Iowa State University Capstones, Theses and Graduate Theses and Dissertations Dissertations
2019 Senecavirus A- a study in immunogenicity, seroprevalence, pathogenesis, and transmission Elizabeth Rose Houston Iowa State University
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Senecavirus A- a study in immunogenicity, seroprevalence, pathogenesis, and transmission by Elizabeth Houston
A thesis submitted to the graduate faculty in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE
Major: Veterinary Preventive Medicine
Program of Study Committee: Pablo Piñeyro, Major Professor James Roth Eric Burrough Luis Giménez-Lirola
The student author, whose presentation of the scholarship herein was approved by the program of study committee, is solely responsible for the content of this thesis. The Graduate College will ensure this thesis is globally accessible and will not permit alterations after a degree is conferred.
Iowa State University Ames, Iowa 2019
Copyright © Elizabeth Houston, 2019. All rights reserved. ii
TABLE OF CONTENTS
LIST OF FIGURES ...... v
LIST OF TABLES ...... vi
NOMENCLATURE ...... vii
ACKNOWLEDGMENTS ...... viii
ABSTRACT ...... x
CHAPTER 1. GENERAL INTRODUCTION ...... 1 Introduction ...... 1 Virus structure ...... 1 History ...... 3 Clinical signs and lesions associated with SVA infection ...... 6 Pathogenesis ...... 9 Immune Response...... 12 Epidemiology and Transmission ...... 13 Diagnosis ...... 15 Oncolytic Properties ...... 17 Thesis Formatting ...... 18 References ...... 18 Tables ...... 27
CHAPTER 2: SEROPREVALENCE OF SENECAVIRUS A IN SOWS AND GROWER- FINISHER PIGS IN MAJOR SWINE PRODUCING STATES IN THE UNITED STATES .... 28 Abstract ...... 28 Introduction ...... 29 Materials and Methods ...... 31 Study design and recruitment of farms ...... 31 SVA recombinant VP1 indirect ELISA (SVA rVP1 ELISA) ...... 32 SVA immunofluorescence assay (SVA IFA) ...... 33 Statistical analysis...... 34 Results ...... 35 Senecavirus seroprevalence in grower-finisher pigs ...... 35 Senecavirus seroprevalence in adult pigs ...... 36 Discussion ...... 37 Conclusion ...... 42 References ...... 42 Figure Legends ...... 46 Tables ...... 51 iii
CHAPTER 3. DEVELOPMENT OF A MOUSE MODEL FOR SENECAVIRUS PATHOGENESIS AND TRANSMISSION...... 53 Abstract ...... 53 Introduction ...... 54 Materials and Methods ...... 56 Determination of mouse strain susceptibility - Study design ...... 56 Determination of mouse infectious route - Study design ...... 57 SVA antibody detection in mice ...... 58 SVA detection in serum, feces and tissue by RT-qPCR...... 59 Histological evaluation and SVA detection in tissue by RNAScope in situ hybridization ... 60 Statistical Analysis ...... 61 Results ...... 61 Senecavirus can cause subclinical infection in different laboratory mouse strains ...... 61 Subcutaneous and intraperitoneal inoculation resulted in subclinical infection ...... 62 Discussion ...... 64 References ...... 68 Figure Legends ...... 71 Tables ...... 80
CHAPTER 4. CHARTERIZATION OF THE IMMUNODOMINANT REGIONS OF SENECAVIRUS A-VP1 STRUCTUAL PROTEIN BY ELISA EPITOPE MAPPING ...... 82 Abstract ...... 82 Introduction ...... 83 Materials and Methods ...... 85 Viruses and cells ...... 85 Generation of SVA-VP1 monoclonal and SVA polyclonal antibodies ...... 85 Indirect rVP1-ELISA and SVA-whole virus ELISA ...... 86 VP1 peptide synthesis and indirect VP1 peptide ELISA ...... 89 Antibody affinity against SVA-rVP1 in blocking ELISA ...... 90 SVA Peptide Blocking ELISA ...... 90 Western Blot ...... 91 Indirect immunofluorescence assay (IFA) ...... 92 Virus Neutralization Assay (VN) ...... 92 Fluorescent Focus Neutralization (FFN) ...... 92 Statistical Analysis ...... 93 Results ...... 94 SVA-rVP1 mAbs generated in vitro do not recognized whole virus particles ...... 94 SVA VP1 mAbs and SVA pAbs can detect virus replication by immunofluorescence assay...... 94 SVA pAbs but not SVA-rVP1 mAbs present neutralizing activity in vitro ...... 95 SVA mAb but not SVA pAb show differential binding activity against SVA-VP1 epitopes by indirect ELISA epitope mapping ...... 95 A group of SVA VP1 linear epitopes shows a differential inhibitory effect on SVA mAb but not in SVA pAbs against rVP1 protein ...... 96 Anatomic location of reactive epitopes in SVA-rVP1 protein ...... 97 iv
Discussion ...... 97 References ...... 101 Figure Legends ...... 105 Tables ...... 112
CHAPTER 5: GENERAL CONCLUSION ...... 113 Summary ...... 113 Conclusions ...... 116
v
LIST OF FIGURES Page CHAPTER 2 Figure 1 Within-herd seroprevalence distribution…………………………………..46 Figure 2 Geographic distribution of grower-finisher seropositive herds by geographical region ……………………………………………………….47 Figure 3 Geographic distribution of sow seropositive herds by geographical region……………………………………………………………………....48 Figure 4 Geographic distribution of grower-finisher seropositive herds by state………………………………………………………………………...49 Figure 5 Geographic distribution of sow seropositive herds by state……………….50
CHAPTER 3 Figure 1 Schematic representation of mouse grouping and housing……………..…71 Figure 2 Serological response to SVA inoculation……………………………….....72 Figure 3 Fecal shedding……………………………………………………………..73 Figure 4 SVA detection in tissue from different mouse strains 14 dpi……...………74 Figure 5 RNAScope detects viral replication in the small and large intestine………75 Figure 6 Humoral immune response by inoculation route…………………………..76 Figure 7 Viremia by inoculation route………………………………………………77 Figure 8 SVA fecal shedding by inoculation route………………………………….78 Figure 9 SVA positive tissues by inoculate route………………………………..….79
CHAPTER 4 Figure 1 SVA pAbs and mAbs characterization by SVA WV and SVA-rVP1 ELISA …………………………………………………………………….105 Figure 2 Characterization of SVA pAbs and mAbs by western blot…………….....106 Figure 3 SVA pAbs and mAbs detect viral replication by IFA…………………….107 Figure 4 SVA pAb and SVA rVP1 mAb neutralizing activity detected by IFA .….108 Figure 5 SVA VP1 epitope mapping evaluated by indirect epitope ELISA …….....109 Figure 6 SVA VP1 epitopes inhibitory effect on SVA mAb and SVA pAb binding activity to rVP1 protein by blocking ELISA ….…………………………..110 Figure 7 SWISS MODEL of SVA VP1…………………………………………...... 111
vi
LIST OF TABLES
Page CHAPTER 1 Table 1 Direct and indirect SVA diagnostic methods……………………………..27
CHAPTER 2 Table 1 ELISA-IFA test agreement………………………………………………..51 Table 2 Pig distribution by state…………………………………………………...52
CHAPTER 3 Table 1 PCR distribution among strains and tissues……………………………….80 Table 2 PCR distribution among routes and tissues………………………………..81
CHAPTER 4 Table 1 VP1 epitope regions……………………………………………………….112
vii
NOMENCLATURE
dpi Days post infection
ELISA Enzyme linked immunosorbent assay
FBS Fetal bovine serum
IFA Immunofluorescence assay mAb Monoclonal antibody pAb Polyclonal antibody
PBS Phosphate buffered saline
PCR Polymerase chain reaction
SVA Senecavirus A
SVA-rVP1 Recombinant SVA-VP1 protein
VI Virus isolation
VN Virus neutralization
viii
ACKNOWLEDGMENTS
I would like to thank my committee chair, Dr. Pablo Piñeyro, my committee members, Dr.
Luis Giménez-Lirola, Dr. James Roth and Dr. Eric Burrough, for their guidance and support throughout the course of this research.
In addition, I would also like to thank my friends, colleagues, the department faculty and staff for making my time at Iowa State University a wonderful experience.
I would like to extend thanks to the following colleagues for their support and tutelage. Dr.
Ronaldo Magtoto for his indispensable guidance in all three projects described here, design of IFA testing, and execution of serological testing for seemingly endless serum samples. Dr. Juan Carlos
Mora-Diaz for design and completion of IFA, VN, and FFA testing, and for teaching me the secrets to cell culture. Dr. Gun Temeeyasen for his wonderful, daily teamwork in examining, weighing, and bleeding the mice, as well as testing of mouse samples. Thank you to Rolf Raul and the
Tetracore, Inc. team for their technical support in completing PCR testing of mouse samples. Dr.
Avanti Sinha provided materials, training, and assistance in completing all western blots for the
VP1 epitope study. To Dr. David Baum for facilitating in farm and veterinarian communication for the seroprevalence study and recruitment of samples. Dr. Fabio Vannucci for design and completion of RNAscope testing of mouse samples. Dr. Nicholas Villarino for statistical analysis of the blocking ELISA tests, and education about using inhibitory concentration 50 when analyzing data.
Finally, my greatest thanks go to Dr. Pablo Piñeyro and Dr. Luis Giménez-Lirola. This work would not have been possible without their unyielding enthusiasm and generous insights. ix
They have provided strong leadership and organization, inspiring me to become a better scientist.
Both have contributed the inciting questions that led to these studies. They have provided the funding, testing platforms, scientific knowledge, and statistical guidance for the three projects described here, along with many more that I have been granted the opportunity to participate in.
x
ABSTRACT
Senecavirus A (SVA), formerly known as Seneca Valley Virus, is a single-strand, positive- sense RNA virus of the family Picornaviridae associated with recent outbreaks of idiopathic vesicular disease and increased neonate mortality in several swine producing countries. SVA is clinically indistinguishable from foot-and-mouth disease and other vesicular diseases, and increased incidence has been seen in the United States and other swine producing countries since
2014. Diagnostic testing must be utilized to confirm infection. The VP1 protein of SVA is a component of the viral capsid which has been used in serologic testing and proven highly immunogenic in other members of the Picornavirus family. Affected and non-affected animals from areas of clinical outbreak have been proven seropositive for IgG against SVA-VP1. In this study, an ELISA using VP1 was completed to determine the seroprevalence of virus antibody in
United States’ swine. A mouse model was explored for use in pathogenicity and transmission.
SVA polyclonal antibodies and SVA recombinant VP1 monoclonal antibodies were generated and characterized through ELISA, western blot, IFA, and viral neutralization.
Serum was collected from 2,433 clinically healthy sows and 3,654 clinically healthy grower or finisher pigs residing at 219 unique commercial swine production sites. SVA seroprevalence was evaluated by SVA rVP1 ELISA and SVA IFA. The estimated seroprevalence for grower-finisher pigs and sows was 12.2% and 34.0%, respectively. The herd prevalence was
42.7 % for grower-finisher farms and 75.8% for sow farms. The SVA rVP1 ELISA and SVA IFA exhibited a fair (sows) and moderate (grower-finisher) agreement at the herd level, while a fair agreement was observed at the individual level for both pig categories evaluated. The McNemar’s test was significant at the individual and herd level (p < 0.05). In this study, we demonstrated the xi
presence of SVA IgG antibodies in pigs from clinically healthy grower-finisher and sow herds.
These results suggest that SVA is circulating subclinically in sow farms and grower-finisher pig farms in major swine producing states in the United States.
Five strains of mice were injected subcutaneously with virus. Then, route of inoculation was explored. Animals were monitored daily. Samples were tested by PCR, ELISA, and IFA to examine viral presence. No mouse showed clinical signs. However, feces and tissue of infected animals tested positive for virus specific nucleic acid. Seroconversion and a transient viremia occurred in mice inoculated by subcutaneous or intraperitoneal route. Mice develop subclinical infection when inoculated with SVA and may play a role in the transmission of SVA as a vector or reservoir for disease.
Eighteen epitopes of the VP1 protein were formed and their reactivity tested through indirect and blocking ELISA. Polyclonal antibodies and monoclonal antibodies were generated and utilized to identify reactive epitopes of the SVA VP1 protein. Four regions of the SVA-rVP1 protein showed high reactivity in multiple testing platforms. Results suggest that immune response generated by SVA-VP1 in mice can be defined by a set of linear epitopes that may be involved in the activity of antibody.
1
CHAPTER 1. GENERAL INTRODUCTION
Introduction
Senecavirus A (SVA) is a non-enveloped, single-stranded RNA virus in the family
Picornaviridae. It is the only member of the genus Senecavirus (1). The virus is composed of four structural proteins (VP1-4) in an icosahedral capsid and was originally discovered in 2002 as a contaminant in cell culture (2). SVA has been associated with vesicular disease and increased neonatal mortality of swine in multiple countries. The extension and distribution of vesicular lesions is indistinguishable from other vesicular diseases including foot and mouth disease virus
(FMDV), vesicular exanthema of swine (VES), swine vesicular disease (SVD), and vesicular stomatitis. The clinical similarity between SVA and other vesicular disease calls for a foreign animal disease investigation at every outbreak of vesicular disease, as FMDV, VES, and SVD are considered exotic for swine in the United States and have severe economic implications. SVA diagnosis is based on clinical signs, detection of the virus, and presence of antibody (3). The disease can also course subclinical and infected pigs can experience a transient viremia, viral shedding, and antibody response without evidence of clinical disease (4).
Virus structure
Members of the family Picornaviridae are characterized by their non-segmented, positive sense single stranded RNA genome. SVA contains a genome approximately 7.2 kb in length and follows a layout typical to the Picornaviruses: leader, four polypeptides of P1, three polypeptides of P2, and four polypeptides of P3 (5, 6). The RNA genome shows an internal ribosome entry site
(IRES) similar to classical swine fever virus and Hepatitis C classified as type IV IRES (5, 7). The
IRES element, located at the 5’ end, assists in cap-independent initiation of protein synthesis (5). 2
These structural and functional resemblances in the IRES between other virus families suggest that genetic exchange between viruses of the Picornaviridae and Flaviviridae family occurred in persistently co-infected pigs (7).
The amino acid sequence of the original, historical SVA isolate (SVV-001) is most similar to the cardioviruses in the family Picornaviridae, with a 35.8% sequence homology (2). The 7,280 base genomic RNA is classified under Genbank Accession No. DQ641257. This encodes a 5’ untranslated region (UTR) 666 bases in length, 4 structural proteins and 7 non-structural proteins matured from polyproteins, then a short 3’ UTR 70 bases in length (5). SVA viral proteins are encoded by the single stranded RNA genome that is translated into one polyprotein. This polyprotein is modified pre- and post-translationally by viral proteases. The 5’ end of the genome contains covalently bound VPg and encodes structural proteins, while the 3’ end has a poly(A) tail and codes for non-structural proteins. SVA replicates at a rapid pace in the cytoplasm of host cells.
The P1 region is translated and cleaved by 3C protease, yielding VP0, VP3, and VP1. VP0 is further processed into VP2 and VP4. VP1, VP2, and VP3 interlace on the external surface of the capsid, with VP4 located internally. The P2 region encodes for 2A protein, which is predicted to play a role in ribosome-skipping. The 2B protein is dissimilar to other picornaviruses in its primary sequence structure and its function is unknown. The 2C protein is similar to helicase and acts in
RNA synthesis. The P3 region of the SVA polypeptide encodes 3A polypeptide and little is known regarding its function. Protein 3C is an enzyme with chymotrypsin-like activity. The 3D polyprotein acts in conjunction with 3AB and 3C in virus replication, uridylation, and as a component of RNA-dependent RNA polymerase (5). 3
The capsid structure of SVA is approximately 30 nm in diameter and is comprised of 60 protomers arranged in an icosahedral manner (8). Each protomer is composed of four structural proteins: VP1, VP2, VP3, and VP4. These proteins have a length of 263, 284, 239, and 72 residues, respectively (8). They show a similar folding pattern to structural proteins in other members of
Picornaviridae. Sixty copies of each protein interlace to create the icosahedral capsid. The capsid of many picornaviruses contains depressions that play a role in receptor binding and evasion of host immune surveillance. The pocket of Senecavirus is similar to that of cardioviruses; the hydrophobic pocket does not contain a lipid pocket factor (9-12). However, unlike cardioviruses, the entrance to the SVA pocket is almost entirely sealed off by residues of the VP1 protein.
Compared to other picornaviruses, characteristics of the SVA capsid proteins are more pronounced, including the BC loop, loop II, and the FMD GH loop of VP1; the puff of VP2; and the knob of VP3. The VP4 protein interacts with internal nucleic acid (8). Additionally, SVA produces an empty procapsid and full virion particles during infection, both of which possess matching antigenic properties. It should be noted that the procapsid appears to require genomic
RNA for stabilization during environmental stressors. Changes in the procapsid could allow for construction of a swine vaccine or nano-carrier in human cancer treatment (13).
History
Seneca Valley Virus (SVV-001) was detected as a contaminant in PER.C6 fetal retinoblast cell culture in 2002 at Genetic Therapy Inc. in Gauthersburg, Maryland, USA, close to the Seneca
Valley geographic region. It is believed that the virus was introduced into cells through bovine serum or porcine trypsin used in cell culture media. The cell culture was cultivating adenovirus-5 based vectors when a rapid onset of cytopathic effect was observed less than 24 hours after infection. SDS-PAGE and electron microscopy yielded the classification of SVV-001 as a 4
Picornavirus (5). Virus isolates were found retrospectively in samples collected from healthy pigs in United States beginning in 1988 (14). In 2016, the International Committee on Taxonomy of
Viruses (ICTV) renamed the only species within the genus Senecavirus from Seneca Valley Virus to Senecavirus A (https://talk.ictvonline.org/ictv-reports/ictv_9th_report/positive-sense-rna- viruses-2011/w/posrna_viruses/234/picornaviridae. Accessed, March 5th, 2018).
During the 1980s, cases of vesicular disease in swine were reported in New Zealand (15,
16) and Australia (17). Although the etiology is still unknown and SVV was not completely ruled out, these cases were associated with contact to vegetation contaminated with the fungus
Sclerotinia sclerotiorum and feeding marine products, respectively. Meanwhile, an outbreak of vesicular disease of unknown etiology was reported in Florida, United States (18). In 2004, a break of vesicular diseases causing lesions on the snout, oral cavity, and coronary bands was reported affecting pigs at several sites of a farrow-to-finish operation in Indiana. A foreign animal disease investigation yielded negative results for foot and mouth disease virus, swine vesicular disease, vesicular stomatitis, and vesicular exanthema of swine by PCR and several serological methods.
