Abstract
We determined tissue localization, shedding patterns, virus carriage, antibody response, and aerosol transmission of Porcine epidemic diarrhea virus (PEDV) following inoculation of 4-week-old feeder pigs. Thirty-three pigs were randomly assigned to 1 of 3 groups for the 42-day study: inoculated (group A; n = 23), contact transmission (group B; n = 5), and aerosol transmission (group C; n = 5). Contact transmission occurred rapidly to group B pigs whereas productive aerosol transmission failed to occur to group C pigs. Emesis was the first clinical sign noted at 3 days postinoculation (dpi) followed by mild to moderate diarrhea lasting 5 more days. Real-time PCR detected PEDV in fecal and nasal swabs, oral fluids, serum, and gastrointestinal and lymphoid tissues. Shedding occurred primarily during the first 2 weeks postinoculation, peaking at 5–6 dpi; however, some pigs had PEDV nucleic acid detected in swabs collected at 21 and 28 dpi. Antibody titers were measurable between 14 and 42 dpi. Although feces and intestines collected at 42 dpi were PEDV negative by PCR and immunohistochemistry, respectively, small intestines from 70% of group A pigs were PCR positive. Although disease was relatively mild and transient in this age group, the results demonstrate that 4-week-old pigs are productively infected and can sustain virus replication for several weeks. Long-term shedding of PEDV in subclinically affected pigs should be considered an important source for PEDV transmission.
Introduction
Porcine epidemic diarrhea (PED) emerged as a new swine enteric disease in North America in the spring of 2013. 13 Porcine epidemic diarrhea virus (PEDV; order Nidovirales, family Coronaviridae, subfamily Coronavirinae, genus Alphacoronavirus) is an enveloped, positive-sense RNA virus. 6 Since initial detection, PEDV has spread rapidly across the United States and has now been reported in 39 states (https://goo.gl/7Iyhtw). The mechanisms for introduction and rapid dissemination throughout pig farms in the United States and North America are still poorly understood; although, transportation, feed, and aerosols have all been implicated as potential contributors.1,2,4,8,11 The peak disease period for this virus is in the late fall and early winter months in temperate climates, and clinical disease typically abates in the late spring, summer, and early fall. Infection of neonatal pigs usually results in extremely high mortality (approaching 100%), caused by malabsorptive diarrhea and dehydration as a result of enterocyte necrosis in the small intestine. 9 Infection of grow-finish animals results in high morbidity but low mortality, with vomiting and mild to moderate diarrhea as the typical clinical presentation. Infection of adult swine is often overlooked given minimal gastrointestinal signs or a complete lack of clinical disease. Following the introduction of PEDV in the United States, ~7 million pigs died within the first year, placing pressure on domestic and foreign markets. 6 Even though PED is very similar to the disease presentation caused by transmissible gastroenteritis virus (TGEV), PEDV and TGEV are genetically and immunologically distinct. 7 For the PEDV challenge in our study, 4-week-old pigs were selected as an intermediate age group, wherein clinical disease was expected to be mild to moderate, and low mortality would allow the analysis of viral shedding, tissue distribution, aerosol transmission, and antibody response over the 42-day period.
Materials and methods
Animals
Experiments involving animals and viruses were performed in accordance with the Federation of Animal Science Socie-ties (FASS) Guide for the Care and Use of Agricultural Animals in Research and Teaching, the USDA Animal Welfare Act and Animal Welfare Regulations, and approved by the Kansas State University Institutional Animal Care and Use Committees and Institutional Biosafety Committees. Thirty-three 3-week-old pigs were obtained from a high-health commercial source negative for PEDV. All pigs were housed in a 66-m2 room at the Biosecurity Research Institute at Kansas State University (Manhattan, Kansas), and maintained under Biosafety Level 3 Agriculture (BSL-3Ag) containment conditions. Pigs were housed in 2 raised stainless steel pens (6.9 and 3.5 m2 for groups A and B and group C, respectively) with slotted fiberglass flooring. The 2 pens were ~3 m apart and separated by a nonpermeable plastic tarp to avoid overt cross-contamination during cleaning. The tarp was placed directly between the 2 pens and covered ~60% of the room length. Floors of the room were chemically disinfected daily. a Pigs were given access to food and water ad libitum. Complete exchange of air within the room occurred 14.5 times/h. Room temperature was 22.8–27.8°C, and humidity was 32–45% during the study.
