Persistent Infection and Transmission of Senecavirus A from Carrier Sows to Contact Piglets

Abstract

Senecavirus A (SVA) is a picornavirus that causes acute vesicular disease (VD), that is clinically indistinguishable from foot-and-mouth disease (FMD), in pigs. Notably, SVA RNA has been detected in lymphoid tissues of infected animals several weeks following resolution of the clinical disease, suggesting that the virus may persist in select host tissues. Here, we investigated the occurrence of persistent SVA infection and the contribution of stressors (transportation, immunosuppression, or parturition) to acute disease and recrudescence from persistent SVA infection. Our results show that transportation stress leads to a slight increase in disease severity following infection. During persistence, transportation, immunosuppression, and parturition stressors did not lead to overt/recrudescent clinical disease, but intermittent viremia and virus shedding were detected up to day 60 postinfection (p.i.) in all treatment groups following stress stimulation. Notably, real-time PCR and in situ hybridization (ISH) assays confirmed that the tonsil harbors SVA RNA during the persistent phase of infection. Immunofluorescence assays (IFA) specific for double-stranded RNA (dsRNA) demonstrated the presence of double-stranded viral RNA in tonsillar cells. Most importantly, infectious SVA was isolated from the tonsil of two animals on day 60 p.i., confirming the occurrence of carrier animals following SVA infection. These findings were supported by the fact that contact piglets (11/44) born to persistently infected sows were infected by SVA, demonstrating successful transmission of the virus from carrier sows to contact piglets. Results here confirm the establishment of persistent infection by SVA and demonstrate successful transmission of the virus from persistently infected animals.IMPORTANCE Persistent viral infections have significant implications for disease control strategies. Previous studies demonstrated the persistence of SVA RNA in the tonsil of experimentally or naturally infected animals long after resolution of the clinical disease. Here, we showed that SVA establishes persistent infection in SVA-infected animals, with the tonsil serving as one of the sites of virus persistence. Importantly, persistently infected carrier animals shedding SVA in oral and nasal secretions or feces can serve as sources of infection to other susceptible animals, as evidenced by successful transmission of SVA from persistently infected sows to contact piglets. These findings unveil an important aspect of SVA infection biology, suggesting that persistently infected pigs may function as reservoirs for SVA.

Keywords: SVA; SVV; Seneca Valley virus; carrier; persistence.

FIG 1

Experimental design. (A) Acute phase (phase 1): effects of transportation stress on acute SVA-induced VD. Group 1 (G1), mock-infected control group (n = 4); group 2 (G2), transportation stress with SVA infection of sows upon arrival (2-h trip from farm to animal facility; n = 5); and group 3 (G3), SVA-infected sows after acclimation (7-day acclimation/no-stress group; n = 5). (B) Chronic/persistent phase (phase 2): occurrence of persistent infection and effect of stressors on chronic/persistent SVA infection. SVA- or mock-infected sows from the first phase of the study were subjected to transportation stress (2-h trip; G1 and G2) or received immunosuppressive dexamethasone treatment (1.5 mg/kg, i.v.) for five consecutive days (G3). The effects of these stressors on persistent SVA infection were evaluated for 14 days. To assess the effect of the stress associated with parturition, a fourth group (G4) consisting of SVA-infected pregnant gilts was included in the study (parturition stress, n = 5). Gilts in G4 were inoculated with SVA upon arrival at the animal facility on gestational day 68. Parturition was induced on day 44 p.i. (gestational day 112), and its effects on SVA persistent infection were evaluated for 14 days. During the chronic/persistent phase, all four groups (G1 to G4) were monitored in parallel.

FIG 2

Clinical presentation of acute SVA-induced vesicular disease following transportation stress or acclimation. (A) Time to lesion development in animals in G2 (transport stress) and G3 (acclimated). Vesicular lesions were first observed on the feet of SVA-infected animals at day 2 p.i. (top panels) and subsequently on the snout on day 3 p.i. (bottom panels). (B) Severity of vesicular lesions in animals in G2 (transport stress) and G3 (acclimated). Vesicular lesions were detected on the snout of SVA-infected sows starting at day 3 p.i. (transport stress group [G2] and acclimated group [G3]). Overall, lesions observed in G2 transport-stressed animals were more severe than those observed in G3 acclimated animals. (C) Clinical scores. Animals were evaluated daily for the presence of vesicles and lameness. One point was assigned for each foot or snout displaying vesicular lesions, and another point was assigned for lameness (maximum of 6 points/animal/day), and daily group averages were calculated. Arrows indicate vesicular lesions.