The etiology in of this outbreak was not determined, and the clinical signs termed porcine idiopathic vesicular disease (PIVD) (19).
Although the role of SVA was still unknown in 2007, SVA was detected by PCR for the first time in vesicular lesions in finisher hogs imported from Canada to a packing plant in
Minnesota (20). In 2012, another report of SVA in association with the presence of vesicular lesion was made in a boar with a spontaneous outbreak of vesicular lesions after purchase at the Indiana
State Fair (21). Several swine sites in Brazil reported vesicular lesions from November 2014 to early 2015 (22). Vesicular fluid, skin scraping, and cutaneous samples from affected and 5 asymptomatic pigs were evaluated for SVA and other vesicular disease. During this outbreak the development of a specific RT-PCR, targeting the VP1 protein with no cross reaction to other members of the Picornaviridae family, allowed SVA detection from clinically affected animals.
These outbreaks represented the first documented cases of SVA outside of North America (23).
Some herds also described increased neonatal losses (30-70%) of piglets one to four days of age, termed epidemic transient neonatal losses (ETNL). Each outbreak was self-limiting and lasted one to two weeks. The University of Minnesota Veterinary Diagnostic Laboratory received samples from vesicular swabs and were able to generate a full-length sequence of the virus. The strains shared 94.2-96.5% nucleotide identity to the previously documented SVV-001 genome (22).
Interestingly, a serological retrospective study demonstrated that no circulating antibodies were present in the major swine production states in Brazil prior 2014 (24).
During July 2015, an increased incidence of vesicular disease outbreaks was observed in exhibition swine, commercial finisher sites, and breeding herds in Iowa (25, 26). Animals showed acute lameness and coronary band vesicles without mortality (27). Further cases have since been documented in Iowa and Ohio (28, 29). Increased pre-weaning mortality and neonatal lethargy were also reported on multiple producer sites (30). The genomic sequence of three isolates revealed
93.9-94% nucleotide identity to SVV-001 (31), 95.9-96.1% identity to a Canadian isolate, and
97.7-97.9% identity to Brazilian isolates (26). The United States isolates appeared to belong to new clade of SVA, with distinct VP1 sequences from those in other countries (27). Sampling of healthy animals moving to cull swine markets indicated that SVA was circulating, and the US strain was most similar to that isolated in Brazil causing vesicles and ETNL (32). 6
The first reports of SVA in China were made in 2015. Two farms broke with vesicular lesions and lameness, with SVA detected by RT-PCR. Illness spread to surrounding pens, with some piglets displaying vomiting and fever. The sequence of the SVA isolate was most similar to the United States and Brazilian strains (33, 34). Currently there are multiple strains of SVA thought to be circulating in China (31, 35) and more recent Chinese strains have shown residue shifts in the VP1 protein creating a divergence of two clusters among local strains.
In February 2016, swine in Colombia broke with vesicular lesions attributable to SVA, with a 98.5-98.9% nucleotide identity to the US strains (KX857728) (36). SVA was next detected in swine in Thailand in October 2016, where vesicular lesions were reported and SVA isolated.
The two isolates described in Thailand were most similar (98.2% sequence homology) to Canadian strain 11-55910-3, with substitutions in the VP1, 3D, and 3A genes (37). Early in 2018, SVA appeared in Vietnam, causing vesicular lesions and showing a 98.5-99% homology with Chinese strains (KX173339, KX173338, KX173340, KY038016) (38).
Since its discovery, SVA has been identified as a cell culture contaminant, a virus causing with vesicular lesions, and a pathogen associated with neonatal losses. It has been attributed with outbreaks in multiple swine producing countries, with cases appearing in new locations each year.
SVA appears to be on the path to becoming a virus with global prevalence.
Clinical signs and lesions associated with SVA infection
SVA specific neutralizing antibodies have been detected in swine, cattle, and mice.
However, these antibodies have not been detected in humans (14). Viral particles were also detectable in swine tissues, mouse feces and intestines, houseflies, and environmental samples
(39), but cattle samples proved negative for SVA particles on RT-PCR. 7
The signature clinical sign of SVA is vesicular lesions on the nose and coronary band, leading to anorexia and lameness in 10-90% of pigs (25). Clinical disease associated with SVA was first described in swine in 2007. Finisher pigs from Canada processed in a packing plant in
Minnesota developed vesicles along snout, coronary band and lameness. The vesicles were a few millimeters in diameter and filled with abundant translucent fluid. Occasionally vesicles were ruptured, presenting with round, necrotic edges. Other animals had red, coalescing erosions or swollen, blanched coronary band lesions. No pyrexia was observed and other causes of vesicular diseases were ruled out through testing at Plum Island Animal Disease Center (40). In Brazil, additional clinical signs were reported in piglets, beginning in 2015. Increased neonate mortality was found in association with SVA in multiple regions. These piglets were in the first week of life and experienced a variety of signs including weakness, salivation, cutaneous hyperemia, neurologic signs, diarrhea, and sudden death. These clinical signs lingered for 3-10 days before ceasing in surviving piglets. Necropsy revealed petechial hemorrhage of the kidneys, tongue ulceration, and coronary band vesicles (41).
Affected skin displayed orthokeratotic and parakeratotic hyperkeratosis, epidermal hyperplasia, ulceration, and infiltration of neutrophils and cellular debris (21). Experimentally infected finisher pigs had multifocal dermal separation with infiltration of inflammatory cells, necrotic keratinocytes, hemorrhage, and fibrin accumulation. In the first 3-7 days after infection, lymphoid hyperplasia was observed in the tonsils, spleen, and lymph nodes, while the lungs contained mild atelectasis and congestion (42). Microscopic lesions have not been found consistently in other tissues of affected animals (21). 8
Among piglets, common histopathologic lesions included interstitial pneumonia and ballooning degeneration of the urinary bladder and renal pelvis epithelium, with rare intracytoplasmic eosinophilic inclusion bodies (41). Atrophic enteritis of the small intestines was also observed, mirroring the diarrheic gastrointestinal contents found on gross necropsy. A nonsuppurative meningoencephalitis and choroid plexitis was noted in some animals experiencing
ETNL. Skin lesions of these piglets were similar in character to those of adults, with hyperkeratosis, necrosis, and crusting of the dermis (43, 44).
Inoculation of swine with contemporary SVA strain SD15-26 by oronasal route caused vesicular disease in 15-week-old finisher pigs. Experimental inoculation resulted in lameness and lethargy persisting for 2-10 days, with vesicles observed beginning 4 days post inoculation (dpi).
These areas began as regions of epidermal erythema, developing into vesicles 0.5-3 cm in diameter. After rupturing at 5-6 dpi, vesicles resulted in areas of epidermal erosion, ulceration covered by an epidermal crust. Lesions self-resolved by 12-16 dpi (42). Similar results were obtained with intranasal administration of SVA to 9-week-old nursery piglets: early development of vesicular lesions causing mild lameness. In these younger piglets, coronary band lesions were observed beginning 4 dpi, with snout and lip lesions occurring at 5-6 dpi (45).
SVA infection in other species seem to have a subclinical presentation. Experimental studies exploring the oncolytic effect of SVA utilizing murine models noted no clinical signs or lesions related to virus administration (46). No clinical signs have been observed in seropositive cattle (14) nor humans in clinical trials (47, 48), suggesting that swine act as the natural host for
SVA infection.
9
Pathogenesis
The specific mechanism of cell entry and receptors associated with SVA causing clinical infection in pigs is still unknown. In vivo studies have been completed in swine and mice with the only observable viral replication occurring in the epidermis, tonsillar dendritic cell and macrophages in lymph nodes. In vitro studies have utilized swine testis (ST), swine kidney (SK-
RST and PK-15), and human small-cell lung cancer cell lines (NCI-H1299) (14, 49-51) and demonstrated that SVA presents several motifs similar to putative domains present in foot and mouth disease. However, all of these domains have an intracytoplasmic location, and may require joint activity (8). SVV-001 has been tested by in vitro loss-of-function tests in human cancer cell lines to identify anthrax toxin receptor 1 (ANTRX1), also known as tumor endothelial marker 8
(TEM8), as an essential receptor for SVV infection (52, 53). The virus utilizes TEM8 as a transmembrane glycoprotein for cell entry, which is upregulated on tumor cells and bind type VI collagen to promote tumor growth; however the role of TEM8 in vivo is still unclear (54). In addition, SVA infection does not affect the immunoglobulin receptors superfamily-like most other picornaviruses (55). During SVA infection in vitro, interferon α and interferon β signaling is downregulated in permissive cells. This is a common trait in certain cancer cell lines (52). SVV-
001 blocks type 1 interferon (IFN) signaling to evade the host innate immune response. Infection does not directly cause an IFN response. Instead, expression of interferon regulatory transcription factor (IRF3) and IRF7 inhibit IFN. The virus targets mitochondrial antiviral signaling protein
(MAVS), TIR-domain-containing adapter-inducing interferon-β (TRIF), and tumor necrosis factor receptor-associated factor family member-associated NF-κβ (TANK) adaptors of the host cell with
3Cpro to signal shutdown of the IFN response and activate the janus kinase signal transducer and activator of transcription proteins (JAK-STAT) pathway (56, 57). Additionally, the retinoic acid- 10 inducible gene 1 (RIG-1) knockout has also been associated with increased SVV replication and propagation, through decreased IFN signaling (58), suggesting the gene involvement in infection.
Experimental studies in vivo showed that a field isolate, collected from a clinically affected pig with vesicular disease, administered by oronasal route to 15-week-old pigs caused lameness and lethargy 4 dpi. Vesicles, ranging from 0.5 to 3 centimeters diameter, developed on the coronary band and snout in 75% of experimentally infected animals. After 5-6 dpi, vesicles ruptured, progressing to dermal erosion and ulceration that healed completely by 12-16 dpi. SVA was detected by RT-PCR and in situ hybridization (ISH) in all dermal lesions. During this acute clinical presentation viremia spanned 3-10 dpi. An acute seroconversion was also observed with detectable
IgG antibodies by 10 dpi. The IgG response possessed neutralizing activity. Viral shedding was detected by PCR in oral and nasal secretions to 28 dpi, and in feces from 3-14 dpi. The presence of SVA in lymph nodes and tonsil was confirmed by RT-qPCR, virus isolation, and ISH.
Histological changes associated with SVA infection were seen only in cutaneous lesions. Lesions were characterized by multifocal separation of dermis from epidermis, necrotic keratinocytes, fibrin, and inflammatory cells. In addition, there was also mild follicular hyperplasia in superficial lymph nodes (59). Clinical evaluation has also shown that the virus can be present subclinically.
In sows, 50.75% of animals belonging to a herd experiencing a clinical outbreak had positive RT- qPCR of tonsil swabs, while the virus was detected in tonsillar swabs only in 28.12% of their piglets. Although a high percentage of sows were positive by RT-qPCR, only 50% of the positive animals presented with vesicular lesions and clinical signs of SVA infection. No significant differences were observed in viremia or viral shedding in feces and oral fluids in piglets born from clinically or subclinically infected sows. Additionally, all sows developed an SVA-specific IgG 11 response in the six weeks following the outbreak, while 42.8% of piglets from clinical sows and
63.6% of piglets from non-clinically affected sows seroconverted during that time (4).
After a clinical re-emergence of SVA in 2015, phylogenetic evaluation has demonstrated that the virus has gone through genetic variation affecting several structural and non-structural proteins. A pathogenesis comparison study evaluated the historical SVV-001 and contemporary
SVA strains in pigs. It was observed that SVV-001 had a lower virulence when compared to a
SVA strain isolated from vesicular lesions. The historical strain did not induce clinical disease, whereas the contemporary strain caused vesicular lesions in inoculated pigs. Nucleotide differences in the 5’ and 3’ UTRs and long polyprotein-encoding ORF may play a part in this virulence difference. Despite the difference in clinical presentation, cross-reaction of neutralizing antibody and T cell responses were observed, suggesting conservation of antigenic regions (60).
Piglets less than one week of age experiencing disease associated with SVA ETNL, showed interstitial pneumonia; lymphoid depletion in the spleen, tonsils, or lymph nodes; ballooning degeneration of urinary bladder epithelium; and atrophy of intestinal villi. The lymph nodes of newborn pigs with natural infection possessed the highest viral load (61). The virus has been detected by ISH in the transitional epithelium in the kidney and ureters, cerebral choroid plexus, and tongue. In animals presenting with diarrhea, the virus has been demonstrated by immunolabeling in cells of the laminal propria of the small intestine. Further confirmation of SVA by ISH and genome amplification from neonates with enteric signs suggest that the virus may disseminate in newborn pigs by an enteric-neurological method, and then spread by urine and gastrointestinal secretions (62, 63). Although it has not been experimentally demonstrated, it seems that SVA may be vertically transmitted, similar to foot and mouth disease (64). 12
Immune Response
In experimentally inoculated 15-week-old pigs, the presence of neutralizing antibodies were first detected at 5 dpi, with maximum concentration observed between 7 and 14 dpi. This increment in neutralizing antibody concentration was associated with reduction of viremia during the first two weeks post infection. Both antibody isotypes IgM and IgG were detected by IFA at 5 dpi and 7 dpi, reaching a maximum concentration at 10 and 14 dpi, and becoming undetectable by
21 and 25 dpi, respectively. It has also been observed that the dynamic of the IgM antibodies correlated with neutralizing antibody levels. The humoral immune response was further assessed using fluorescent microsphere immunoassays (FMIA) against structure proteins VP1, VP2, and
VP3. The FMIA demonstrated that the presence of IgM and IgG was strongly associated with structural proteins VP2 and VP3 (50, 65). IgG levels in sows increased two to three weeks after onset of clinical signs and remained elevated until six weeks after onset. No study has monitored the IgG levels past this time. In naturally infected sows, specific SVA IgG was present one week after clinical outbreak. During this clinical evaluation, no differences were observed in the antibody dynamic nor levels of IgG between clinically affected and non-affected animals; in addition, piglets born from seropositive sows showed high IgG levels at 7 days of age, with no significant difference based on the sows’ clinical status. These antibody levels declined and disappeared by approximately 6-7 week of age, suggesting that maternal colostrum is involved in neonatal protection rather than an active immune response (4).
SVA infection can also induce a strong cellular immune response. Experimentally infected animals have shown IFN-γ specific T-cells as early as 3-7 dpi. The T cell phenotype observed during SVA infection is also characterized by an early and stronger CD4+ cells response compared to CD8+. The dynamic of the CD4+ response correlates with the humoral response and more 13 specifically with the presence of neutralizing antibody, while CD8+ T cell activity correlated with resolution of clinical signs and viral clearance from replications sites such as the skin. However,
T cell responses did not correlate with viral clearance from the tonsils and lymph nodes, and viral
RNA remained detectable in this these tissues at 14 dpi. Double positive CD4+CD8+ T cells were stimulated by SVA infection and caused an IFN response. This subset of T cells was detected after
35 dpi, past the resolution of clinical disease, suggesting a role as SVA-specific memory T cells.
Both αβ- and γδ-T cells were specifically found in response to VP1, VP2, and VP3 structural proteins at similar frequencies, with a slightly higher response to VP2 (65).
Currently there are no prevention strategies for SVA introduction and no protection methods outside of husbandry and biosecurity. One experimental vaccine study has been completed, in which a strain of SVA isolated in China was inactivated using binary ethylenimine and mixed with oil adjuvant. One dose of vaccine yielded neutralizing antibody in pigs, and vaccinated animals showed no clinical signs after virus challenge (66). Work by Li et al. suggests that use of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR- associated protein-9 nuclease (Cas9) genome editing could be used for cell modification to create an efficient virus cell line for vaccine production (58).
Epidemiology and Transmission
The clinical presentation of SVA seems to be associated with multiple factors. During clinical outbreaks it has been demonstrated that not all infected animals develop vesicular lesions and animals can remain subclinical, suggesting that other factors are necessary to cause disease.
In addition, there is no correlation between both SVA clinical presentations. Not all farms reporting PIVD also report ETNL, and only some herds with SVA associated ETNL report 14 vesicular disease. ETNL in Brazil broke in clusters, even in closed herds. Although is not clear, after the first natural exposure affected farms may develop herd immunity since there are no official reports of clinical re-breaks (67). Risk of SVA infection has a high variation between herds and farm types. Risk factors may include a high number of breeding females, more employees, and biosecurity gaps (25). Commingling of piglets at the time of weaning may contribute to spread of SVA (68). Accelerated hydrogen peroxide has shown efficacy in disinfecting areas contaminated by SVA. (69).
Although swine are the only natural host of SVA, clinical studies have shown that rodents and insects can contribute to the spreading of the virus. During these studies the virus was detected by RT-qPCR on mouse fecal samples, mouse small intestine, and fly samples. SVA presence was later confirmed by virus isolation. During the same clinical evaluation, the virus was detected from swabs of internal and external surfaces on the farm. This suggested that SVA may remain viable in the environment, as well as in mice, which may act as natural reservoir and a potential vector
(70).
Samples collected from swine and their environment at several sites over a twelve-month period from 2015 to 2016 showed that high genetic diversity occurs in SVA over a short time period. Although most of the differences were synonymous, nonsynonymous changes were present in VP1-VP3 regions, 3A, 3C, and 3D. It has been proposed that mutation pressure acts as the main driver of SVA evolution, rather than natural selection (71). It is not yet clear if these structural changes may increase the virulence of the virus in the host; however SVV-001 monoclonal antibodies were not able to bind all these new mutated strains, indicating that changes in binding site may affect the hosts’ humoral protection (72). 15
Diagnosis
The diagnosis of SVA infection can be achieved by direct and indirect pathogen detection
(Table 1). Based on the characteristic cytopathic effect of SVA infection, traditional virus isolation has been reported in PER.C6 fetal retinoblast cells (8), PK-15 cells (23), swine testis cells (26),
H446 small cell carcinoma cells (73), and H1299 human non-small cell carcinoma cells (59).
Cytopathic SVA can be isolated from vesicular swabs and scrapings (27).
The detection of SVA nucleic acid by RT-PCR has improved in diagnosis and helped to understand the pathogenesis of this virus. Numerous RT-PCR assays with different viral targets have been reported and are currently available. A RT-qPCR has been developed for the SVA 3D polymerase gene (74) and is available as a commercial kit (Tetracore, Inc., Rockville, MD). A
Taq-man based RT-qPCR for a conserved region of the VP1 protein (75) and a nested RT-PCR assay (76) have also been described in detail. A reverse transcription loop-mediated isothermal amplification (RT-LAMP) assay has been described as a method for rapid detection that could be used in the field for SVA diagnosis (77, 78).