Experimental design
Pigs were randomly assigned to 1 of 3 groups including an inoculated group (group A; n = 23), contact transmission group (group B; n = 5), and aerosol transmission group (group C; n = 5; Table 1). The pigs were given 1 week to acclimate on arrival. At 4 weeks of age, group A pigs were given a PEDV “feedback” inoculum via the oral and intranasal routes, with 5 mL of inoculum per route. The inoculum was gut-derived intestinal contents that had been used as “feedback” inoculum for controlled exposure of a sow herd in a commercial swine production unit. b The PEDV inoculum (isolate USA/KS/2013, GenBank accession KJ184549.1) had a quantification cycle (Cq) of 22 as determined by the Kansas State Veterinary Diagnostic Laboratory (KSVDL) real-time PCR assay. This Cq value is equivalent to 1.42 × 105 TCID50 (50% tissue culture infective dose). Routine PCR testing by KSVDL showed that the inoculum was negative for Porcine reproductive and respiratory syndrome virus (PRRSV), Porcine circovirus 2 (PCV-2), and Rotavirus A, B, and C.
Experimental design for Porcine epidemic diarrhea virus (PEDV) transmission in 4-week-old feeder pigs.
Inoculation via oral and intranasal routes; 5 mL/route.
Comingled with group A 6 h postinoculation.
Housed in a separate pen in the common animal room.
Group A pigs were individually removed from the group pen and inoculated. Group B pigs were not inoculated and were housed separately from group A pigs during inoculation. Approximately 6 h postinoculation of the group A pigs, groups A and B were allowed to comingle and were housed together for the remainder of the study. Group C pigs were not inoculated and were housed in a separate pen in the same animal room as groups A and B (described above). Nasal swabs, fecal swabs, oral fluids, and serum samples were collected from all pigs prior to challenge on day –3 and on 0–7, 9, 14, 21, 28, 35, and 42 days postinoculation (dpi). One pig on 0, 2, and 4 dpi and 2 pigs on 7, 9, 14, 21, and 28 dpi were randomly selected for euthanasia and autopsy. All remaining pigs were euthanized on 42 dpi. To assess the presence of virus in the environment, swabs were collected from the V-troughs used for blood collection, walls, pens, and food bins from both the groups A and B and group C sides of the room on 14 dpi.
Clinical evaluation
Pigs were evaluated daily throughout the study period for the presence of clinical signs associated with PED. During the acute period of infection from 0 to 11 dpi, each pig was physically examined by a veterinarian for clinical signs such as decreased body condition, dehydration, lethargy, and evidence of diarrhea or vomiting. Lethargy was assessed through response to stimuli and interaction with pen mates. Body condition scores were assigned using an adapted 5-point scale. 12 Dehydration was assessed through evaluation of enophthalmos and third eyelid protrusion, nasal planum moisture and coloration, mucous membrane moisture and coloration, and skin turgor. Fecal consistency scores were assigned using a 5-point scale: 0 = no feces; 1 = normal feces; 2 = soft but formed feces; 3 = brown diarrhea with particulate fecal material; 4 = brown diarrhea without particulate fecal material; 5 = clear, watery diarrhea.
Gross anatomic and histologic examination
Pigs were sequentially euthanized throughout the study to assess gross anatomic and microscopic changes during infection. Pigs were euthanized with pentobarbital sodium, autopsied immediately, and the following tissues were collected from each pig: inguinal lymph node, submandibular lymph node, tonsil, thymus, thyroid gland, esophagus, trachea, lung (1 section from each lobe), tracheobronchial lymph node, heart, liver, adrenal gland, kidney, spleen, stomach, mesenteric lymph node, duodenum with pancreas, jejunum (3 locations), ileum, cecum, spiral colon (2 locations), descending colon, nasal turbinates, bone marrow, and brain. Tissues were frozen at −70°C and fixed in 10% neutral buffered formalin. After 7 days, formalin-fixed tissues were routinely processed in an automated tissue processor and embedded in paraffin. Tissue sections were cut at 4 μm, mounted on glass slides, and stained with hematoxylin and eosin (H&E). Autopsies and histologic evaluations of tissues were by a board-certified pathologist.
Polymerase chain reaction
A commercial viral RNA isolation kit c was used together with a magnetic particle processor d for all sample types. Tissue samples were homogenized using a laboratory paddle blender. e One milliliter of 1× phosphate buffered saline buffer was added to ~0.5 g of the feces or a nasal swab tube, vortexed briefly, and allowed to sit for 2–3 min. The supernatant was then used for RNA extraction. For all sample types, 70 μL of liquid was used for RNA extraction. The extracted RNA was frozen at −20°C until analysis by real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR).
