at birth were observed in those pigs that seroconverted and were the progeny of sows challenged at D35 and D55. Only one pig at D70 had no detectable antibody and it was stillborn. Antibody titres in the D90 group were generally low at birth presumably due to the short interval between infection and birth. However, the detection of viraemia indicates that all pigs in the D90 litters were infected prior to birth. Pigs infected post-natally generally seroconvert from 12–14 days post-infection [23]. Therefore, based on the D90 data, it is considered that transplacental and subsequent foetal infection had occurred within 8–10 days of intra-nasal exposure of the sow.

Unfortunately, farrowings were not monitored throughout the night and in some cases pigs were able to feed prior to precolostral blood samples being collected. While not optimal, we were still able to identify pigs in the D35 and D55 groups that did and did not mount a humoral immune response in utero and demonstrate that the time of gestation at which infection occurs has an influence on the ability of the porcine foetus to mount a humoral immune response to Bungowannah virus. The porcine foetus becomes competent to respond to infection with CSFV between days 70–90 of gestation and the serological findings of this study would suggest that the timing is similar, although likely closer to 70 days for Bungowannah virus [13,14,21]. The failure of some pigs in the D35 and D55 litters to seroconvert is indicative of immunotolerance, which occurs following infection prior to foetal immunocompetence and is recognised with other in utero pestivirus infections [5,13,14,21]. In the D55 group it was in Litters 8 and 10, that were challenged at 56 and 55 days of gestation, respectively, that the birth of litters with a mix of both seronegative and seropositive pigs resulted. A previous study [25], suggested that direct in utero transmission of Bungowannah virus to adjacent foetuses is probable and this has also been suggested for CSFV [22]. We speculate that the mixed outcomes of seronegative and seropositive pigs at birth in the D55 and to a lesser extent the D35 group are a

consequence of the timing of foetal infection following transplacental transmission, and timing of any subsequent infections that result from direct in utero transmission.

While the current study did not measure neutralising antibody, the findings have similarities to those of Frey et al. [21] who did not detect neutralising antibody against CSFV in progeny at term when pregnant gilts were infected with CFSV at 65–67 days of gestation. In contrast, 3/4 (75%) of foetuses from gilts infected at 85 days of gestation had neutralising antibodies at term; additionally, antibodies were only detected in 2/6 (33%) of infected pigs when infection occurred after 94 days of gestation. As these foetuses were collected via hysterectomy at term it is not possible to determine if the failure to detect a neutralising antibody response is due to insufficient time between infection and sampling or the result of immunotolerance. Interestingly, Van Oirschot [13] with one exception did not detect precolostral antibodies in any pigs following challenge of the dam with CSFV at 40, 65 or 90 days gestation although the ability of some pigs (highest percentage at 90 days) to clear their infection by 2 weeks of age indicates they were not immunotolerant. Further comparison of antibody titres as measured by PLA and the virus neutralisation test may clarify the nature of the humoral immune response that develops in the period immediately following the development of immunocompetence compared with later in gestation.

The results also suggest that the cell-mediated immune response to Bungowannah virus may be suppressed if the foetus is infected before 90 days gestation or that the antibody response at this time is non-neutralising. In those foetuses from dams infected at 75 days of gestation or earlier that mounted an antibody response, even after ingestion of colostrum, the time taken for these animals to clear the infection was generally delayed compared to those from dams infected at 90 days gestation.

The identification of viraemic pigs that were seronegative following infection of the sow with Bungowannah virus at 35 or 55 days of gestation suggests that persistent infections may result. Persistent pestivirus infections occur following in utero infection of the foetus prior to it becoming immunocompetent and have been described following infection prior to day 70 to 90 of gestation for CSFV, bovine viral diarrhoea virus (BVDV) and Border disease-like viruses in the pig [13,14,30,31]. Despite two pigs in D35 not becoming PI, infection at this stage of gestation results in a high probability of occurrence. By day 55 of gestation the likelihood of maternal infection resulting in persistent infections is declining and may result in a mixed outcome for foetuses within that litter. Based on the results of this study and direct foetal inoculation [25], infection of the pregnant pig after approximately day 60 of gestation (approximately day 70 for foetal infection) seems unlikely to result in persistent infections.

At birth, PI animals could only be differentiated in the laboratory by their seronegative status. While the mean viral load in serum at birth was higher than for those animals born seropositive, individual animal variation was sufficient to prevent the titre of viraemia being a distinguishing feature. Furthermore, it was not until maternal antibodies began to wane from approximately 4–5 weeks of age in the PI pigs that the viral load in serum was higher than the non-PI animals. In contrast, the viral load on oropharyngeal and rectal swabs from PI animals remained elevated throughout the study period but the viral load on swabs for those animals seropositive at birth declined from 3 weeks of age. PI pigs from the D35 group grew poorly and died or were euthanased by 75 days of age [24]. While those from the D55 group were stunted [24], they had survived longer.