Although viremia is relatively short and does not last more than 10 days in experimentally infected animals, nucleic acid can be detected by RT-PCR. SVA RT-PCR detection in serum during clinical break has demonstrated that infection can occur subclinically (79). PCR detection has helped to determine viral shedding in feces and oral fluids. Although during the viremic phase of the disease, RT-PCR can detect the virus in almost all tissues from affected animals, after viremia disappears this technique is not capable of detecting the virus outside of a high burden in the tonsils and lymph nodes. 16
The virus can also be detected in situ by ISH and IHC. Specific RNA probes targeting areas of the VP1 gene demonstrated the presence of SVA in vesicular lesions of affected sows, ulcerative lesions in the tongue of piglets, and various other tissues with no evidence of histological lesions (80). The RNAScope ISH technique was capable of detecting virus in situ in SVA PCR positive sample ranging from 12 to 32.6 Ct. Monoclonal antibodies have been used for IHC detection. In neonates without specific vesicular lesions associated with ETNL, viral antigen has been detected in the transitional epithelium of the renal pelvis and urinary bladder, as well as the choroid plexus, enterocytes, and tongue of piglets (81). A serum neutralizing assay has also been described for diagnosis of infection (82).
Experimental studies have shown that after five dpi SVA specific antibodies can be detected in serum. Different structural proteins including VP1, VP2 and VP3 have been evaluated as potential antigen targets in swine serum for IgG detection through indirect ELISA. While the
VP3 protein showed minimal immunoreactivity, both VP1 and VP2 reacted with serum from naturally infected animals (83). A VP1 indirect ELISA yielded a sensitivity of 93% and a specificity of 99% (4). The VP2 protein ELISA was optimized and validated, with 94.2% sensitivity and 89.7% specificity. IFA testing of swine serum showed similar results, with increased sensitivity of the ELISA test during early infection (83). Cross-reactivity has been demonstrated between historical SVV strains and contemporary SVA strains (60). Comparative serological studies have shown that a competitive ELISA using mAbs decreased cross-reactivity and allow a more rapid test conclusion than IFA (50, 84).
17
Oncolytic Properties
Due to its cytopathic effect on cell culture, mostly in neoplastic cell lines, SVA was proven to possess oncolytic properties and has been under investigation as virotherapy for cancer in humans. Some of the characteristics that make this virus desirable for oncolytic treatment are: 1) the virus does not contain a DNA phase during replication, therefore cannot integrate into the mammalian genome; 2) no insertional mutagenesis was noted after several cycles in vivo replication; 3) the virus can induce apoptosis in neoplastic cells; 4) this RNA virus showed a tropism for oncogenic cells; 5) SVV-001 has demonstrated an ability to target and enter solid tumors after intravenous administration.
Senecavirus was able to enter neuroendocrine tumor cells and cause cytotoxicity. SVV-
001 possessed the ability to cause cytopathic effects in neuroblastoma, rhabdomyosarcoma, medulloblastoma, Wilms tumor, and glioblastoma cells (2, 85, 86). The virus required α2,3- and
α2,6- linked sialic acids to infect pediatric glioblastoma multiforme cells (46). Importantly, after viral therapy to human cancer patients, SVA posed a small risk of excretion. The risk of animal infection from humans due to viral excretion was less than 1% (87). Early phases of testing for safety and clinical efficacy began in adult and pediatric cancer patients. Neotropix started development of SVV-001 as drug NTX-010 with Alliance for Clinical Trials in Oncology and
United States national Cancer Institute. Acting to stimulate apoptosis, Phase I trials were completed on neuroendocrine tumors in 2010 (88). Phase II trials were conducted on small cell lung cancers (89) and presented in 2013 at the Annual Meeting of the American Society of Clinical
Oncology. An infectious clone of the virus was able to enter only a small population of small cell lung carcinoma cells and target these cancer cells in mice with tumors (73). However, in 2017, 18 these phase II trials were ended as several studies demonstrated no efficacy in late state disease
(http://adisinsight.springer.com/drugs/800024958, accessed 7/25/2017).
Thesis Formatting
This thesis contains five chapters. The first chapter introduces the project, with a critical review of literature that is currently available regarding Senecavirus A. The second chapter discusses the seroprevalence of Senecavirus A in the United States’ swine producer herds. In the third chapter, a mouse study for pathogenicity and transmission of the virus is described. The fourth chapter describes a project in determining the immunodominant regions of Senecavirus A-
VP1 protein through use of ELISA epitope mapping and other techniques. The fifth chapter provides a summary and general conclusion of the projects discussed in this thesis.
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27
Tables
Table 1- Multiple direct and indirect detection methods are under research for SVA.
SVA Test Published ELISA (VP1) Gimenez-Lirola et al, 2017 Journal of Clinical Microbiology Dvorak et al, 2017 BMC Research ELISA (VP2) Dvorak et al, 2017 BMC Research ELISA (VP3) Dvorak et al, 2017 BMC Research ELISA (WV) Data not published cELISA Goolia et al, 2017 Journal of Veterinary Diagnostic Investigation IFA Goolia et al., 2017 Journal of Veterinary Diagnostic Investigation Dvorak et al., 2017 BMC Research Joshi et al., 2016 Journal general Virology Montiel et al., 2016 Emerging Infectious Disease VNT Goolia et al., 2017 Journal of Veterinary Diagnostic Investigation Saporiti et al., 2017 Veterinary Research Communication Joshi et al., 2016 Journal general Virology ISH Joshi et al., 2016. Journal of General Virology IHC Leme et al., 2016. Journal of Comparative Pathology PCR Bracht et al., 2016., PLoS One Fowler et al, 2017. Journal of Virological Methods Dall Agnol et al., 2017. Molecular Cell Probes Feronato et al., 2017.Tropical Animal Health and Production RT-LAMP Armson et al., 2018. Transboundary and Emerging Diseases Zeng et al., 2018. Journal of Virological Methods
WV= SVA whole virus, cELISA= competitive ELISA, IFA=immunofluorescence assay, VNT= virus neutralization test, ISH= in situ hybridization, IHC= immunohistochemistry, RT-LAMP= reverse transcriptase loop-mediated isothermal polymerization 28
CHAPTER 2: SEROPREVALENCE OF SENECAVIRUS A IN SOWS AND GROWER-
FINISHER PIGS IN MAJOR SWINE PRODUCING STATES IN THE UNITED STATES
Elizabeth Houston, Luis Gabriel Giménez-Lirola, Ronaldo Magtoto, Juan Carlos Mora-Díaz,
David Baum, Pablo Enrique Piñeyro
Modified from a manuscript published in Journal of Preventive of Veterinary Medicine: April
2019.
Abstract
Senecavirus A (SVA) is a single-stranded RNA virus in the family Picornaviridae.
Recently, SVA has been associated with idiopathic vesicular disease and increased neonate mortality outbreaks in the United States, Brazil, China, Colombia, and Thailand, with increasing incidence since 2014. Indirect detection by antibody detection methods, including indirect immunofluorescence assay (IFA), virus neutralization assay, and competitive or indirect enzyme- linked immunosorbent assays (ELISAs), have been reported in clinical and experimental trials.
The objective of this study was to determine the seroprevalence of SVA in nonclinically affected herds in the United States. Individual samples were collected from 3,654 and 2,433 clinically healthy grower-finisher pigs and sows, respectively, from 219 unique commercial swine production sites. SVA seroprevalence was evaluated by SVA rVP1 ELISA and SVA IFA. The estimated seroprevalence for grower-finisher pigs and sows was 12.2% and 34.0%, respectively. 29
The herd prevalence was 42.7 % for grower-finisher farms and 75.8% for sow farms. The SVA rVP1 ELISA and SVA IFA exhibited a fair (sows) and moderate (grower-finisher) agreement at the herd level, while a fair agreement was observed at the individual level for both pig categories evaluated. The McNemar’s test was significant at the individual and herd level (p < 0.05). In this study, we demonstrated the presence of SVA IgG antibodies in pigs from clinically healthy grower-finisher and sow herds. These results suggest that SVA is circulating subclinically in sow farms and grower-finisher pig farms in major swine producing-states in the United States.
Key Words: Senecavirus A, SVA, seroprevalence, SVA recombinant VP1 indirect ELISA, SVA immunofluorescence assay
Introduction
Senecavirus A (SVA) is a nonenveloped, single-stranded RNA virus of the genus
Senecavirus in the family Picornaviridae (1). The virus possesses four structural proteins (VP1-4) composing an icosahedral capsid and seven nonstructural proteins (2). SVA was originally discovered as a contaminant in PER.C6 culture cells in 2002 and has been associated with swine idiopathic vesicular disease (IVD) in Canada and the United States (2, 3). However, retrospective studies demonstrated the presence of SVA in the U.S. swine population for over 30 years (4). Since
2015, several outbreaks of IVD and epidemic transient neonatal losses (ETNL) associated with the presence of SVA have been reported in the United States (5, 6). In addition, numerous countries, including Brazil (7, 8), China (9, 10), Colombia (11), and Thailand (12), have reported an increase in the incidence of IVD cases associated with the presence of SVA. A phylogenetic analysis of the contemporary SVA isolated in the United States, Brazil, China, Thailand, and Colombia shows 30 small genomic differences (< 6%) compared with the historical Seneca Valley Virus strain (11-
15).
The diagnosis of the disease should be based on the presence of clinical signs, detection of the virus, and presence of antibodies against SVA (16). Outbreaks of SVA IVD are characterized by abrupt onset of vesicles that progress to ulcerative dermatitis on the nostril and coronary bands, which affects a high percentage of sows and/or finisher pigs (6, 17). Lesions associated with SVA are clinically indistinguishable from those observed in foot-and-mouth disease, vesicular stomatitis, swine vesicular disease, and vesicular exanthema of swine.
Etiological diagnosis can be achieved by RT-qPCR, virus isolation from vesicular material
(18, 19), in situ hybridization (20, 21), and immunohistochemistry (22). Antibody detection methods currently available include indirect immunofluorescence assay (IFA), virus neutralization assays, competitive enzyme-linked immunosorbent assays (cELISAs), and indirect ELISAs targeting different structural proteins (16, 19, 23-25). However, it has been demonstrated that animals can shed the virus and develop SVA-specific IgG response without evidence of clinical disease, suggesting that the virus may be circulating subclinically (16).
It has been demonstrated clinically and experimentally that SVA infection induces antibodies that can last for at least six weeks post infection, and the dynamic of antibodies detected during the infection may vary based on the antigen target of the serological test used (16, 20, 24).
Thus, the presence of IgG against VP1 protein can be observed as early as 7-10 days post infection
(dpi) (16, 24), whereas the response against VP2 seems to appear earlier, approximately 5 dpi, and can be detected for approximately 60 dpi (24). In contrast, previous reports have shown a weaker and shorter IgG response to VP3, which lasted approximately only two weeks post infection (24).
The duration of the detection of SVA IgG antibody by IFA has been reported to be between 10 31 and 40 dpi (19, 23, 25). Neutralizing antibodies also seem to appear early during infection, and they last for approximately seven weeks post infection (19, 23).
There is no information currently available regarding the seroprevalence or virus circulation of SVA in commercial swine herds, particularly in those which have not reported outbreaks of SVA-associated vesicular disease or increased neonatal mortality. Therefore, the objective of this study was to determine the seroprevalence of SVA in grower-finisher and sow herds in the U.S. using a combination of two serological methods: SVA rVP1 indirect ELISA and
SVA IFA.
Materials and Methods
Study design and recruitment of farms
The study was carried out between June 2016 and December 2016, ensuring that all animals assessed were present in commercial production during the same time period, and encompassing one cycle of rearing market-value animals. Participants in the study were derived from commercial swine farms distributed across the major swine producing states in United States. Referral veterinarians or site manager were interviewed telephonically in order to assess if sites had not had a previous SVA diagnosis, a presence of detectable vesicular lesions, or an increase in neonatal mortality associated with the presence of SVA. After interview screening for clinically negative
SVA farms, farmers (respondents) were asked for their cooperation and willingness to participate, and after that, a verbal consent was obtained. Those who were willing to participate were included in the study. No specific sampling was performed for this study and serum samples were selected from regular submissions for PRRSV monitoring received at the Iowa State University Veterinary
Diagnostic Laboratory (ISU-VDL) and collected from clinically normal herds from multiple 32 geographic regions of the major swine-producing states (n = 18) that voluntarily agreed to participate in the study. Once serum samples or whole blood samples were received from the farm, samples were aliquoted and stored at -80°C until use. Submission samples were collected from breeding herds, gilt development units, farrow-to-finish operations, or wean-to-finish sites.
Assuming a default 50% seroprevalence value with a 95% confidence interval and a 17% absolute error, it was calculated that it was necessary to include at least 30 animals from each site in the study (26). Each site was identified by a unique premise identification number and geographical location. The herd prevalence was presented by geographical regions as follow;
Region 1: (North Dakota (ND), South Dakota (SD), Nebraska (NE), Minnesota (MN), Iowa (IA),
Wisconsin (WI), Illinois (IL); Region 2: Michigan (MI), Indiana (IN), Ohio (OH), Pennsylvania
(PA); Region 3: Wyoming (WY), Colorado (CO), Kansas (KS), Missouri (MO), Texas (TX),
Oklahoma (OK), North Caroline (NC), Idaho (ID). Due to the lack of specific information regarding the age of the animals included in this study, the animals were grouped into two general categories: grower-finisher pigs (6–26 weeks of age) and sows (sows and gilts > 26 weeks old).
Pigs under six weeks of age were not included in the study to avoid the detection of SVA circulating maternal antibodies (Gimenez-Lirola et al., 2016). All serum samples were tested by
SVA rVP1 ELISA and SVA IFA.
SVA recombinant VP1 indirect ELISA (SVA rVP1 ELISA)
The SVA rVP1 ELISA used in this study has been described previously (16). In brief,
NUNC MaxiSorp™ high protein-binding capacity polystyrene 96-well ELISA plates
(ThermoFisher Scientific) were coated with rVP1 protein at 0.18 µg/ml. Positive and negative plate controls, SVA IgG antibody positive and negative serum samples, were run in duplicate on each ELISA plate. One-hundred µL of sera diluted 1:50 were added to each well and incubated at 33
37°C for 1 h. Plates were washed five times with 350 µL phosphate buffered saline containing
0.1% Tween 20 (PBS-T) before adding 100 µL of horseradish peroxidase-conjugated goat anti- swine IgG (Fc) antibody diluted 1:30,000. Peroxidase activity was monitored by adding 100 µL of tetramethylbenzidine-hydrogen peroxide (TMB) substrate solution (Surmodics, Eden Prairie,
MN) for five min at room temperature in the dark. Then, 100 µl of stop solution (Surmodics) was added to each well and the plates were read (450 nm) using an ELISA plate reader (BioTek
ELx800, Biotek® Instruments Inc. Winooski, VT, USA) operated with commercial software
(GEN5TM, Biotek® instruments Inc.). The serum IgG antibody response was represented as sample-to-positive (S/P) ratios.
SVA immunofluorescence assay (SVA IFA)
NCI-H1299 (ATCC® CRL-5803TM, Manassas, VA) cells were seeded (4 x 104 cells/well) in 96-well clear-flat-bottom, black polystyrene surface-treated microplates
(CellBIND®; Corning, Corning, NY) and grown in Roswell Park Memorial Institute 1640 medium
(ATCC® 30-2001TM, Manassas, VA) supplemented with 5% heat-inactivated fetal bovine serum
(FBS, ATCC®, Manassas, VA), 1% penicillin/streptomycin (GibcoTM, Thermo Scientific,
Waltham, MA), and 100 μg/ml gentamicin (Gibco®TM), and incubated at 37°C with 5% CO2 for
24 h. Cell confluency was then evaluated using an imaging cytometer (SpectraMax® MiniMax
300, Molecular Devices, Sunnyvale, CA) operated with SoftMax Pro 6.5 (Molecular Devices,
Sunnyvale, CA) software. When confluency reached > 90% (± 2%), cells were inoculated with
100 µl of SVA field isolate at 5,000 TCID50/ml mixed with the media described above but without
FBS, and incubated at 37°C with 5% CO2. After 24 hours, the inoculum was removed and the cells were fixed with 80% acetone (50 μl/well) for 15 min at room temperature. Plates were air dried, sealed, and stored at -20°C until use. Cell confluency was re-evaluated and plates with < 34
90% cell confluency were rejected. Samples and controls (positive and negative) were tested in duplicate (100 µl/well) diluted 1:40 in phosphate buffered saline pH 7.4 (PBS; Gibco®, Thermo
Scientific, Waltham, MA) with 0.1% BSA (Jackson ImmunoResearch, West Grove, PA), and incubated at 37°C for 1 h. After washing the plates three times with 200 μl of PBS (Gibco®), 50
μl of DyLight® 650 anti-pig IgG (Abcam, Cambridge, MA) diluted in PBS (Gibco®) with 0.1%
BSA (Jackson ImmunoResearch) was added to each well and incubated for 40 min at 37°C. Plates were then washed three times with 200 μl of PBS/well, followed by the addition of 100 μl of PBS per well before reading. Plates were read on the SpectraMax i3x® instrument operated with
SoftMax Pro 6.5 (Molecular Devices, Sunnyvale, CA) software in image-acquisition mode, as follows: one site per well (4.32 mm2), wavelength 713 nm, 30 ms exposure, and -20 μm focus adjustment.
Statistical analysis
The cutoffs and associated diagnostic sensitivity and specificity for both SVA rVP1 ELISA and SVA IFA were estimated using samples of known SVA immune status collected and tested in a previous study (16). For the SVA rVP1 ELISA, we selected an S/P cutoff of 0.4, giving a diagnostic specificity of 99.8% and 93% diagnostic sensitivity (16). For the SVA IFA, we used a
1:40 serum dilution cutoff, which produce a diagnostic specificity of 98.2% and a diagnostic sensitivity of 95 to 100% between 3 to 4 weeks post-exposure (data not published). The diagnostic sensitivity and, more specifically, the time of detection and kinetic of the antibody detection varied with the post-exposure time (16, 20, 24, 27). Based on selected cutoffs, the overall prevalence and
95% confidence interval (CI) was estimated for the total number of animals tested (n = 6087) in each group (grower-finisher and adults) for each serological test (SVA rVP1 ELISA and SVA
IFA). In order to account for the lack of concordance between the results of the two diagnostic 35 assays in individual animals, the within-herd prevalence was also reported using parallel interpretation of the results of two diagnostic assays. However, in order to complement the potential lack of specificity introduced by parallel interpretation of tests within a herd, a farm was considered positive when at least two samples were positive either by SVA rVP1 ELISA or SVA
IFA or combination of both tests. The herd prevalence and 95% CI was calculated for grower- finisher pigs and adult pigs using SVA rVP1 ELISA and SVA IFA. To determine the individual and herd diagnostic agreement between SVA rVP1 ELISA and SVA IFA, a weighted Cohen’s kappa coefficient (k) with a 95% CI was calculated and interpreted as follows: poor, < 0.20; fair,
0.20–0.39; moderate, 0.40–0.59; good, 0.60–0.79; and excellent, ≥ 0.80. Paired results of the two serological methods were analyzed by using McNemar’s Chi squared test to evaluate significant difference between the test results. Statistical analyses and geographical plots were performed using JMP 13.1.0 (SAS Institute Inc., Cary, NC, USA). The within-herd prevalence frequency for grower-finisher pigs and adult pigs was evaluated and plotted using GraphPad Prism® (GraphPad
Software Inc., La Jolla, CA, USA).