A duplex qRT-PCR was designed for the dual purpose of detecting PEDV nucleocapsid (N) protein gene and the host 18S ribosomal RNA subunit to monitor extraction efficiency. Primers and probe sequences for PEDV were designed from the N gene of genome KJ184549.1: PEDVn-F2 (GCTATGCTCAGATCGCCAGT; 27,331-27,350 nt), PEDVn-R2 (TCTCGTAAGAGTCCGCTAGCTC; 27,423-27,402 nt), and PEDVn-Pr2 probe (FAM-TGCTCTTTGGTGGTAATGTGGC-BHQ1; 27,373-27,394 nt). Primers and probe sequences for 18S were: 18S-F (GGAGTATGGTTGCAAAGCTGA), 18S-R (GGTGAGGTTTCCCGTGTTG), and 18S-Pr probe (Cy5-AAGGAATTGACGGAAGGGCA-BHQ2). A commercial RT-PCR kit f was used for all real-time PCR reactions. The qRT-PCR reactions in 20 µL consisted of 1.5 µL of nuclease-free water, 10 µL of 2× reaction buffer, 1 µL of 10 µM PEDVn forward and reverse primers, 1 µL of 10 µM 18S forward and reverse primers, 1 µL of 10 µM 18S probe, 0.5 µL of PEDV probe (10 µM), 1 µL of enzyme mix, f and 4 µL of extracted RNA. Each qRT-PCR reaction plate was run on a thermal cycler g under the following conditions: 48°C for 10 min; 95°C for 10 min; followed by 45 cycles of 95°C for 10 s and 60°C for 40 s. Positive and negative PCR amplification controls and a negative extraction control were included in each run.
Immunohistochemical staining
A primary goal for immunohistochemical (IHC) staining was to determine if virus replication occurred in the respiratory tract as well as in the small intestine. Therefore, tissues from the gastrointestinal and respiratory tracts were prioritized for IHC staining after collection from pigs euthanized between 0 and 28 dpi. Preliminary IHC examination of tissues for PEDV antigen was performed at the Iowa State University Veterinary Diagnostic Laboratory (Ames, Iowa). Subsequently, IHC was performed on these same tissues at the KSVDL, where positive and negative results were confirmed independently.
IHC staining at the KSVDL was performed on formalin-fixed, paraffin-embedded tissues that were sectioned at 4 μm thickness onto positively charged slides. Slides were stained using a commercial autostainer h and detection kit. i The PEDV primary antibody (SD 6-29 j ) was diluted k to 1:100,000. Epitope retrieval was performed by incubating slides in citrate (pH 6.0) for 20 min at 100°C. Tissue sections were incubated with the primary antibody for 15 min at ambient temperature. Polymerization l was performed for 25 min at ambient temperature. Visualization was done with DAB and counterstained with hematoxylin. Positive and negative controls were included with each run.
Antibody response
Serum samples were collected throughout the study and stored at −80°C for antibody testing using 2 methods. First, serially diluted serum samples were assayed for PEDV antibodies using an indirect fluorescent antibody test (IFAT) in a 96-well format. The IFAT antigen was obtained by infecting swine testicular (ST) cells with a standardized stock of PEDV (Colorado 2013 isolate m ). IFAT endpoints were calculated as the reciprocal of the last serum dilution that gave a positive IFAT response when viewed with a fluorescent microscope.
PEDV neutralizing antibody levels were determined using a 96-well microtiter system with African green monkey kidney (Vero) cells as the substrate and a standardized trypsin independent stock of PEDV as the indicator virus. Serial dilutions of serum were mixed with a constant quantity of PEDV (50–300 TCID50), incubated for 1 h at 37°C, and inoculated into 4 replicate wells of 3-day-old Vero cells in 96-well plates. Cultures were incubated for 3 days at 37°C, and the presence of virus was determined by the presence of virus-associated cytopathic effect. Serum neutralization titers were based on 50% inhibition of the indicator virus, and 50% endpoints were then determined by the method of Spearman and Karber. 5
Results
Transmission
Group B pigs rapidly became infected with PEDV after comingling with group A pigs. In contrast, aerosol transmission did not occur as group C pigs failed to develop a productive PEDV infection during the 42-day study period.
Clinical evaluation
Clinical signs associated with PEDV infection occurred in groups A and B between 3 and 8 dpi. Clinical disease correlated well with the time course of peak fecal and nasal PEDV shedding between 4 and 7 dpi (Fig. 1). The first clinical sign associated with PED was emesis noted on 3 dpi. Vomitus contained partially digested food particles without blood or mucus. The most severe diarrhea observed was assigned a score of 4 out of 5 and included brown liquid feces without particulate fecal material, blood, or mucus. This diarrheal score was recorded in 30% of the group A and B pigs between 4 and 7 dpi. Evidence of clinical dehydration was documented in ~70% of group A and B pigs between 4 and 8 dpi. Dehydration was mild to moderate and resulted in pigs having enophthalmos with third eyelid protrusion, dry nasal planums, tacky mucous membranes, and decreased skin turgor. During this time, most pigs also had mild to moderate lethargy, including reluctance to rise and ambulate, decreased resistance to handling and restraint, and reduced responsiveness. Starting on 4 dpi, most pigs in groups A and B (16 of 26) had body condition scores that were less than ideal (<3 of 5). However, body condition scores gradually improved after resolution of clinical disease and, by 11 dpi, most group A and B pigs (13 of 21) had ideal body condition scores (≥3 of 5). No pigs were considered moribund during the study, and there were no mortalities.