Three pigs, from the D55 group (8-01, 8-05 and 10-01) were monitored over a 6 to 11 month period to assess their virological and serological status over time and for any abnormal clinical signs [24]. Unexpectedly, these animals that appeared to be PI seroconverted at varying times during this observation period. This phenomenon has been described rarely in pigs infected with BVDV or border disease virus (BDV) [30,31]. Like these previous studies with BVDV and BDV, the Bungowannah virus viraemia ceased after seroconversion but virus could still be detected in many tissues for a further 5–6 months and at high levels for the female (8-05). For the male (8-01), the highest viral load was 9.8 log10 copies/mL in epididymal semen and was generally above 6.0 log10 copies/mL for other testicular samples (Table 2). Such high virus levels have also been observed in a PI bull [32] and in the pig 8-01 presumably remained high following seroconversion due to the immunologically privileged status of the testes. In contrast, for the female (8-05) the highest viral loads were detected in lymphoid tissues (tonsil and lymph node) (Table 2). It is also of interest to note the high quantity of Bungowannah virus detected in the seminal fluid of the male in the absence of spermatozoa [24]. This has also been described in a boar PI with BVDV [32]. The mechanism for the failure of virus to be cleared from tissues at the same time as the resolution of the viraemia is not clear but may be related to clearance from infected cells being dependent on cell-mediated immunity rather than just the presence of antibody and its inability to access immunologically privileged sites.

Chronic infections resulting from in utero infections with pestivirus have not been reported in ruminants with one exception of BDV in a sheep [17]. It seems probable that the mechanism for development of the chronic infections observed in pigs are related to differences in the time required for maturation of the immune system of ruminants and pigs given that the porcine foetus becomes immunocompetent at a proportionately later stage of gestation compared to ruminants.

While the two chronically infected pigs followed to 11 months of age were quite stunted and would not have been selected as breeding animals [24], they have the potential to be more important in the ongoing transmission of Bungowannah virus than the PI pigs that die early in life. To this end, we were able to show that Bungowannah virus is readily transmitted by these animals to naïve pigs in close proximity.

At least 100–1000 times more Bungowannah virus RNA was detected on oropharyngeal and faecal swabs from pigs at birth in this study compared with peak viral shedding of transiently infected animals [23]. The difference in quantity of virus shed by persistently/chronically infected animals in oropharyngeal swabs compared with peak shedding by transiently infected animals remained approximately 100 times higher until seroconversion was detected in the chronically infected animals. In addition, while the virus load in faeces is negligible in the transiently infected pig, faeces along with urine appear to be an important route of shedding in the PI pig (100–1000 times greater quantity), emphasising the role of PI animals in the epidemiology of pestivirus infections. Placental and vaginal secretions also appear to be an important source of virus for environmental contamination and ongoing transmission and has previously been recognised for BVDV [33]. High quantities of viral RNA could be detected in the placental and vaginal secretions of recently farrowed animals that carried infected foetuses. Although the quantity detected decreased rapidly over the first 5–7 days after farrowing, these reproductive materials could provide a significant source of virus in a population of pregnant animals to sustain transmission cycles even in the absence of a PI animal. This study indicates that all recently farrowed Bungowannah virus infected litters are also potentially adding large quantities of virus into the environment. Further studies are required to determine if these animals are shedding an infectious virus and, therefore, a source of virus for ongoing transmission, although given the quantities of Bungowannah virus RNA detected on oropharyngeal swabs we believe this is highly likely to be the case.

Finally, examination of the virus loads in tissue samples indicates that Bungowannah virus is widely disseminated in the porcine foetus at birth and provides an insight into sample selection for diagnostic purposes from live and dead animals. While the virus could usually be detected in a wide range of samples collected at birth, the optimal tissues to sample for virus detection were serum/body fluid, oropharyngeal swabs, tonsils and lungs. In older animals, virus can also be readily detected in a wide range of samples from PI animals, although serum, oropharyngeal swabs and lymph nodes are preferred for the greatest chance of detection of infection in animals greater than 10 days of age that may not be PI. The high quantity of virus detected in tonsillar samples suggests that the tonsils are the primary source of Bungowannah virus found in oropharyngeal secretions.

#### **5. Conclusions**

The results of this study provide a unique insight into the biology of Bungowannah virus infections in the porcine foetus and the subsequent pre- and post-weaning period following infection at different stages of gestation. While the birth of PI animals was expected based on our knowledge of pestivirus infections in other species, the birth of chronically infected pigs that later went on to seroconvert was unexpected. This study provides further information about how the porcine foetus responds to in utero pestivirus infections.

**Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1999-4915/12/6/691/s1. Summary of the number of animals from which serum or body fluid was collected at each time-point for Bungowannah virus PCR (Table S1).

**Author Contributions:** Conceptualization, D.S.F. and P.D.K.; methodology, D.S.F. and P.D.K.; investigation, D.S.F.; resources, D.S.F. and P.D.K.; data curation, D.S.F.; writing—original draft preparation, D.S.F.; writing—review and editing, D.S.F. and P.D.K.; visualization, D.S.F. and P.D.K.; supervision, P.D.K.; project administration, D.S.F. and P.D.K.; funding acquisition, P.D.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Australian Pork Limited (Project No. 2188), the Australian Biosecurity Cooperative Research Centre for Emerging Infectious Disease and NSW Department of Primary Industries.

**Acknowledgments:** We gratefully acknowledge the staff of the Virology Laboratory at the Elizabeth Macarthur Agricultural Institute (EMAI) for technical assistance (particularly Katherine King), veterinary assistance (Andrew Read), and assistance with sample collection, and both farm and laboratory staff, especially Glenda Macnamara, for their care of the animals used in this study.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*