Results
Senecavirus seroprevalence in grower-finisher pigs
Serum samples collected from 3,654 grower-finisher pigs (6–26 weeks of age) in 131 premises without previous history of SVA from 16 states throughout the United States were evaluated by SVA rVP1ELISA and SVA IFA. The overall seroprevalence based on SVA rVP1
ELISA was 8.7% (CI ± 0.92; 7.83% to 9.67%), whereas the seroprevalence based on SVA IFA was 5.7% (CI ± 0.75; 4.95% to 6.45%). The overall prevalence obtained by a combination of both serological methods was 12.2% (CI ± 1.06; 11.18% to 13.30%). The herd prevalence was 27.3% 36
(CI ± 7.6; 19.74% to 34.94%) based on SVA rVP1 ELISA and 25.8% (CI ±7.46; 18.34% to
33.26%) based on SVA IFA. The test agreement at the individual and herd level was fair and moderate respectively (Table 1), while McNemar’s test was significant at the individual (p = <
0.001) and herd level (p= 0.009) (Table 1). The estimated herd prevalence obtained using a parallel testing of SVA rVP1 ELISA and SVA IFA results (at least two positive animals either by SVA rVP1 ELISA or SVA IFA or both tests combined) was 42.7 % (CI ± 8.5; 34.2% to 51.1%). The frequency of farms with a within-herd prevalence ranging from 1% to 50%, from 51% to 70%, and from 71%–100% was 70.8%, 5.5%, and 1.8% respectively (Figure 1).
The proportion of positive serum samples among the total grower-finisher pig samples tested by state is shown in Table 2. The three geographical regions showed similar herd prevalence
(Figure 2). The geographical distribution of grower-finisher pigs showed that 15 of the 16 (93.7%) states included in this study were seropositive for SVA antibodies. No seropositive animals were detected in Texas (Figure 4). The highest concentration of positive results at individual and herd levels was found in the Midwest region of the U. S (Table 2; Figure 4).
Senecavirus seroprevalence in adult pigs
A total of 2,433 serum samples collected from clinically healthy sows and gilts (> 26 weeks old) without previous history of SVA, from 91 commercial production sites distributed over 18
U.S. states, were evaluated by SVA rVP1 ELISA and SVA IFA. The overall prevalence obtained with each immunoassay was 26.2% (CI ± 1.76; 24.2% to 27.7%) by SVA rVP1 ELISA and 15.9%
(CI ± 1.46; 14.5% to 17.4%) by SVA IFA. The overall prevalence calculated based on the combination of SVA rVP1 ELISA and SVA IFA results was 34.0% (CI ± 1.88; 32.1% to 35.9%).
The percentage of seropositive farms was 71.4 % (CI ± 9.28; 62.15% to 80.71%) according to
SVA rVP1 ELISA, 48.3 % (CI ± 10.27; 38.1% to 58.6%) according to SVA IFA, and 75.8% (CI 37
± 8.8; 67.0% to 84.6%) according to a combination of these serological methods. The agreement between SVA rVP1 ELISA and SVA IFA at the individual and herd level was fair (Table 1), while
McNemar’s test was significant at the individual level (p = < 0.001) and herd level (p= 0.0027)
(Table 1). The percentage of farms with a within-herd prevalence ranging from 1% to 50%, from
51% to 70%, and from 71% to 100% was 58.2%, 10.98%, and 18.68% respectively (Figure 1).
The proportion of positive animals evaluated by state is shown in Table 2. The herd prevalence with the three geographical regions varied from 69.32% to 83.3% (Figure 3). The geographical distribution of sows herds showed that 14 of the 18 (77.7%) U.S. states included in this study present at least one SVA seropositive premise (Figure 5). No seropositive farms were detected in
Idaho, Texas, Wisconsin or Wyoming. The highest prevalence of SVA seropositive adult pigs, with more than 60% positive samples from the total evaluated was detected in Nebraska, and
Missouri (Figure 5).
Discussion
SVA has been linked to sporadic cases of swine idiopathic vesicular disease (IVD) for over
30 years in the United States (4). However, since 2015 the incidence of IVD and ETNL associated with the presence of SVA has increased (5, 6). Previous individual reports have described only clinical outbreaks of SVA; however, there is no information available regarding the distribution and prevalence of SVA in the U.S.
In the present study, the moderate overall frequency of SVA seropositive grower-finisher pigs detected by a combination of SVA rVP1 ELISA and SVA IFA (12.2%) raises the possibility that animals are being exposed during grower and finisher stages. In this study, we evaluated a broad age range of animals, varying from six weeks old to market weight (on average, 24 weeks old). Previous studies have demonstrated that colostral antibodies decline rapidly and are almost 38 undetectable by six weeks of age in piglets born from seropositive but nonclinically affected sows
(16). Although, we evaluated pigs at six weeks of age and older, the presence of colostral antibodies in the youngest pigs evaluated in this study cannot be completely ruled out.
Furthermore, while all serum samples evaluated were collected from clinically negative sow farms, the overall prevalence in reproductive animals, based on results obtained by a combination of both tests was 34.0%, indicating that SVA is also circulating in reproductive herds. In a previous study, we demonstrated that both clinically and non-clinically affected animals, in a SVA positive farms showed the same antibody kinetic i.e., time of detection, duration of the detection, and magnitude of the antibody response regardless clinical status (Gimenez-Lirola et al. 2016). In the same study performed in a small cohort of neonatal pigs, we observed that animals farrowed from seropositive sows can clear the virus within 6 weeks of age and did not present clinical disease (16). Although there is no definitive information regarding the potential role of lactogenic immunity against SVA associated disease in neonates, these results could have potential implications in reproductive herds during the introduction of non-immune/naïve gilt in to general population. The overall prevalence rate detected at individual level in sows and grower-finishers showed significant differences between both tests used in this study (grower-finishers: 8.7% SVA rVP1 ELISA vs 5.7% SVA
IFA; sows: 26.2% SVA rVP1 ELISA vs 15.9% SVA IFA). Although, both tests showed a high and similar specificity (>98%), estimates of SVA seroprevalence may be influenced by antibody specificity and the serologic method used. Thus, the differences in detection rate between SVA rVP1 ELISA and SVA IFA could associated not only with the performance of these two tests but also with the biological response/variation between individuals evaluated in this cross-sectional study. 39
A seroprevalence study carried out in Brazil indicated that SVA was not circulating in this country before 2014 (2007–2013); nevertheless, SVA was circulating between 2014 and 2016 with an overall seroprevalence of 36.4%, including clinically and nonclinically affected animals (28).
By contrast, our study evaluated the seroprevalence strictly in nonclinically affected herds; therefore, the difference in the overall prevalence reported in our study and the prevalence reported in Brazil could be due to differences in the clinical status of the animals evaluated. Further limitations of the study by Saporiti et al. (28) were the number of samples evaluated (n = 594) and the use of virus neutralization, which is more appropriate to assessing protection compared to the detection of isotype (e.g., IgG) specific antibodies to evaluating SVA seroprevalence. Although it may be reasonable to assume that an individual who is seronegative according to ELISA and IFA likely does not have neutralizing antibodies, no assumptions can be made about the levels of neutralizing antibodies in individuals who are seropositive according to ELISA and IFA.
Regarding the herd prevalence, in this study, 42.7% of the grower-finisher pig farms evaluated were seropositive, but the within-herd prevalence was variable, which may indicate a variable infectious pressure during the grower-finisher stage. Thus, only a low percentage of positive farms (1.8%) exhibited a high within-herd seroprevalence (> 70% seropositive animals), and more than 50% of the farms exhibited a low seroprevalence varying from 1% to 50% seropositive animals. Because no risk factors such as pig density, AIAO/continuous flow, animal movements, and disinfection programs were evaluated in this study, predisposing factors associated with this low–moderate prevalence require further investigation. Here, we demonstrated that SVA is highly prevalent in sow herds in the absence of SVA clinical signs. The frequency of positive sow farms was 75.8%; while the herd prevalence was high, the within-herd prevalence remained low (< 20%) in a high frequency of farms (Figure 1). In a previous longitudinal study, 40 we demonstrated detectable levels of IgG antibodies in sows from an endemic SVA infected farm for a period of six weeks after first detection of the virus by qPCR, regardless of the presence of clinical signs (16). It is also noteworthy that herd detection rate varied between serological techniques methods used in this study. While, there is not a real consensus regarding the number of seropositive animals to define a positive farm, due to the nature of this study and based on parallel testing used to determine the individual overall prevalence and the within-herd prevalence, we decided to set a stringent cut-off of two positive animals to define a positive farm. Thus, observed variation in the herd detection rate by individual testing versus parallel testing could also be associated with the low within-herd prevalence observed in multiple farms.
Numerous field and experimental studies have shown differences in the dynamic of the humoral response against different SVA structural proteins in response to SVA infection (29, 30).
In this study, VP1 protein was selected as the target antigen for ELISA because it is highly conserved (95.8% to 100% amino acid identity) within genus Senecavirus but highly variable, with a low percentage (< 30%) of homology in its amino acid sequence with other members of the family Picornaviridae (e.g., Cardiovirus). Moreover, it has been reported that VP1 elicits an antibody response that allows differentiation among picornaviruses (Kleid et al., 1981).
Experimental and field studies have shown that IgG against VP1 appear within 7–11 days post infection/clinical outbreak and are detectable for at least 6–7 weeks thereafter (16, 24, 27). The use of IFA to detect SVA IgG antibody has been widely described. SVA IFA IgG antibody has been reported from 10 dpi up to 40 dpi (19, 23, 25). To overcome potential differences in test performance due to (1) differences in the dynamic/affinity of the antibody response after natural infection, (2) differences in assay platforms or antigen display, (3) differences in target antigens, 41 and (4) differences in diagnostic specificity, we decided to include two different immunoassay methods for IgG antibody detection methods (SVA rVP1 ELISA and SVA IFA).
A previous study showed a high percentage of agreement (93.7%) between cELISA and
IFA at the individual diagnostic level (23). In this study, we observed a fair agreement between
SVA rVP1 ELISA and SVA IFA at the individual diagnostic level (Table 1). However, significant differences in test agreement was observed between both techniques at individual level and herd level. The proportion of individual animals detected concomitantly by both techniques was lower than the proportion of animals detected by a single technique (Table 1). In a cross-sectional study like this one, the dynamic of antibody response, time of exposure, infectious exposure dose, and immune status of the host are not taken in consideration. Those extrinsic factors, among others, bear no relation to test performance. but could also contribute to the observed differences in detection rate between both techniques. Even though we set a stringent cutoff of two positive animals to determine a positive farm, both tests showed a fair and moderate agreement for sows and grower-finisher herd diagnosis, respectively. Although there was significant difference in the proportion of farms detected by the SVA rVP1 ELISA vs. the SVA IFA, the combination of both techniques greatly improved the SVA antibody detection in sows and grower-finisher herds.
Clearly, these two different techniques detected different SVA seropositive farms; therefore, the additive effect of parallel testing improved the herd detection rate. In addition, the use of parallel testing not only has an impact in the farm detection rate but more importantly, it would help to determine the within-herd prevalence. In non-clinically affected farms, the detection of individual seropositive animals, which seems to be quite frequent, would help to target individual animals for partial or complete eradication programs or prevent movement of seropositive animals that could be a potential risk when introduced in naïve populations. 42
Conclusion
In summary, we established baseline data for the prevalence of SVA in major swine producing states in U.S. swine herds. In total, 3654 serum samples from 219 farms from 18 swine production states throughout the U.S. were evaluated for the presence of SVA antibodies. SVA antibody detection by SA rVP1 ELISA or SVA IFA provided strong evidence of the exposure and immune response against SVA in sows and grower-finisher pigs in the major swine producing- states in the U.S. To date, this is the most comprehensive survey of swine exposure to SVA in the
U.S. swine sow herds and grower-finisher population. Overall, the results obtained in this study showed that SVA is widely distributed and circulating subclinically in the major swine producing states in the U.S. swine population.
Acknowledgments
We would like to thank Calista Knoeke and Nancy Nelson for their technical support.
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46
Figure Legends
30 Grower-finisher pigs
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a R 0 0 0 0 0 0 0 0 0 0 0 -1 -2 -3 -4 -5 -6 -7 -8 -9 0 1 1 1 1 1 1 1 1 1 -1 1 2 3 4 5 6 7 8 1 9 Within herd prevalence (%)
Figure 1: Within-herd seroprevalence distribution. White bars represent the frequency of grower-finisher farms (n =131) with different levels of within-herd seroprevalence. Black bars represent the frequency of sow farms (n =91) with different levels of within-herd seroprevalence.
Only the results of herds with at least one positive were displayed.
47
Figure 2: Geographic distribution of grower-finisher seropositive herds by geographical region. The map represents the percentage of grower-finisher farms detected by geographical regions. Region 1: (North Dakota (ND), South Dakota (SD), Nebraska (NE), Minnesota (MN),
Iowa (IA), Wisconsin (WI), Illinois (IL); Region 2: Michigan (MI), Indiana (IN), Ohio (OH),
Pennsylvania (PA); Region 3: Colorado (CO), Kansas (KS), Missouri (MO), Texas (TX), North
Caroline (NC). The table represent the total number and percentage of positive farms detected by region with the 95% CI (confidence interval).
48
Figure 3: Geographic distribution of sow seropositive herds by geographical region. The map represents the percentage of sow farms detected geographical regions. Region 1: (North Dakota
(ND), South Dakota (SD), Nebraska (NE), Minnesota (MN), Iowa (IA), Wisconsin (WI), Illinois
(IL); Region 2: Michigan (MI), Indiana (IN), Ohio (OH), Pennsylvania (PA); Region 3: Wyoming
(WY), Kansas (KS), Missouri (MO), Texas (TX), Oklahoma (OK), North Caroline (NC), Idaho
(ID). The table represent the total number and percentage of positive farms detected by region with the 95% CI (confidence interval).
49
Figure 4: Geographic distribution of grower-finisher seropositive herds by state. The map represents the percentage of grower-finisher farms detected by states. The table represent the total number and percentage of positive farms detected by stat with the 95% CI (confidence interval).
50
Figure 5: Geographic distribution of sow seropositive herds by state. The map represents the percentage of sow farms detected by states. The table represent the total number and percentage of positive farms detected by stat with the 95% CI (confidence interval).
51
Tables
Table 1 ELISA-IFA Test Agreement
ELISA-IFA test agreement ELISA ELISA Kappa statistic McNemar Test† negative positive (SE; 95% CI) Sow individuals IFA, negative 1591 448 0.21 (0.02; 0.17-0.25) < 0.001 IFA, positive 198 191 Grower-Finisher individuals IFA, negative 3103 238 0.23 (0.02; 0.17-0.28) < 0.001 IFA, positive 130 72 Sow farms IFA, negative 11 20 0.31 (0.1; 0.11-0.51) 0.0027 IFA, positive 5 55 Grower-Finisher farms IFA, negative 58 24 0.49 (0.07; 0.34-0.63) 0.009 IFA, positive 9 40 Cohen kappa coefficient: poor, <0.20; fair, 0.20–0.39; moderate, 0.40–0.59; good, 0.60–0.79;
and excellent, ≥0.80. † Values <0.05 were considered significant.
52
Table 2 Pig Distribution by State
Percentage of positive grower-fishier pigs by Percentage of positive sows by state Pig state inventory by state State Total samples Percentage of 95% CI Total samples Percentage of 95% CI (1,000 head) evaluated (n) positive (%) evaluated (n) positive (%) Iowa 677 16.2 13.42-18.98 295 34.2 28.79-39.61 22,800 North Carolina 199 6.03 2.72-9.34 320 16.2 12.21-20.29 9,000 Minnesota 80 8.7 2.52-14.88 101 14.8 7.92-21.78 8,500 Illinois 703 6.8 4.94-8.66 415 47.9 43.14-52.76 5,350 Indiana 247 16.6 12.0-21.28 61 42.6 30.19-55.01 4,050 Nebraska 165 22.4 16.04-28.76 110 73.3 65.1-81.62 3,600 Missouri 75 4 0.0-8.86 185 61.1 54.05-68.11 3,400 Ohio 396 6.1 3.74-8.46 186 35.5 28.6-42.36 2,950 Oklahoma n/a n/a n/a 55 58.2 45.16-71.24 2,220 Kansas 370 7.6 4.9-10.3 25 20 4.32-35.68 2,110 South Dakota 25 76 59.26-92.74 89 12.3 5.51-19.19 1,526 Pennsylvania 381 18.6 14.69-22.51 277 34.6 29.0-40.2 1,250 Michigan 60 3.3 0.0-7.82 145 12.4 7.04-17.76 1,190 Texas 30 0 n/a 30 3.33 0-9.75 1,040 Colorado 109 29.3 20.76-37.84 n/e n/e n/e 750 Wisconsin 49 6.1 0.0-13.54 28 3.57 0-10.4 350 North Dakota 88 31.8 22.07-41.53 60 28.3 16.93-39.73 147 Wyoming n/a n/a n/a 30 0 n/a 90 Idaho n/e n/e n/e 21 0 n/a n/a Total 3654 12.7 11.62-13.78 2433 34.04 32.16- 35.92 Table 2; 95 % CI confidence interval; n/e not evaluate; n/a: not available
53
CHAPTER 3. DEVELOPMENT OF A MOUSE MODEL FOR SENECAVIRUS
PATHOGENESIS AND TRANSMISSION
Elizabeth Houston, Gun Temeeyasen, Luis Gabriel Giménez-Lirola, Ronaldo Magtoto, Rolf
Rauh, Fabio Vannucci, Pablo Enrique Piñeyro
Abstract
Senecavirus A (SVA) is a RNA virus in the family Picornaviridae. SVA has recently been identified as a causative agent of vesicular lesions and increased neonate mortality in swine. SVA neutralizing antibodies and nucleic acid have been reported in mouse samples from swine facilities. The objective of this study was to evaluate transmission of SVA in experimentally infected mice. To evaluate mice strain susceptibility, ten 8-week-old mice (BALB/C, SWISS,
SJL/J, and C57BL/6) and 4-week-old mice (SCID) were inoculated subcutaneously (SC) with 100
9 µL of SVA at 10 TCID50/mL. Transmission was evaluated by adding 1 mock inoculated mouse per cage for the duration of the study (14 dpi). Intraperitoneal (IP), intranasal (IN), and SC routes of inoculation were assessed for 10 BALB/C mice per group and evaluated for 28 dpi. In all experiments, mice were monitored every 6 hours for the first 72 hours post-infection and daily thereafter. Body weight, clinical signs and virus shedding in feces were collected daily from individual mice. Blood and a select set of tissues (brain, heart, lung, liver, spleen, kidney, and intestine) were collected for viral detection by PCR, histopathology, and in situ hybridization. No differences in clinical signs were observed among strain and route. Except for SCID and control 54 animals, all infected and sentinel mice developed SVA IgG response. Viral RNA was detected in all tissues from directly inoculated mice at 14 dpi, regardless of strain. Viral shedding in feces was observed in infected mice. Sentinel animals had inconsistent shedding between 5 to 10 dpi. In mice inoculated by SC and IP route viremia was detected as early as 3 dpi,. SVA caused subclinical infection in mice and induce seroconversion. Viral shedding in sentinel mice supports previous observations that mice could be a reservoir or vector of SVA in swine herds.