FIG 3

Kinetics of viremia and virus shedding during acute SVA infection in sows (0 to 35 dpi). Viremia (A) and virus shedding in oral (B) and nasal (C) secretions and in feces (D) were determined by RT-qPCR quantitated based on a relative quantitation method.

FIG 4

SVA neutralizing antibody levels during acute SVA infection of sows and detection of SVA in tonsil. (A) Virus-neutralizing (VN) antibody responses elicited by SVA infection were evaluated weekly from day 0 to day 35 p.i. NA titers represent the reciprocal of the highest serum dilution capable of completely inhibiting SVA infectivity. The gray line represents the cutoff value of the VN assay. (B) Viral load in the tonsil of SVA-infected animals. To confirm viral presence in the tonsil, one animal from each group was necropsied on day 44 p.i. SVA presence in the tonsil was determined by RT-qPCR, and viral load was determined by a relative quantitation method.

FIG 5

Dexamethasone-induced changes in peripheral white blood cells count in SVA-infected sows. Peripheral white blood cell count and leukocyte discrimination were performed by flow cytometry during SVA infection of control sows (G1) and sows subjected to transport stress (G2), dexamethasone treatment (G3), or parturition stress (G4). (A) A representative flow cytometry gate strategy used to determine white blood cell subpopulations is shown. The plot shows red blood cells, platelets, and cell debris exclusion and leukocyte selection (based on forward scatter area [FSC-A] and side scatter height [SSC-H]; top left plot). A dot plot depicts lymphocyte, monocyte, and granulocyte populations (top right plot). This plot served as confirmatory gates used for selection based on the expression of cell markers (bottom plots). B and T lymphocytes were selected based on the expression of CD21 and CD3, respectively (panel 1 plots). Granulocytes and monocytes were determined by the expression of CD172a, as well as by their differential size and complexity properties (panel 2 plot). (B) Absolute white blood cell counts. (C) Relative granulocyte count (CD172a⁺; high complexity). (D) Relative T cell count (CD3⁺). (E) Relative monocyte count (CD172a⁺; low complexity). (F) Relative B cell count (CD21⁺). Percentages in panels C to F are based on the total WBC count.

FIG 6

Kinetics of viremia and virus shedding during chronic/persistent SVA infection (42 to 60 dpi). Viremia (A) and virus shedding in oral (B) and nasal (C) secretions and in feces (D) were determined by RT-qPCR and quantitated based on a relative quantitation method.

FIG 7

SVA neutralizing antibody levels in serum of SVA-infected animals during chronic/persistent SVA infection. Virus-neutralizing (VN) antibody responses during chronic/persistent SVA infection from day 42 to 60 p.i. NA titers represent the reciprocal of the highest serum dilution capable of completely inhibiting SVA infectivity.

FIG 8

Detection of SVA RNA in the tonsil of SVA-infected animals on day 60 p.i. (A) Viral load in the tonsil of SVA-infected animals, as determined by RT-qPCR and quantitated based on a relative quantitation method. (B) Representative in situ hybridization demonstrating the presence of SVA nucleic acid (in red) in the tonsils of SVA-infected sows on day 60 p.i. G1 control, SVA RNA detected; G2, transport stress; G3, immunosuppression by dexamethasone; and G4, parturition. Animal numbers are indicated. Multifocal staining was observed in tonsillar epithelial (TE) cells, crypt epithelial (CE) cells, subepithelial lymphoid tissue (SLT), and lymphoid follicles (LF).

FIG 9

In vivo detection of viral dsRNA in tonsil of SVA-infected pigs during chronic/persistent infection. Frozen tonsil sections from mock-inoculated (top) or SVA-infected animals at day 60 p.i. dsRNA was stained with a mouse monoclonal antibody specific for viral dsRNA (J2 clone), and the cell nuclei were counterstained with DAPI (right panel; blue). Red fluorescence indicates the presence of dsRNA in the cytoplasm of cells. G1, control animal 979; G2, transport stress animal 1623; G4, parturition stress animal 1807.

FIG 10

Detection of SVA RNA in oro-rectal secretions from piglets born to sows in the parturition stress group (G4). (A) Oro-rectal secretions from piglets born to sows in G4 (5 piglets from each sow were randomly sampled) were collected daily. Levels of SVA RNA were determined by RT-qPCR and quantitated by a relative quantitation method.