Key Words: Senecavirus, SVA, transmission, mice, shedding
Introduction
Senecavirus A (SVA) is a nonenveloped, single-stranded RNA virus belonging to the genus
Senecavirus in the family Picornaviridae (1). SVA is composed of four structural proteins (VP1-
4) and seven nonstructural proteins (2). It was discovered as a contaminant of PER.C6 cell culture in 2002 and has been associated with swine idiopathic vesicular disease (IVD) (2, 3). SVA has been described in the U.S. swine population for over 30 years (4). In 2015, it was associated with outbreaks of IVD and epidemic transient neonatal losses (ETNL) (5, 6). Such outbreaks were then described in many countries including Brazil (7, 8), China (9, 10), Colombia (11), and Thailand
(12).
Mouse models had been used to study multiple viruses of the family Picornaviridae, including closely related foot and mouth disease virus (FMDV) (13-15). Rodents are near ubiquitous on swine farms and are a biosecurity concern, as they have also been shown to act as a reservoir for encephalomyocarditis virus (16). Research in FMDV epidemiology has determined 55 that mice are susceptible to the virus under experimental conditions, though they are not involved in disease epidemiology (17). Mice were first used in FMDV studies beginning in 1951, where unweaned mice were inoculated by intraperitoneal route (18). This route has since been used in a variety of studies, inducing clinical signs including muscular paralysis when infected before 3-4 weeks of age. Mice do not develop vesicular lesions following infection with FMDV (19). Adult mice show no signs of disease, but some strains are capable of undergoing subclinical infection:
CH3, Swiss, BALB/C, SJL, and nude strains. Infected animals experienced a transient viremia and developed neutralizing antibodies (20, 21). Some serotypes and variants proved lethal for adult
C57BL/6 and BALB/C mice. Additional studies have described inoculation through intramuscular
(22) and subcutaneous foot pad (21), as well as the original intraperitoneal route.
Swine appear to be the only natural host for SVA; however, retrospective studies demonstrated the presence of SVA neutralizing antibodies in mice and cattle. Recent studies showed that environmental samples collected from SVA outbreak sites yielded detection of SVA nucleic acid in both the feces and small intestines of mice sampled on site. Further evaluation confirmed the presence of live infective virus by virus isolation (23).
Mice have also been used in many in vivo models examining the oncolytic properties of
SVA in attempt to develop a therapeutic agent for humans. Neutralizing antibody to the virus has been found in mice used in these studies (24). Liu et al. used an orthotopic xenograft Rag2 severe combined immunodeficient (SCID) mice for a model of pediatric glioblastoma multiforme. For this study, mice were inoculated intravenously with 5 x 1012 viral particles per kg body weight of
SVV-001. The only clinical signs observed in this study were related to the neurologic effects of the tumor (25). Additional studies have used Rag2 SCID mice for medulloblastoma (26) and Y79 56 retinoblastoma models (27). In these experiments, SVV-001 was successful in reducing tumors, but did not directly elicit recorded clinical signs of illness.
The objective of this study was to explore SVA transmission and clinical outcomes in an experimental mouse model. Given the success of previous studies using mice and their prevalence in livestock facilities, they have potential to act as a model for SVA infection, transmission, and pathogenesis.
Materials and Methods
Determination of mouse strain susceptibility - Study design
Three male and three female 8-week-old mice from BALB/C, C57BL/6, SJL/J, and Swiss strains, as well as three male and three female 5-week-old SCID mice were used to determine mice strain susceptibility against SVA infection. Mice were allowed one week of acclimatization in the laboratory animal facility at Iowa State University. At the time of arrival, mice were weighed and ears notched for identification. Mice were housed by strain and sex in groups of three and received standard mouse feed and water ad libitum. Wood shaving bedding and standard enrichment was provided and not changed during the duration of the study.
Each mouse was inoculated subcutaneously in the rear left footpad (RLFP) with 100 µL of
9 SVA at 10 TCID50/mL in RPMI-1640 media. For direct contact transmission evaluation, one female from each strain was designated the sentinel animal and was sham inoculated with RPMI-
1640 media (Figure 1). Mice were monitored every four hours for the first 72 hours, then once daily for the duration of the fourteen-day study. Body weight and clinical signs were evaluated daily. Clinical signs included difficulty breathing, cutaneous rash, and lameness. Viral shedding in feces was evaluated after feces collection from each individual animal once daily. At 14 dpi, 57 mice were administered 65 mg/kg ketamine with 4mg/kg xylazine via intraperitoneal route for anesthesia prior to cardiac puncture for blood collection. Without regaining consciousness, mice were then euthanized using carbon dioxide, followed by cervical dislocation. A necropsy was performed, and a full set of tissues including heart, lungs, spleen, liver, intestine, kidney, and brain were collected. Fresh tissues were stored at -80°C for further SVA PCR evaluation. Tissues were also collected and stored in 10% neutral buffered formalin for histopathological examination and viral detection by in situ hybridization.
Determination of mouse infectious route - Study design
In order to determine the most efficient infectious route, mice were inoculated by several methods. Ten BALB/c mice were inoculated either subcutaneously in the RLFP, intraperitoneally
9 (IP), or intranasally (IN) with 100 µL of 10 TCID50/mL SVA. Mice were clinically monitored as previously described for the mice strain susceptibility study, every 6 h for the first 72 h and daily thereafter for 28 dpi. Feces and weight measurements were collected daily. Blood was collected from the submandibular vein at 0, 3, 5, 7, 10, 14, 17, 21, and 28 dpi. Each mouse was manually restrained and the vein punctured using 23G needle. A maximum of 100 µL of blood was collected in EDTA blood capillary tubes and spun at 2000g for 10 minutes to separate serum. After venipuncture, pressure was applied with gauze until bleeding ceased. At 28 dpi mice were administered 65 mg/kg ketamine with 4mg/kg xylazine via intraperitoneal route for anesthesia prior to cardiac puncture for blood collection. Without regaining consciousness, mice were then euthanized using carbon dioxide, followed by cervical dislocation. A necropsy was performed for macroscopic evaluation and tissue collection. Fresh tissues were stored at -80°C for further SVA
PCR evaluation. Tissues were also collected and stored in 10% neutral buffered formalin for 58 histopathological examination in viral detection by in situ hybridization. This experiment was run in two biological duplicates.
SVA antibody detection in mice
The presence of specific SVA IgG was evaluated in terminal serum by three serological methods including rVP1-ELISA, whole virus ELISA, and indirect immunofluorescence assay
(IFA) for the mice strain susceptibility study. The IgG antibody dynamic was evaluated by IFA for the mouse infectious route susceptibility study.
A rVP1-ELISA was run as previously described (28) for swine serum. In brief, 96-well high binding plates were coated with recombinant VP1 protein at 0.18 µg/mL. One-hundred µL of sera diluted 1:50 were added and incubated at 37°C for 1 h. Plates were washed five times with phosphate buffered saline containing 0.1% Tween 20 (PBS-T) before 100 µL of horseradish peroxidase-conjugated goat anti-mouse IgG (Fc) antibody (Pierce Protein Biology) was added, diluted 1:5,000. Peroxidase activity was monitored by adding 100 µL of tetramethylbenzidine- hydrogen peroxide (TMB) substrate solution for five minutes at room temperature in a darkened environment. The reaction was terminated by addition of 100 µL of stop solution. Finally, optical density was measured at 450nm using an automated plate reader (BioTek ELx800, Winooski, VT,
USA) operated with commercial software. Positive control sera was SVA polyclonal antibody generated in a mouse. Negative control sera was collected from an uninfected laboratory mouse.
Samples were also tested by SVA whole virus ELISA. SVA was clarified and centrifuged for 15 minutes at 4,000 rpm and 4°C. Supernatant was then spun for two hours 28,000 rpm and
4°C. The resulting pellet was resuspended in PBS. One-hundred µL of viral particles were diluted
1:400 in PBS and coated onto 96-well high binding microtiter plates (Thermo Fisher Scientific). 59
Whole virus ELISA plates were incubated for 16 hours at 4°C. Plates were washed five times with
PBST and blocked with 1% bovine serum albumin solution (Jackson ImmunoResearch Inc.) for two hours at 25°C. Samples were diluted 1:50 and a total 100 µL per well were then incubated for one hour at 37°C followed by five washes with PBST. Reaction was revealed by adding 1:10,000 dilution of peroxidase conjugated goat anti-mouse IgG (Fc) antibody (Pierce Protein Biology) for
30 minutes at 37°C. Addition of 100 µL per well of TMB allowed for visualization of the peroxidase reaction. Reactions were stopped with 100uL Nova-stop after 5 minutes. Optical density was read at 450 nm.
Finally, an indirect immunofluorescence assay (IFA) was performed on all samples from
3 NCI-H1299 cells were grown in 96-well plates. Each well was inoculated with 1x10 TCID50/mL
SVA. After 24 hours, cells were fixed with 200 µL cold 90% acetone for 10 minutes at 4°C. Cells were washed with 200 µL PBS, then incubated with 50 µL two-fold serial dilution mouse serum at 37°C for one hour. Wells were washed three times with 200 µL PBS and incubated with 50 µL of FITC labeled anti-mouse IgG (KPL Scientific, Montreal, Quebec, CAN), diluted 1:100, at 37°C for 30 minutes. Plates were washed three times with PBS and examined under a fluorescence microscope.
SVA detection in serum, feces and tissue by RT-qPCR
The presence of SVA was evaluated in feces, serum, and tissue samples using RT-qPCR by Tetracore Inc. Two fecal pellets were added to 0.5 mL PBS and vortex agitated for 20 seconds.
Five grams of tissue, when possible, were suspended in 1:10 weight/volume ratio in PBS, macerated for 10-15 min using a Stomacher 80 lab blender (Seward Laboratory Systems Inc.,
Bohemia, NY), and centrifuged at 1500 × g for 10 min. The obtained tissue homogenate 60 supernatants were aliquoted and stored at -80C until usage. All samples were manually extracted using the QIAamp Viral RNA Mini Kit (Qiagen, Germantown, MD, USA) following the manufacturer’s recommendations. To control the integrity of the extraction and PCR reaction, 6
µL of 800 copies per µL of internal control RNA from Tetracore, Inc. was added to the lysis buffer.
The viral RNA was eluted in 90 µL of elution buffer. Seven µL of each extracted sample was tested using 18 µL of real-time RT-PCR EZ-SVA reagents from Tetracore, Inc. This reagent targeted the conserved sequence of the 3D region of the SVA genome. Real-time RT-PCR was performed on an ABI 7500 Fast instrument (Life Technologies) with the following conditions in standard mode: 1 cycle of 48°C for 15 minutes, 1 cycle of 95°C for 2 minutes, then 45 cycles of
95°C for 5 seconds and 60°C for 40 seconds. To analyze PCR data, the baseline was set to automatic. Threshold settings were set to the maximum of the positive control for SVA and internal control. For a negative result to be considered valid, the internal control for the sample required a
Ct value less than 40.
Histological evaluation and SVA detection in tissue by RNAScope in situ hybridization
All tissues samples collected during necropsy were fixed in 10% buffered formalin, processed for routine histopathologic examination, and stained with hematoxylin and eosin. In situ hybridization (ISH) was performed through the RNAScope platform (Advanced Cell Diagnostics,
Inc., CA). The test targeted the specific reverse complimentary nucleotide sequence of the 301-
345 region of the VP1 gene (GenBank: EU271758.1). Positive signals correlated with metabolically active SVA characterized by the mRNA coding for the VP1 protein. Unstained paraffin tissue sections were processed as previously described (29). These sections were deparaffinized and treated with hydrogen peroxide for 10 minutes. The slides were hybridized 61 using a hybridization buffer and sequence amplifiers were added. Red colorimetric staining detected the SVA hybridization signal and counterstaining occurred with hematoxylin.
Statistical Analysis
Group sizes were established using multiple power analyses using ANOVA and ANCOVA models. GraphPad Prism 7.03 software was used to visualize mean and standard deviation of serologic tests at each time point. For serological and virus detection, an unpaired t test was run to analyze each group against the negative control, using the Holm-Sidak method to correct for multiple comparisons and using a p-value of 0.05 to assess significance.
Results
Senecavirus can cause subclinical infection in different laboratory mouse strains
Throughout the duration of the 14-day study, no vesicular lesions, systemic signs of illness, or significant loss in body weight were observed among the evaluated strains. Clinically, only
BALB/c mice developed mild clinical signs including a rough coat, poor body condition, and a slight decrease in body weight in the three days immediately following infection. However, these changes were not significant compared with other mouse strains or sentinel mice. All infected animals, except SCID mice, developed a SVA IgG antibody response detected by SVA-rVP1 and whole virus indirect ELISA, and IFA (Figure 2A-C). Sentinel mice, housed with infected animals, did not develop a humoral response by 14 dpi. Different levels of viral shedding in feces were observed in infected mice throughout the study. At 24 hours post inoculation, 100% of the SWISS,
BALB/C and SJL mice and 80% of the SCID shed virus in feces. Interestingly, 80% of the sentinel mice housed with subcutaneously infected mice shed virus at 4 dpi. Viral shedding in feces was 62 still detectable in a small subset (approximately 20%) of infected and sentinel mice at the end of the study (Figure 3). Nucleic acid was detected at 14 dpi in all tissues and all inoculated mouse strains (Figure 4). SVA nucleic acid was not detected in tissues of sentinel mice by RT-qPCR, except two mice with a positive detection in the kidney (Table 1).
During histological evaluation, no significant changes were observed among different strains of infected or sentinel mice. Viral replication in tissue was evaluated by RNAScope in situ hybridization. The presence of viral replication was only observed in the small and large intestines of infected mice, mainly in in enterocytes and lymphocytes of the lamina propria. Positive signal was detected in the small and large intestine of BALB/C (3/3), C57BL (2/2), and SJL mice (2/2).
SCID and SWISS mice showed positive signal in (1/2) of both small and large intestinal samples
(Figure 5). No positive hybridization signal was observed in negative control mice.
Subcutaneous and intraperitoneal inoculation resulted in subclinical infection
No significant clinical signs or weight loss were observed throughout the duration of the
28-day study regardless of the inoculation route. No vesicular lesions or systemic signs of illness were observed among the different inoculation route groups compared with the negative controls.
The inoculation route appeared to have an important effect on SVA humoral response.
Mice inoculated by subcutaneous or intraperitoneal route developed a strong IgG antibody response detected by IFA at 7 and 10 dpi respectively. Antibody response increased gradually throughout the study with the highest IgG concentration observed by 17 dpi. No significant differences in fluorescence intensity level (IgG levels) were observed between IN and control groups (Figure 6). 63
A transient viremia was observed in a high percentage of mice inoculated by SC and IP route. Viral RNA was detectable at 3 dpi in 60% of SC and IP inoculated animals, with a gradual decrease in the number of viremic animals in both groups as the study progressed. At 21 dpi, none of the SC mice and only one IP mouse had a detectable viremia. However, an increase in the percentage of viremic animals was observed at 28 dpi, particularly in the IP group. A small subset of mice showed moderate viremia at 3 and 28 dpi in the IN group. Negative control remained negative throughout the study (Figure 7).
Viral shedding was observed in animals inoculated by all three infectious routes. SVA nucleic acid was detected in the feces of 100% of the SC and IN mice, and 50% of the IP inoculated animals 1 dpi. Feces from negative control animals remained negative throughout the study (Figure
8).
Viral RNA was detected in multiple tissues of the SC, IP, and IN groups. Although no clinical signs or significant gross lesions were observed, animals in the IP group showed the highest percentage of detection among the different inoculation routes in the heart, lung, liver, spleen, kidney, and intestine (Table 2). In addition, the IP group also presented significantly higher
Ct values in the heart, liver, kidney, and intestine (Figure 9). SVA nucleic acid was also detected at lower percentage in the SC groups, with a higher percentage of detection in liver and spleen
(Table 2). The liver and spleen also contained the highest observables Ct values among all tissues evaluated for this group (Figure 8). In the IN group, SVA nucleic acid was only detected in heart, and liver with significant lower Ct (Figure 9) values and lower percentage (Table 2) of positive animals compared with IP or SC groups. Interestingly, brains were negative at 28 dpi regardless of the infectious route (Figure 9, Table 2). 64
Discussion
Murine models have been described for the study of many members of the family
Picornaviridae (13, 14). These models have helped develop understanding and dissect the pathogenesis of FMDV and Cardioviruses. In addition, mouse models have been used to lead all advances associated with SVA oncolytic therapy for human cancer, including pediatric glioblastoma multiforme (25), medulloblastoma (26), and retinoblastoma (27).
Previous studies with foot and mouth disease virus have shown that mouse models can reproduce vesicular disease and mortality. However, these clinical outcomes appear to be dependent on mouse strain and age (19). Intraperitoneal inoculation of lactating mice caused development of epithelial ballooning and degeneration of keratinocytes in the footpad. While, different adult mouse strains including CH3, Swiss, BALB/C, and SJL developed subclinical infection after intraperitoneal inoculation with FMDV (20, 21). In this study, five different mouse strains including BALB/C, C57BL/6, SJL/J, Swiss, and SCID were inoculated subcutaneously with SVA. All mouse strains in this study had detectable viremia by 14 dpi, in addition to viral detection in multiple tissues. No significant difference in SVA detection rate were observed among strains. Finally, all mouse strains except for SCID mice developed a strong humoral response by
14 dpi. However, despite this strong evidence of viral replication and systemic spreading of the virus we did not observed lethality or significant clinical signs of systemic illness in any of the evaluated strains. In addition, gross and histological evaluation showed that SVA does not induce cutaneous vesicles or significant lesions in other tissues after 14 dpi. Based on these results, we suggest that SVA can produce subclinical infection in multiple strains of laboratory mice by subcutaneous inoculation. 65
Since no difference in mouse strain susceptibility was observed in our first study, we aimed to determine if changes in infectious route could affect SVA infectivity in BALB/C mice. Previous studies with FMDV explored different infectious routes in adult mice. Mice are more susceptible to SC and IP infection against FMDV, resulting in neurological disease or increased mortality (21).
Although no clinical differences were observed among the different infectious routes in this study,
IP and SC inoculation appeared to cause subclinical infection. In both groups there was acute viremia and viral shedding in feces detected within three dpi. Unlike pigs, fighting and grooming play a large role in the spread of viral disease in mice (30). Our results suggest that SC and IP route better mimic the natural exposures that would occur through biting, grooming, and physical contact than IN route. While mucosal transmission may play a role in mouse infection, IN inoculation without disruption of mucous membranes was not sufficient in causing infection. This ecological information may be crucial in understanding why direct contact between mice is important for viral spread.
Due to the lack of vesicular lesions or lethality associated with SVA infection in different mouse strains and different infectious routes under the experimental conditions used for these studies, a mouse model does not seem to be an appropriate model to study vesicular disease induced by SVA. Although experimental SVA inoculation can produce vesicular disease in grower-finisher pigs, clinical evaluation of sows and their offspring during a break of SVA associated IVD showed that both can be subclinically infected with observable viremia, viral shedding in feces and oral fluids, and acute seroconversion (31). Similar results were observed in this study for multiple mouse strains, and by SC and IP inoculation. The main co-factors necessary to trigger SVA associated vesicular disease in naturally infected pigs are still unknown. Therefore, further studies to evaluate potential co-factors triggering clinical disease in mice would be 66 necessary. Despite the lack of vesicular disease, mice could still be an important model to understand the subclinical presentation of SVA in pigs.
The viremia detected in these studies resembles a pattern of viremia observed in experimentally infected pigs, with an acute onset observed approximately 3 dpi and a duration varying from 10-15 dpi (32). The detection of SVA in multiple tissues by RT-qPCR could be due to 1) active tissue replication or 2) extended viremia. However, virus replication was only detected by RNAScope in situ hybridization in small intestine and large intestine. These results may indicate that nucleic acid detection in other tissues could be the result of viremia without active viral replication. Viral cell tropism during in vivo studies performed in pigs suggested that virus replicated in dendritic cell and macrophages. Antigen detection by IHC showed viral activity in enterocytes of neonatal pigs after an ETNL break (33). Our mouse model showed subclinical infection with persistent viremia and viral shedding in feces, similar to observations in neonatal swine during herd breaks of IVD.
Swine appear to be the only natural host for SVA; however, retrospective studies demonstrated the presence of SVA neutralizing antibodies in mice and cattle (24). In the current study, we evaluated the presence and dynamic of the IgG antibody response in different mouse strains and different infectious routes. The production of IgG antibody after SVA infection in mice is acute, with antibodies levels detectable after 7 dpi in all inoculated strains. This rapid development of specific humoral response is consistent with results observed in experimentally inoculated pigs. Although the antibody neutralizing activity was not evaluated in this study, previous experiments in swine demonstrated that neutralizing activity is likely related to the presence of IgM (34). The IgG antibody response was not affected by strain. All mouse strains 67 used in this study, expect SCID mice, showed a strong humoral response by 14 dpi. Interestingly, sentinel mice showed viral shedding in feces and presence of nucleic acid in the kidney, which may indicate lateral infection. However, no specific antibody response was observed. In addition, in our infectious route study, animals inoculated IN did not showed a specific IgG response. A low infectious dose due to oral infection in sentinel mice and infectious route could be factors associated with the humoral response in experimentally infected mice.
One of the important factors associated with SVA epidemiology and ecology is its persistence in the environment and potential viral vectors or reservoirs on pig farms. Previous studies demonstrated the presence of viable virus in environmental samples surrounding swine herds after an IVD break. These studies also demonstrated that SVA can be re-isolated from mouse feces and intestinal content (23). In this study, viral shedding in feces was observed in all animals, regardless of strain or infectious route. Viral shedding seemed to be transient and only lasting for two weeks in the mouse strain study; however, due to the experimental design, animals housed in the same environment for longer periods of time could be exposed to reinfection by oral exposure, as was observed in animals inoculated IP. Lateral transmission and persistence in the environment are additional factors to consider during eradication or preventive program in swine herds.
Transmission among mice could perpetuate SVA in the environment without clinical evidence of mouse infection. A high percentage of sentinel mice showed viral shedding after three day of cohabitation with experimentally infected mice. Although they remained subclinical, perhaps due to the infectious dose or infectious route, they shed virus for approximately 10 days. These results demonstrate that mice could act as reservoirs or potential viral sources in the environment.
In conclusion, mice may act as a reservoir or vector for SVA infection in swine, with transmission occurring through fecal shedding, urine, or direct contact. Wild mice are widespread 68 across the United States and ubiquitous in production animal housing. Virus may be transmitted mouse to mouse through direct animal or fecal contact, and would allow the virus to travel far distances, reaching previously uninfected swine facilities. Should this be the case, increased pest management and changes in biosecurity could prevent spread of disease among swine herds.
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Figure Legends
Figure 1- Schematic representation of mouse grouping and housing. Mice were housed by sex in groups of three. One sentinel animal was sham inoculated with RMPI-1640 media and housed with two mice inoculated with SVA. 72
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Figure 3- Fecal shedding. Viral shedding was evaluated by RT-qPCR of feces, collected daily. A high percentage of infected animals from all five strains showed fecal shedding beginning 1 dpi, with a small subset of animals in each strain shedding by 14 dpi. Sentinel animals displayed intermittent shedding between 4 and 9 dpi.
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Figure 4- SVA detection in tissue from different mouse strains 14 dpi. The presence of SVA was detected in all tissues evaluated in all infected strains. Bars represent mean and standard deviation of Ct values in five mice per strain. No significant difference in Ct values were observed among tissues or subcutaneously infected mouse strain.
75
Figure 5- RNAScope detects viral replication in the small and large intestine. RNAScope in situ hybridization detected viral replication enterocites and the lamina propria of the small and large intestine of mice from all five strains. No viral replication was observed in sentinel mice. 76
I F A
1 0 0 C o n t r o l
8 0 S C
e c
n I P
e 6 0 c
s IN e
r 4 0
o
u
l F 2 0
0
0 3 5 7 0 4 7 1 8 1 1 1 2 2
D a y s P o s t I n o c u l a t i o n
Figure 6- Humoral immune response by inoculation route. Mice inoculated by SC or IP route developed a humoral immune response detected by IFA beginning 7-10dpi. IN and negative control animals did not demonstrate a humoral immune response throughout the duration of the
28-day study.
77
Figure 7 - Viremia by inoculation route. Mouse inoculated by SC and IP route experienced a transient viremia peaking 3 dpi and gradually decreasing. Viremia was detectable to the end of the study, 28 dpi, in animals from both groups. IN experienced a less consistent viremia, with less than
20% of IN mice positive by PCR only on 3 and 28 dpi. Control animals developed no viremia at any time point.
78
100
Control (-)
s l
a 80 SC
m i
n IP a
60 e
v IN
i
t i
s 40
o
p
f o
20 % 0 0 1 3 7 10 14 21 28 Days post inoculation
Figure 8- SVA fecal shedding by inoculation route. Viral RNA was detected in the feces of
BALB/C mice inoculated with SVA by SC, IP, and IN route beginning 1 dpi. The highest percentage of animals shedding SVA in feces was observed 3 dpi regardless the inoculation route..
SVA was not detected after 10 dpi for IP and IN inoculated animals, and after 14 dpi for SC animals. 79
Figure 9- SVA positive tissues by inoculation route. The study was terminated at 28 dpi and tissues were collected. Using a cut-off of 38, RT-qPCR revealed positive tissues in SC, IP, and IN groups. The highest inverse Ct values were found in the IP group of animals, particularly in the intestines. Additional positive tissues included heart, lung, liver, spleen, and kidney. No brain tissues were positive and control animals had no detectable viral RNA. 80
Tables
Table 1- PCR distribution among strains and tissues. RT-qPCR was run on tissues collected at necropsy. This chart shows the number of positive samples compared to the number of samples tested (in parenthesis) for each tissue type. Every infected strain and tissue possessed nucleic acid.
Sentinel animals did not demonstrate nucleic acid on PCR, with the exception of two kidney samples.
Positive/total (%) Strain Brain Heart Lung Liver Spleen Kidney Intestine SCID 1/5 (20) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) C57BL 4/5 (80) 4/5 (80) 4/5 (80) 4/5 (80) 4/5 (80) 5/5 (100) 5/5 (100) BALB/C 4/5 (80) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) SJL 3/5 (60) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) SWISS 4/5 (80) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) 5/5 (100) Sentinel 0/5 (0) 0/5 (0) 0/5 (0) 0/5 (0) 0/5 (0) 2/5 (40) 0/5 (0)
81
Table 2- PCR distribution among routes and tissues. RT-qPCR revealed SVA specific RNA in multiple tissues when comparing inoculation routes. IP route resulted in the most positive samples, including heart, lung, liver, spleen, kidney, and intestine. The most frequently positive tissue by
IP route were intestine and liver with 95% of animals PCR positive. SC and IN route also resulted in detection of RNA in multiple tissues. However, a lower percentage of animals yielded PCR positive tissues. No SVA specific RNA was detected in any control animal tissue or in the brain of any tissue.
Positive/total (%) Route Brain Heart Lung Liver Spleen Kidney Intestine Subcutaneous 0/10 2/20 0/20 15/20 18/20 2/20 9/20 (0) (10) (0) (75) (90) (10) (45) Intraperitoneal 0/10 7/20 15/20 19/20 15/20 6/20 19/20 (0) (35) (75) (95) (75) (30) (95) Intranasal 0/8 2/18 8/18 2/18 1/18 0/18 2/18 (0) (11.1) (44.4) (11.1) (5.5) (0) (11.1) Control (Neg) 0/10 0/17 0/17 0/17 0/17 0/17 0/17 (0) (0) (0) (0) (0) (0) (0)
CHAPTER 4. CHARTERIZATION OF THE IMMUNODOMINANT REGIONS OF
SENECAVIRUS A-VP1 STRUCTUAL PROTEIN BY ELISA EPITOPE MAPPING
Elizabeth Houston, Luis Giménez-Lirola, Avanti Sinha, Juan Carlos Mora-Díaz, Nicholas
Villarino, Pablo Piñeyro
Abstract
Senecavirus A (SVA) is a RNA virus in the family Picornaviridae. The virus has been recently detected in swine production systems, causing vesicular disease and neonate mortality.
The viral capsid is composed of four structural proteins: VP1-VP4. Although the VP1 protein has been reported as the most immunogenic proteins in vivo, no information on immunodominant regions of the SVA polyprotein is available. The objective of this study was to identify immunodominant regions of SVA polyprotein using epitope mapping. The binding effect of SVA polyclonal antibody (SVA-pAb), SVA-VP1 monoclonal antibodies (mAb), and SVA positive sera from clinically affected animals were characterized using a set of SVA-VP1 derived peptides by indirect and blocking ELISAs. Epitope array was based on 18 SVA-VP1 peptides, overlapping 20- mer and shifted by five amino acids. All VP1-derived peptides yielded significant signal against
SVA-pAb and VP1-mAb by indirect ELISA. One peptide (aa 1-20) showed significantly higher
OD on rVP1 and SVA whole virus indirect ELISAs. Peptides spanning aa 1-20, 60-80, 105-125,
120-140 showed a significant half maximal inhibitory concentration (IC50) on VP1-mAb against 83
rVP1 by blocking ELISA. Although several peptides showed variable binding inhibitory effect of
SVA-pAb and SVA-positive clinical samples, none of the peptide reach a 50% inhibitory effect.
These results suggest that the humoral immune response against SVA-VP1 can be defined by a set of linear epitopes. Further investigation is warranted to determine if these peptides are indeed responsible for neutralizing antibodies against SVA.
Key Words: Senecavirus, VP1, immunogenicity, epitopes, ELISA
Introduction
Senecavirus A (SVA), formerly known as Seneca Valley Virus, is the only member of the genus Senecavirus in the family Picornaviridae. This single-stranded, positive-sense RNA virus was originally discovered as a contaminant in PER.C6 culture cells and described retrospectively in United States’ swine herds (1-3). Recently, SVA infection has been associated with idiopathic vesicular disease. In 2015, outbreaks of vesicular disease and neonatal mortality related to SVA were reported in numerous swine producing countries, including the United States (4-7), Brazil
(8), Colombia, Thailand, China (9), and Vietnam (10). SVA vesicular presentation is characterized by the presences of vesicles on the snout or coronary band, causing transient lameness, anorexia, and fever. Vesicular lesions associated with SVA are clinically indistinguishable from those seen in foot-and-mouse disease, vesicular stomatitis, swine vesicular disease, and vesicular exanthema of swine (8, 11, 12). A final etiological diagnosis is required to distinguish SVA from other vesicular disease. 84
SVA has a genome length of approximately 7.3kb, which encodes three polyproteins, named polyprotein 1 to 3 (P1, P2, P3). The P1 region is both pre- and post-translationally cleaved into structural proteins: viral protein 1-4 (VP1-4) (3). Sixty copies of each viral protein make up an icosahedral capsid, with VP1-3 interlocking on the surface and VP4 found on the interior. While the pathogenic role of VP1 is unknown in SVA, it has proven highly immunogenic in SVA and other viruses of the Picornaviridae family (13, 14). The VP1 protein is also highly specific to each
Picornavirus, with low sequence similarity between species. It is highly conserved within the species, with different strains showing 99-100% nucleotide identity and 100% amino acid identity
(15). Crystallography has shown that SVA VP1 has multiple exposed loops that may be responsible for cell tropism and binding, including the BC loop, loop II, and FMDV loop. Unlike other picornaviruses, the hydrophobic pocket formed by the capsid proteins of SVA possess an entrance without a pocket factor that is nearly sealed. Due to the pocket structure, receptors and host cell binding does not mimic that described for other members of Picornaviridae (16). While recent studies have revealed affinity of the ANTXR1 cell receptor, commonly upregulated in tumor cells (17-20), no work has been completed with swine samples and information regarding the immunodominant regions of VP1 is not yet available. Additional knowledge on the antigenic regions of the VP1 protein would allow for better understanding of SVA pathogenesis and immunodiagnostics.
In this study, SVA polyclonal antibodies and monoclonal antibodies of the SVA VP1 protein were produced and characterized to map antigenic epitopes on the VP1 protein. These epitopes were further evaluated by SVA antisera from experimentally infected mice and swine samples collected from a clinical outbreak. 85
Materials and Methods
Viruses and cells
A SVA strain was isolated from grower-finisher pigs with vesicular lesions confirmed SVA positive by PCR at the Iowa State University Veterinary Diagnostic Laboratory. A NCI-H1299 cell line (ATCC® CRL-5803) was maintained in RPMI 1640 (Gibco®, Carlsbad, CA, USA) at
37°C and 5% CO2. Media was supplemented with 10% heat-inactivated fetal bovine serum (FBS,
ATCC), 1% antibiotic-antimycotic (Gibco®) and 0.1% gentamicin (Gibco®).
Generation of SVA-VP1 monoclonal and SVA polyclonal antibodies
Polyclonal antibodies (pAbs) were generated by intraperitoneal inoculation of mice with
6 inactivated SVA whole virus. A total of 10mL of 1x10 TCID50/mL of passage one SVA was treated with 5% formalin and ultracentrifugated for two hours at 35,000 rpm. The viral pellet was resuspended in 400 µL of phosphate buffered saline (PBS) and an aliquot of 50 μL was mixed with incomplete Freund's adjuvant at 1:1 volume ratio. Three six-week-old BALB/C mice were inoculated intraperitoneally, followed by a booster injection two weeks after primary inoculation.
Beginning two weeks after the booster inoculation, blood was harvested and presence of SVA antibodies evaluated weekly by SVA-rVP1 ELISA. When SVA-rVP1 antibody titer reached a maximum concentration, mice were treated intraperitoneally with Pristane (Sigma-Aldrich, St.
Louis, MO) and three days later given an intraperitoneal injection of SP2/O cells (approximately
5 to 8 x 106 cells) to produce polyclonal ascites fluid. After 10 days, all mice were euthanized.
Ascites fluid was collected and stored -20ºC. 86
Monoclonal antibodies (mAbs) against the recombinant SVA-VP1 (SVA-rVP1) protein were generated by GenScript (Piscataway, NJ, USA). Five six-week-old BALB/c mice were injected subcutaneously with 0.1 mg of SVA-rVP1 protein using MonoExpress immunization technique, twice at two-week intervals. Spleen cells were gathered and fused with SP2/0 myeloma cells. Hybridoma supernatants of parental clones were suspended in Dulbecco’s Modified Eagle’s
Medium (DMEM) with 10% FBS, 1x HT supplement, 1x penicillin streptomycin, and 0.02%
Sodium Azide. A total of 20 hybridomas were screened using indirect SVA-rVP1 ELISA and three hybridomas were selected for mAb production and purification: 4D10, 8F12, 10C2.
The concentration of all SVA VP1-mAbs was normalized to 0.092 mg/mL for all ELISA testing. Isotyping was completed for using a rapid mouse immunoglobulin ELISA (Antagen
Pharmaceuticals Inc, Boston, MA, USA) following vendor protocol. Briefly, each SVA VP1-mAb was diluted to 2 µg/mL and 100 µL was added to each cassette sample well and incubated for five minutes at room temperature. Antibody isotyping was read by red colorimetric changes in the specific site for each antibody class and subclass.
Indirect rVP1-ELISA and SVA-whole virus ELISA
A recombinant VP1 protein was generated in an Escherichia coli expression system. A codon-optimized version of the VP1 gene, 1,359 nt in size (Genbank #DQ641257) was synthesized in vitro (Sanghai Genery Biotech Co. Ltd, Shanghai, China) and amplified using a forward primer
(5’- CAT CAT CAT CAT CAT CAT ATG TCT ACA GAT AAT GCA GAA ACG- 3’) and reverse primer (5’- AGA CTG CAG GTC GAC AAG CTT TTA ACC TGA CTG CAT CAG CAT
C-3’). This PCR product was cloned into pCold II expression plasmid using the NovoRec © PCR
One Step Directed Cloning Kit (Novoprotein Scientific Inc, Shanghai, China) and confirmed 87
through sequencing (Genewiz Inc, Suzhou, China) before being transformed into E. coli BL21
(DE3) pLysD (Rosetta cells; InvitrogenTM, Carlsbad, CA, USA). This bacterial clone was grown in Luria-Bertani (LB) medium (InvitrogenTM) with 100µg/mL ampicillin and shaking 250 rpm at
16 °C. After sixteen hours, cells were chilled to 4 °C and harvested by centrifugation at 3,500g for fifteen minutes. Cells were then resuspended in 20mM PBS, 500mM NaCl pH 7.4 and lysed by ultrasonication. The extracts were centrifuged at 50,000g for thirty minutes at 4 °C and fractions analyzed by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). The
VP1 protein at 30.9 kDa was expressed as an inclusion body in the precipitate, and solubilized using a denaturing buffer including 20mM Tris, 6M Guanidine-HCl, 10mM β-Mercaptoethanol pH 8.0. The rVP1 protein was refolded in vitro to the native conformation and concentrated by dialysis with a refolding buffer consisting of 50mM Tris, 240mM NaCl, 10mM KCl, 2mM MgCl2,
0.4M sucrose, 0.5M arginine, 0.05% Triton X-100, and dithiothreitol pH 8.2. The 278 amino acid sequence identity was confirmed.
The rVP1 protein was utilized in ELISA testing as described by Gimenez et al. (2016).
Ninety-six well high binding microtiter plates (Maxisorp, Nunc, Thermo Fisher Scientific,
Agawam, MA, USA) were coated with 100µL of rVP1 at 0.18 µg/mL/well in a carbonate buffer, pH 8.2. Plates were incubated sixteen hours at 4 °C, then washed five times with phosphate buffered saline (PBS), pH 7.4, containing 0.1% Tween 20 (PBST) and blocked with a 1% weight/volume bovine serum albumin solution (Jackson ImmunoResearch Inc., West Grove, PA,
USA) for two hours at 25 °C. Plates were dried at 37 °C for three hours and stored at 4 °C in a sealed bag with desiccant packets. 88
One-hundred µL of sample was added per well and incubated at 37°C for one hour.
Samples were run in triplicate with a positive and negative control. Mouse serum, including mAbs and pAbs, was diluted in a newborn calf serum-based diluent at 1:50. Swine serum was diluted in a fish serum based diluent at 1:50. Plates were washed five times with PBST and then incubated with 100 µL of horseradish peroxidase-conjugated goat anti-mouse IgG Fc antibody (Pierce
Protein Biology) diluted 1:5,000. Alternatively, for swine serum, goat anti-swine IgG Fc HRP conjugated antibody (Bethyl Laboratories, Montgomery, TX) was used at a 1:30,000 dilution.
Peroxidase activity was monitored by adding 100 µL of tetramethylbenzidine-hydrogen peroxide
(TMB) substrate (Surmodics, Eden Prairie, MN, USA) for 5 minutes at room temperature in a darkened environment. The reaction was terminated by addition of 100 µL of Nova-stop
(Surmodics). Finally, optical density was measured at 450nm using an automatic plate reader
(BioTek ELx800, Biotek ® Winooski, Vermont) operated with a commercial software (GEN5TM,
Biotek ®).
For SVA whole virus ELISA, virus was harvested by three cycles of freezing and thawing of the NCI-H1299 cell line (ATCC® CRL-5803) cells previously infected with SVA, recovered from a clinical case of vesicular disease. Cell culture supernatant was centrifuged for 15 minutes at 4,000 rpm and 4°Csupernatant was then spun for two hours 28,000 rpm and 4°C, and the resulting pellet was resuspended in PBS. One-hundred µL of viral particles were diluted 1:400 in
PBS and coated onto 96-well high binding microtiter plates (Thermo Fisher Scientific). Whole virus ELISA plates were incubated for 16 hours at 4°C. Plates were washed five times with PBST and blocked with 1% bovine serum albumin solution (Jackson ImmunoResearch Inc.) for two hours at 25°C. Samples were diluted 1:50 and a total 100 µL per well were then incubated for one hour at 37°C followed by five washes with PBST. Reaction was revealed by adding 1:10,000 89
dilution of peroxidase conjugated goat anti-mouse IgG (Fc) antibody (Pierce Protein Biology) for
30 minutes at 37°C. Addition of 100 µL per well of TMB allowed for visualization of the peroxidase reaction. Reactions were stopped with 100uL Nova-stop after 5 minutes. Optical density was read at 450 nm.
VP1 peptide synthesis and indirect VP1 peptide ELISA
Eighteen overlapping 20-mer peptides shifted by five amino acids, spanning the complete
SVA-VP1 amino acid sequence, were synthesized (Novoprotein Scientific INC., Summit, NJ) and conjugated to keyhole limpet hemocyanin (KLH) at the N-terminus and hydroxyl at the C-terminus
(Table 1). Reactivity of synthetic peptides was measured by two SVA-pAbs and three VP1-mAbs.
Each peptide was thawed and diluted in phosphate buffered saline (PBS) (Life Technologies, CA) to a concentration of 1 µg/mL. One-hundred µL was added to high binding plate (Thermo
Scientific, MA) and incubated sixteen hours at 4°C. After incubation, plates were washed five times with PBST using an automated plate washer (BioTek,Winooski, VT). Plates were blocked with 300 µL of bovine serum albumin (BSA) for two hours at room temperature (RT) and dried for 4 h at 37°C.
SVA pAbs were diluted in newborn calf serum-based diluent via two-fold dilution from
1:50 to 1:6400. One-hundred µL of two-fold serial dilutions of SVA pAbs, or 1:10 dilution of rVP1 mAbs were incubated in 96 well plates. One-hundred µL of goat anti-mouse IgG Fc conjugated with horseradish peroxidase (HRP) (Pierce Protein Biology) at 1:10,000 was incubated for 1 h at 37°C followed by five washes with 1 x PBS. One-hundred µL of tetramethylbenzidine- hydrogen peroxide substrate (Surmodics) was incubated for 10 min at RT, and enzyme-substrate reaction was stopped with 100 µL Nova-stop (Surmodics). The reaction was quantified by 90
detection of absorbance at 450nm on an automated plate reader (BioTek). As negative control of
100 µL of pre-immunization mouse serum at 1:100 dilution was incubated with each peptide.
Antibody affinity against SVA-rVP1 in blocking ELISA
Antibody affinity selection of SVA pAb, mAb, and swine clinical samples were evaluated by SVA-rVP1 blocking ELISA. SVA pAb, mAb, and pre-immunization mouse serum were diluted in newborn calf-serum diluent. SVA-rVP1 protein was diluted to a concentration of 0.1, 1, 10, 25,
50, and 100 µg/mL. SVA-rVP1 was added to serum in a 1:1 volume ratio and incubated for one hour at 37°C, then loaded into high-binding plates coated in SVA-rVP1 protein as described for
SVA-rVP1 indirect ELISA. After one hour at 37°C, plates were washed and goat anti-mouse IgG
(Fc) HRP (Pierce Biology) was added for one hour at 37°C. Plates were again washed and TMB was added for five minutes before stopping the reaction and reading optical density at 450 nm.
This procedure was repeated using serum from five sows testing positive onSVA-rVP1 ELISA in a 2015 clinical outbreak (4). Samples and SVA-rVP1 were diluted in a fish serum diluent. A goat anti-swine IgG Fc HRP (Bethyl Laboratories, Montgomery, TX, USA) conjugate was used. Serum from five specific pathogen free pigs were run as negative controls.
SVA Peptide Blocking ELISA
SVA VP1 peptide affinity was also evaluated by SVA peptide blocking ELISA. Each of the eighteen peptides was diluted in distilled deionized water to a concentration of 0.2, 2, 20, 50,
100, and 200 µg/mL. An equal volume of mAb 4D10 at 5.75 x 10-5 µg/µL was added to each sample and incubated at 37°C for one hour with shaking. Sample was loaded into SVA-rVP1 coated plates for one hour at 37°C. After washing five times with PBST, goat anti-mouse IgG Fc 91
HRP (Pierce Biology) was added for one hour at 37°C. Plates were washed again and substrate added. After stopping the enzyme-substrate reaction, plates were read at 450nm. Blocking ELISA was repeated with pAb 769L. A blocking SVA-rVP1 ELISA was run using serum collected from five sows testing SVA-rVP1 ELISA positive in a 2015 clinical outbreak (4). Samples were diluted
1:50 in a fish serum-based diluent, then incubated for one hour at 37°C with an equal volume of each peptide diluted in a fish serum diluent to 0.2, 2, 20, 50, 100, and 200 µg/mL. The conjugate used for these samples was goat anti-swine IgG Fc HRP (Bethyl Laboratories, Montgomery, TX,
USA) conjugate. Serum from five specific pathogen free pigs were run as negative controls.
Western Blot
Reactivity of both SVA pAbs and rVP1 mAbs against whole virus particles was evaluated
6 in six serial dilutions of a 1x10 TCID50/mL SVA. Reactivity against SVA-rVP1 was evaluated against both antibodies in six different dilutions of SVA-rVP1. Both SVA particles and rVP1 were mixed in Laemmli loading buffer (Bio-Rad, Hercules, CA, USA) with 5% 2-mercaptoethanol
(Sigma Aldrich), denatured for five minutes at 100°C and loaded into an 8-16% Tris Glycine gel
(Bio-Rad). Proteins were transferred to nitrocellulose membranes (Bio-Rad) utilizing the mini protean transfer system from Bio-Rad using TGX buffer with 20% methanol. The membranes were then blocked for 1 hour at room temperature with Licor blocking buffer containing 0.01% Tween-
20 (LBT) and washed twice with PBST. SVA pAbs and SVA-rVP1 mAbs were diluted in LBT at
1:10,000 and 1:500 respectively and incubated for one hour at room temperature. The membranes were washed four times with PBST prior to incubation with Alexa Fluor 790 goat anti-mouse IgG
H+L (InvitrogenTM, Carlsbad, CA, USA) at 1:5,000 in LBT for one hour at room temperature.
Blots were read using an Odyssey Infrared Imaging System (LICOR Biosciences, Lincoln, NE). 92
Indirect immunofluorescence assay (IFA)
NCI-H1299 cells were grown in 96-well plates. Each well was inoculated with 1x103
TCID50/mL SVA. After 24 hours, cells were fixed with 200 µL cold 90% acetone for 10 minutes at 4°C. Cells were washed with 200 µL PBS, then incubated with 50 µL two-fold serial dilution mouse serum at 37°C for one hour. Wells were washed three times with 200 µL PBS and incubated with 50 µL of FITC labeled anti-mouse IgG (KPL Scientific, Montreal, Quebec, CAN), diluted
1:100, at 37°C for 30 minutes. Plates were washed three times with PBS and examined under a fluorescence microscope.
Virus Neutralization Assay (VN)
Virus neutralization assay was performed using serum samples that were previously heat inactivated at 56°C for 30 minutes. mAb and pAb were not heat inactivated. The sera were two- fold serially diluted (1:20 to 1:40,960) in RPMI 1640 (25 µl/well) in 96-well plate. An equal
2 volume of 1x10 TCID50/mL SVA was added to the mouse sera. The plate was incubated for one hour at 37°C, then the 50 µl serum-virus mixture was inoculated into NCI-H1299 cells in a 96- well plate. The plate was incubated at 37°C for 24 hours and then fixed with 90% acetone. The infection of the cells was checked by SVA IFA described above. The virus neutralization titer was determined as the highest serum dilution where IFA was negative. Each VN assay was performed in duplicate.
Fluorescent Focus Neutralization (FFN)
NCI-H1299 cells (ATCC CRL-5803) were cultured in 96 well flat bottom black tissue culture treated microplates (Corning) in RPMI-1640 media containing 1% Penicillin-Streptomycin 93
and 5% FBS at 37°C and 5% CO2 for 24 hours. Serum samples were heat inactivated for 30 minutes at 56°C. Monoclonal and polyclonal antibodies were not heat inactivated. In a dilution plate, a two-fold dilution from 1:20 to 1:40,960 was completed for all samples and controls. SVA stock was diluted to TCID50 of 10,000 in RMPI. One-hundred µL of virus was added to 100 µL diluted sample and incubated at 37°C for one hour. Next, the virus-sample combination was added to cells, and incubated 1.5 hours at 37°C and 5% CO2. The test plate was washed, then inoculation media added for 24 hours at 37°C and 5% CO2. Media was then removed, and cells fixed with 50
µL of 80% acetone for 15 minutes at room temperature. After removing acetone, plates were dried for 30 minutes. SVA VP1 monoclonal antibody was diluted 1:200 in PBS with 0.1% BSA, and 50
µL added per well for 1 hour at 37°C. Plates were washed three times with PBS with 0.1% BSA, then 50 µL of FITC-labeled goat anti-mouse IgG (KPL Scientific) was added for 45 minutes at
37°C. Plates were washed and PBS added to each well. Finally, plates were read using a Spectra
Max i3x cytometer using SoftMax Pro software (6.5) at 456 nm excitation and 541 nm emission, reading one site per well.
Statistical Analysis
All experiments were performed in duplicate or triplicate technical replicates and analyzed using GraphPad Prism Version 6. Mean and standard deviation were charted. In characterizing generated antibodies, a two-way ANOVA with Tukey corrections made post-test for multiple comparisons was used to determine significance of results in comparison to a negative control. An unpaired t-test with no correction for multiple comparisons was used to evaluate results between
ELISA tests for each antibody. An alpha level of 0.05 was used for all analyses. A two-tailed, nonparametric t-test was used for analysis of IFA and VN. The epitope ELISA was analyzed using 94
an unpaired t-test with no corrections for multiple comparisons. For blocking ELISAs, inhibitory concentration (IC50) was used. A curve was generated using degradation of positive controls.
Results
SVA-rVP1 mAbs generated in vitro do not recognized whole virus particles
Binding activity of two SVA pAbs generated against the whole virus particle and three mAbs generated against a recombinant VP1 protein were evaluated by a whole virus ELISA, SVA- rVP1 ELISA, and western blot. No significant difference in binding activity against rVP1 protein was observed amongst SVA mAbs or SVA pAbs in a SVA-rVP1 ELISA (Figure 1). Both SVA pAbs showed significant binding activity against SVA viral particles in a WV ELISA; however, none of the SVA mAbs showed significant reactivity compared with negative controls (Figure 1).
The binding activity of SVA pAbs and mAbs against SVA WV particles and SVA- rVP1protein was further confirmed by western blot. Serial dilution of detergent treated virus
(molecular weight 65 kDa) and serial dilutions of SVA rVP1 protein (molecular weight 31 kDa) were incubated with two SVA pAbs. SVA pAbs binding activity against SVA WV particles was detectable to a minimum of concentration of 104 virus particles (Figure 2A) and a minimum concentration of 50 ng of SVA rVP1 protein (Figure 2B). All mAbs showed strong reactivity against SVA rVP1 protein (Figure 2C); however, no reactivity was observed against SVA WV particles (figure not shown).
SVA VP1 mAbs and SVA pAbs can detect virus replication by immunofluorescence assay
An indirect immunofluorescence assay showed that both SVA-rVP1 mAb and pAb were able to detect SVA infection in NCI-H1299 cells. No significant difference in IFA titers between 95
SVA mAb and pAb was observed (Figure 3A). Both antibodies showed strong cytoplasmic fluorescent signal (Figure 3B).
SVA pAbs but not SVA-rVP1 mAbs present neutralizing activity in vitro
Viral neutralizing activity of SVA pAb and SVA rVP1-mAb was evaluated in vitro. Serial dilutions of SVA pAb 769L and SVA mAb 4D10 were incubated with SVA and neutralizing activity was evaluated by IFA. SVA pAb showed neutralizing activity to a 1:2,560 dilution, while no neutralizing activity was observed with mAb (Figure 4). In addition, five clinical swine samples positive by SVA-rVP1 ELISA showed strong neutralization titers varying from 1:80 to 1:2,560
(data not shown). No detectable neutralizing activity was observed in five specific pathogen free pig samples used as negative controls.
SVA mAb but not SVA pAb show differential binding activity against SVA-VP1 epitopes by indirect ELISA epitope mapping
To localize linear B-cell epitopes of SVA VP1 involved in immunogenicity, the binding activity of SVA pAb and mAb was evaluated against sequential, overlapping peptides of the SVA- rVP1 protein by indirect peptide ELISA. SVA pAb showed significant reactivity against all peptides spanning the entire SVA VP1 protein compared to a negative control. In addition, no significant differences in intensity were observed among peptides by SVA pAb detection (Figure
5A). However, SVA mAb was capable of detecting significantly higher signal from only four non- overlapping amino acid residues spanning amino acids 1-20, 45-140, 150-170, and 180-230. The peptide spanning amino acid 1-20 showed a particularly high and significant reactivity compared to the peptides spanning amino acids 45-140, 150-170, and 180-230 (Figure 5B). 96
In order to rule out potential cross reactivity between the SVA VP1 peptides used in this study and other members of the family Picornaviridae, a phylogenetic analysis using a NCBI
BLAST (version 2.5.0) was performed. No sequence homologies were seen between other members of the Picornavirus family and the eighteen SVA rVP1 peptides.
A group of SVA VP1 linear epitopes shows a differential inhibitory effect on SVA mAb but not in SVA pAbs against rVP1 protein
Specific peptide inhibitory activity on SVA pAb and SVA mAb against SVA rVP1 protein was evaluated by blocking ELISA. Ten log dilution concentrations of each peptide were incubated for 1 hour either with SVA pAb or mAb. Then, inhibitory binding concentration 50 (IC50) against
SVA rVP1 protein was evaluated by SVA rVP1 ELISA. As a positive control, inhibitory activity of the whole rVP1 protein was also evaluated. Pre-incubation of rVP1 protein with SVA mAbs resulted in significant binding inhibition in a dose-dependent manner, while SVA pAbs showed a minimal reduction in binding activity against rVP1 protein. When SVA pAb binding activity against rVP1 was evaluated after individual peptide blockage, all epitopes showed partial binding inhibitory effect. However, none of the peptides demonstrated a 50% or greater inhibitory effect on SVA pAb binding activity to SVA rVP1 protein (Figure 6A). All eighteen peptides also possessed some ability to block binding activity of SVA mAb to SVA-rVP1. However, four peptides spanning amino acids 1-20, 60-80, 105-125, and 120-140 of SVA-rVP1 showed potent inhibition, reducing SVA mAb binding activity to rVP1 protein by more than 50% (Figure 6B).
97
Anatomic location of reactive epitopes in SVA-rVP1 protein
Peptide interaction and availability was predicted using SWISS MODEL (Figure 7).
Peptide 1, spanning the first twenty amino acids of the VP1 protein, had high substrate availability as was highly exposed in the individual protein.
Discussion
The VP1 protein of SVA is a structural protein that has been used in diagnostic testing, for
ELISA and PCR (21, 22), due to its highly conserved sequence, immunogenicity, and virus specificity. While the other capsid proteins are highly conserved among picornaviruses, VP1 is unique to the Senecavirus genera and involved in cell tropism (2). This study aims to examine the role of linear epitopes of the VP1 protein in the SVA humoral immune response.
SVA pAb and SVA rVP1 mAb characterization showed that SVA rVP1-mAb did not react against a whole virus on both ELISA and western blot. It is possible that conformational epitopes are responsible for this interaction. Therefore, differences in folding and interaction among conformational epitopes in whole virus particles may mask specific binding sites needed for SVA
VP1 mAb antigen interaction. In addition, this lack of reactivity against whole virus particles could be due to differences in antibody affinity due structural differences in protein folding of the rVP1 compared with proteins naturally expressed during viral replication in live cells (23). SVA VP1 is part of a polyprotein, and interaction with other areas of the polyprotein may be vital for viral infection and replication. In foot and mouth disease virus, the GH loop demonstrates flexibility and an ability fold in different configurations (24). The hydrophobic pocket of VP1 has been shown in other picornaviruses to be related to capsid stability and genome release. This could be a 98
contributing aspect to the immunogenicity observed for SVA VP1. The pocket is located under a canyon in the protein structure and its entrance is almost entirely sealed by other residues of VP1
(2, 25). In other viruses, mAb for VP1 has been shown to bind deep in the canyon and have more than one region of receptor binding (26). The SVA-rVP1 could possess differences in this structural pocket, due to folding changes, that prevent natural epitopes from being exposed. Thus, a combination of these VP1 protein structural features could contribute to the difference in SVA mAbs binding activity to whole virus particles.
Although SVA rVP1-mAbs were not capable of detecting WV particles by WV ELISA and western blot, both SVA pAb and mAb recognized SVA replication through immunofluorescence staining, with virus localized mainly in the cell cytoplasm. This difference may be explained by the biological virus-cell interaction allowing exposure of natural epitopes during viral replication. Whole virus ELISA and western blot depend on virus interaction with the plate or viral degradation through detergent treatment to allow conformational changes in regions necessary for antibody-antigen interaction (27, 28). SVA and other picornaviruses have internal ribosomal entry sights and other antigenic regions that must be post-translationally modified by cellular proteases to reach an active configuration (29). Without cellular interaction, proteins expressed in the virus particles are not post-translationally modified and would not express the full repertoire of specific, active binding sites necessary for antibodies generated against VP1 protein.
Antibody generated against whole viral particles demonstrated an ability to neutralize
SVA, while no neutralizing activity was observed with SVA rVP1-mAbs. All monoclonal antibodies evaluated in this study were IgG isotype. Previous experiments in swine demonstrated that neutralizing activity is likely related to the presence of IgM (30). In addition, the same study 99
also showed an earlier and higher neutralizing activity against VP2 compared with VP1 antibodies.
In this study, serum collected from clinically affected sows showed also neutralizing activity, which might suggest that that VP1 is not sufficient to generate antibody protection, and interaction with multiple structural proteins expressed during SVA replication might be necessary.
The SVA VP1 epitope, spanning amino acids 1-20 of the SVA-rVP1 protein, showed a high reactivity and was capable of inhibiting interaction between SVA mAb and SVA rVP1. These results suggest that this linear region of SVA-VP1 plays an important role in the humoral immune response. A SWISS-MODEL of the VP1 protein showed that this region, residues 1-20, is highly exposed when the protein is examined alone. This vulnerable position may contribute to the ability of this amino acid region to induce or be part of induction of an IgG response. However, the spatial interaction of this amino acid region, as well as the whole VP1 protein with the remaining capsid proteins, must be considered to better understand the exposure and interactions of this epitope.
While other peptides showed variable binding activity and inhibitory effects on SVA mAb interaction with rVP1 protein, peptides spanning amino acid 60-80, 105-125, and 120-140 were capable of significantly inhibiting mAb binding activity to SVA-rVP1. These regions may act as additional linear epitopes capable of inducing a humoral immune response. The twenty amino acid segments generated in this study were all recognized by SVA pAb, but could not alone interfere with its interaction with the viral protein. None of the peptides evaluated were able to significantly reduce binding activity of SVA pAb or swine serum against rVP1 protein. It should be noted that the swine serum was collected from sows during a clinical outbreak, while mouse serum was collected from experimentally inoculated animals boostered with a specific, known dose of virus at specific time intervals. Thus, differences observed in clinical samples could be associated with 100
time and dose of exposure. From both, it can be concluded that epitopes of rVP1 alone cannot induce antibody inhibitory effect in SVA pAb generated during experimental or natural infection.
Other protein interactions are necessary to drive an inhibitory humoral response (23).
Recent studies have shown that SVA interacts with cells through the anthrax toxin receptor
1 (ANTXR1), also known as tumor endothelial marker-8 (TEM8). This receptor is upregulated on tumor cells, which may explain the success of SVA as an oncolytic agent in vitro. While no work has been done in vivo or with swine cells, SVA has shown a high affinity for this receptor and some regions of the capsid proteins have been identified as putative areas of interaction with
ANTXR1. Cryo-electron microscopy single-particle analysis showed ANTXR1 amino acid interaction with the BC loop (residues 48-72), loop II (92-107), and FMDV loop (183-213) of
VP1, as well as the puff of VP2 (170-198) and the knob of VP3 (55-71). Additionally the BC loop and loop II formed hydrogen bonds with the receptor (19). Inhibitory activity of SVA mAb against rVP1 induced by epitopes spanning amino acid residues 60-80 and 105-125 might reflect the binding effect observed by ANTXR1 to BC loop (residues 48-72) and loop II (residues 92-107).
It is important to note that the BC loop and loop II of VP1 interact with the puff of VP2 to form the maximum capsid diameter, 325 Å, at the five-fold axis of the capsid (2). This elevated exposure may contribute to reactivity with host cells. Additionally, binding resulted in minor changes in capsid conformation (19), indicating that models and bioinformatics can give only a limited amount of information.
Zhao et al. utilized bioinformatics to predict antigenic epitopes of the VP1 protein to evaluate SVA evolution. Their analyses predicted multiple B and T cell epitopes on the VP1 protein. Of the nine predicted B cell epitopes, residues 1-6, 14-18, 57-63, and 144-148 were 101
included. These regions align with three of the areas identified in our swine and mouse-based mapping study. Software also predicted that secondary structures were of high importance for immune recognition (31). These regions may be involved in conformational epitopes.
This study focused on the role of VP1 protein in the host humoral immune response, particularly IgG. It is important to acknowledge to role of IgM, which has been shown to be highly mediated by the VP2 and VP3 proteins. Additionally, while the humoral response is required to control picornavirus infection through limiting viremia, tissue spread, and severity, the cellular immune response also plays a role in virus defense. Flow cytometry studies showed that the external capsid is essential for CD4+ T cell response and early neutralizing antibody production
(30). More information is necessary to determine if this set of linear epitopes with inhibitory activity within VP1 protein can act as decoy epitopes, delaying antibody neutralizing activity against SVA. Assessment of these linear regions forming conformational epitopes may provide further information regarding their potential role in the humoral response during SVA infection.
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105
Figure Legends
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Figure 1- SVA pAbs and mAbs characterization by SVA WV and SVA-rVP1 ELISA.
Reactivity against SVA recombinant VP1 protein was seen between both pAbs and all three mAbs on SVA-rVP1 ELISA when compared to a pre-injection negative control. Significant reactivity against SVA WV particles on WV ELISA was only with SVA pAbs and no significant reactivity was observed with any of the SVA rVP1-mAbs. Values represent mean and SD of three technical replicates. After completion of ELISA characterization, pAb 769L and mAb 4D10 were selected to be representative of the generated antibodies. 106
Figure 2- Characterization of SVA pAbs and mAbs by western blot. Western blot was completed using a titrated concentration of detergent treated SVA WV particles and SVA VP1 recombinant protein. SVA pAb 769L showed reactivity against WV particles (2A) and producing one band correlating to the molecular weight of SVA-VP1 (2B). SVA mAbs showed binding activity against the SAV-rVP1 (2C), but not SVA WV particles (figure not shown).
107
Figure 3- SVA pAbs and mAbs viral replication detection by IFA. SVA replication in NCI-
H1299 cells was evaluated by IFA with SVA pAbs and mAbs. Both antibodies were capable of specifically detecting SVA replication after 24 hpi. No significant difference in viral detection was observed between SVA pAb 769L and SVA mAb 4D10 (3A). Both antibodies demonstrated a high intracytoplasmic viral replication detection (3B). Values represent mean and SD of three technical replicates.
108
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Figure 4- SVA pAb and SVA rVP1 mAb neutralizing activity detected by IFA. Significant differences were observed between viral neutralizing activity of SVA pAb and SVA rVP1-mAb.
Both mAb 4D10 and pre-immunized serum negative control did not showed neutralizing activity against SVA, while SVA pAb was capable of neutralizing virus. Values represent mean and SD of three technical replicates.
109
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Figure 5- SVA VP1 epitope mapping evaluated by indirect epitope ELISA. SVA-VP1 epitopes were developed and utilized in indirect ELISA. pAb showed reactivity against all eighteen epitopes
(5A). High reactivity was seen with mAb against peptides 1-20 of VP1 on indirect ELISA (5B).
Values represent OD means and SE of three technical replicates.
110
Figure 6 - SVA VP1 epitopes inhibitory effect on SVA mAb and SVA pAb binding activity to rVP1 protein by blocking ELISA. All epitopes showed a partial inhibitory effect on binding; however, none of peptides demonstrated a 50% or greater inhibition effect on SVA pAb binding activity to SVA protein (6A). All eighteen peptides also possessed some ability to block binding activity of SVA mAb with SVA-rVP1 (7B). However, four peptides spanning amino acids 1-20,
60-80, 105-125, and 120-140 of SVA-rVP1 showed potent inhibition, with SVA mAb to rVP1 protein reduced by more than 50% (6 B, C). 111
Figure 7- SWISS MODEL was used to generate a model of SVA-rVP1.
112
Tables
Table 1: VP1 epitope regions. Eighteen peptides spanning the length of SVA-rVP1 were generated. Peptides were 20 amino acids and length and overlapping by five peptides. Peptide purity was at 85% and all peptides except SVA-VP1-PEP5, 8, and 17 were optimized through addition of cysteine.
Name of peptide Sequence of peptide Position of amino acid Length (aa) SVA-VP1-PEP1 STDNAETGVIEAGNTDTDFS 1-20 20 SVA-VP1-PEP 2 DTDFSGELAAPGSNHTNVKF 15-35 20 SVA-VP1-PEP 3 TNVKFLFDRSRLLNVIKVLE 30-50 20 SVA-VP1-PEP 4 IKVLEKDAVFPRPFPTQEGA 45-65 20 SVA-VP1-PEP 5 TQEGAQQDDGYFCLLTPRPT 60-80 20 SVA-VP1-PEP 6 TPRPTVASRPATRFGLYANP 75-95 20 SVA-VP1-PEP 7 LYANPSGSGVLANTSLDFNF 90-110 20 SVA-VP1-PEP 8 LDFNFYSLACFTYFRSDLEV 105-125 20 SVA-VP1-PEP 9 SDLEVTVVSLEPDLEFAVGW 120-140 20 SVA-VP1-PEP 10 FAVGWFPSGSEYQASSFVYD 135-155 20 SVA-VP1-PEP 11 SFVYDQLHVPFHFTGRTPRA 150-170 20 SVA-VP1-PEP 12 RTPRAFASKGGKVSFVLPWN 165-185 20 SVA-VP1-PEP 13 VLPWNSVSSVLPVRWGGASK 180-200 20 SVA-VP1-PEP 14 GGASKLSSATRGLPAHADWG 195-215 20 SVA-VP1-PEP 15 HADWGTIYAFVPRPNEKKST 210-230 20 SVA-VP1-PEP 16 EKKSTAVKHVAVYIRYKNAR 225-245 20 SVA-VP1-PEP 17 YKNARAWCPSMLPFRSYKQK 240-260 20 SVA-VP1-PEP 18 SYKQKMLMQ 255-264 9
113
CHAPTER 5: GENERAL CONCLUSION
Summary
SVA is a single-strand, positive sense RNA virus in the family Picornaviridae that has recently been associated with outbreaks of vesicular disease and epidemic transient neonatal losses in swine. Such outbreaks have been described in multiple countries including the United States,
Brazil, Canada, China, Vietnam, Colombia, and Thailand. Clinical signs of SVA infection have only been described in swine and cannot be from other vesicular diseases including FMDV, VES,
VS, and SVD. Diagnosis is achieved through multiple direct and indirect methods including PCR,
VN, ISH, ELISA, and IFA. After natural exposure, the virus produces viral shedding and a short viremia. Viral particles are detectable during a short viremia and in the feces of clinically affected animals, as well as those without clinical signs. Additionally, clinically healthy animals at the site out herd outbreaks have been shown to develop an IgG response, suggestive a subclinical presentation. More information is needed regarding the pathogenesis and transmission of SVA in order to develop sound protocols for biosecurity, vaccination, and procedures during outbreaks.
Serological studies show that the proportion of seropositive animals in SVA positive farms did not differ between clinically and non-clinically affected animals. However, there is not information available regarding the seroprevalence on United States’ farms without clinical IVD or ETNL associated with SVA. Here we determined the seroprevalence of SVA in commercial swine in the USA. A total of 2,433 sows and 3,654 grower/finisher serum samples were collected from farms with no history of IVD, ETNL, and SVA diagnosis during 2016 at ISU-VDL. Thirty animals were selected from each unique site n=219 (n=88 sows; n=131 grower-finishers) representing 18 USA states. Sera were tested using SVA-rVP1 ELISA and indirect 114
immunofluorescence assay (IFA). The overall seroprevalence detected by rVP1-ELISA was
26.2% (CI ±1.76) and 8.7% (CI ±0.92), and by IFA was 15.9% (CI ±1.46) and 5.7% (CI ±0.75) for sows and grower-finishers, respectively. The between farm prevalence was 75.8% (CI ±8.8) and 42.7 % (CI ±8.5) for sow and grower-fisher herds. The within herd prevalence for sows had the following distribution: 58.2% of sites had 1-50% seropositive rates, 10.98% ranging from 51-
70%, and 18.68% between 71-100%. For grower-finisher premises, 70.8% of farmed fell into the
1-50% seropositive range, 5.5% from 51-70%, and 1.8% in 71-100%. Clinically healthy sows and grower-finisher pigs possessed antibody against SVA with variable within herd prevalence. The presence of antibodies in grower-finishers indicates that animals are subclinically infected during the grower-finisher stage or lesions are going undetected. These results strongly suggest that SVA is circulating subclinically in United States’ swine population.
Murine models have been described for the study of many members of the family
Picornaviridae. Previous studies with foot and mouth disease virus have shown that mouse models can reproduce vesicular disease and mortality. However, these clinical outcomes appear to be dependent on mouse strain and age(1)(1)(1). In this study, five different mouse strains including
BALB/C, C57BL/6, SJL/J, Swiss, and SCID showed detectable viral shedding during the 14 day study, in addition to viral detection in multiple tissues. All mouse strains, except for SCID mice, developed a strong humoral immune response by 14 dpi. Despite this strong evidence of viral replication and systemic spreading of the virus, we did not observed lethality or significant clinical signs of systemic illness. In addition, gross and histological evaluation showed that SVA does not induce cutaneous vesicles or significant lesions in other tissues at 14 dpi. Based on these results, we suggest that SVA can produce subclinical infection in multiple strains of laboratory mice by 115
subcutaneous inoculation. No clinical differences were observed among the different infectious routes in this study. IP and SC inoculation appeared to cause subclinical infection. In both groups there was acute viremia and viral shedding in feces detected within three dpi. Unlike pigs, fighting and grooming play a large role in the spread of viral disease in mice. Our results suggest that SC and IP route better mimic the natural exposures that would occur through biting, grooming, and physical contact than IN route. Swine appear to be the only natural host for SVA; however, retrospective studies demonstrated the presence of SVA neutralizing antibodies in mice and cattle.
In the current study, we evaluated the presence and dynamic of the IgG antibody response in different infectious routes. The production of IgG antibody after SVA infection in mice is acute, with antibodies levels detectable after 7 dpi. This rapid development of specific humoral response is consistent with results observed in experimentally inoculated pigs. Mice may play a role in the transmission of SVA as a vector or reservoir for disease, allowing for virus to spread from one swine site to another through shedding in the feces or urine, or through direct contact. As wild mice as widespread across the United States and within production animal housing, viral transmission and movement of mice would allow SVA to spread over a wide geographic distance.
Risk of SVA infection in mice may be decreased through increased pest management and biosecurity.
The VP1 protein of SVA is a highly conserved and immunogenic structural protein. While the other capsid proteins are highly conserved among picornaviruses, VP1 is unique to the
Senecavirus genera and involved in cell tropism. This study aims to examine the role of linear epitopes of the VP1 protein in the SVA humoral immune response. Immunogenic regions of the
SVA-VP1 protein were examined through epitope mapping. The binding activity of SVA pAbs and mAbs were evaluated eighteen epitopes of the rVP1 protein. Epitopes were twenty amino acids 116
in length with a 5-mer overlap. While other peptides showed variable binding activity and inhibitory effects on SVA mAb interaction with rVP1 protein, peptides spanning amino acid 60-
80, 105-125, and 120-140 were capable of significantly inhibiting mAb binding activity to SVA- rVP1. Additionally, the epitopes spanning aa 60-80 and 105-125 lie on the BC loop and loop II of the VP1 protein, respectively. Cryo-electron microscopy has shown that these regions interact with the ANTXR1 receptor in vitro. Our results suggest that a set of four linear epitopes of the VP1 protein are responsible for IgG humoral response against VP1 during SVA infection.. Additional work is warranted to determine the role of these regions as in their conformational, native structure; their interaction with other immunogenic regions of the VP1 protein and other structural proteins; and their ability to induce a neutralizing antibody response.
Conclusions
SVA appears to be circulating subclinically in healthy swine herds distributed across the
United States, with seropositive animals detected in both breeding and grower/finisher herds.
Laboratory mice undergo subclinical SVA infection and are capable of shedding viral particles in the feces. Wild mice could act as a source of transmission between swine herds as a reservoir host or vector. SVA VP1 protein plays a role in the humoral response of the host driven by a set of linear epitopes. However, these linear epitopes are not sufficient to confer protection and may need to be expressed in their native structure or interact with other immunogenic regions of the other structural proteins.