Time course of clinical disease and Porcine epidemic diarrhea virus (PEDV) detection in group A pigs after PEDV inoculation. Clinical signs consistent with PEDV were not present after 10 days postinoculation (dpi).
Gross anatomic, histologic, and IHC examination
Except for the mild to moderate dehydration observed clinically, gross lesions were not observed in any of the euthanized pigs. All tissues collected from group A pigs autopsied from 0 to 28 dpi were examined histologically, and significant microscopic changes were present in the small intestines of 3 of 13 pigs. There was diffuse, moderate to severe atrophy and fusion of villi in the jejunum and ileum of the pig autopsied on 4 dpi and in 1 of the 2 pigs autopsied on 7 dpi. Mild to moderate villus atrophy and fusion was present in the jejunum and ileum in 1 of the 2 pigs autopsied on 28 dpi.
IHC staining results from Iowa State University and the KSVDL were identical. PEDV was detected by IHC staining in the small intestines of 6 pigs (Table 2) and in the mesenteric lymph nodes of 3 pigs. All 5 small intestinal samples from the pig euthanized at 4 dpi and in 1 of the 2 pigs euthanized at 7 dpi were positive. One of 5 sections from both pigs euthanized at 14 dpi, and 2 of 5 sections from 1 of the 2 pigs euthanized at 21 and 28 dpi were positive. In the pigs in which all small intestinal sections were positive, PEDV antigen was multifocal in the villus epithelium of the duodenum and diffuse in the villus epithelium of the jejunum and ileum. In pigs in which only a portion of the sections was positive, viral antigen was limited to the jejunum and/or ileum and was absent from the duodenum. In positive sections with Peyer’s patches, positive staining was often present within lymphoid tissue as well as the villus epithelium.
Immunohistochemistry summary of small intestinal sections collected sequentially from group A pigs after Porcine epidemic diarrhea virus (PEDV) inoculation.*
Pigs were randomly selected throughout the study for sequential collection. Five sections of small intestine were collected from each pig and evaluated for the presence of PEDV antigen by immunohistochemical (IHC) staining.
The percent of villus epithelial cells in IHC-positive sections was estimated and assigned scores as follows: + = 0–25% positive; ++ = 26–50% positive; +++ = 51–75% positive; ++++ = 76–100% positive.
On 4 dpi, PEDV antigen was uniformly detected in group A pigs along the epithelial border, associated with villus atrophy and fusion (Fig. 2A). Between 7 and 28 dpi, PEDV antigen became less consistent along the epithelial border (Fig. 2B–2E). Taken together, these data show a high degree of variability in IHC detection of PEDV antigen, particularly after 7 dpi.
Porcine epidemic diarrhea virus (PEDV) immunohistochemistry of jejunal sections from group A pigs sequentially collected after PEDV challenge. Images are shown from tissues collected:
PEDV shedding and viremia
In general, fecal and nasal shedding patterns were very similar between group A and group B pigs (Fig. 3, Supplemental Tables 1 and 2, available at http://vdi.sagepub.com/content/by/supplemental-data). Surprisingly, all samples were negative for the virus at 24 h postinoculation. Fecal and nasal shedding of group A was first observed at 48 h postinoculation. Nasal shedding was detected in the group B pigs at 48 h postinoculation, and fecal shedding was present in all 5 group B pigs by 72 h postinoculation. Peak fecal shedding occurred between 5 and 6 dpi and was significantly greater than nasal shedding in both groups A and B (compare Fig. 3A and 3B). Fecal shedding was typically detected at a 10-fold or greater level than that observed in the nares. In groups A and B, most animals were negative for fecal and nasal shedding by 21 dpi. However, 3 of 14 pigs in group A and 2 of 5 pigs in group B were still shedding virus in feces at 21 dpi. In addition, 1 of 12 pigs from group A was shedding PEDV in feces and nasal secretions at 28 dpi. Although low levels of PEDV nucleic acid were detected in 5 of 5 group C nasal swabs during the early part of the study, the absence of fecal shedding and lack of antibodies suggested that the PEDV detected in nares was noninfectious.
PCR detection of Porcine epidemic diarrhea virus (PEDV) in fecal swabs (
In general, the amount of PEDV nucleic acid was lower in serum compared to fecal and nasal samples (Fig. 3C, Supplemental Table 3, available at http://vdi.sagepub.com/content/by/supplemental-data). Viremia peaked around 5 dpi, similar to fecal and nasal shedding. PEDV viremia was clearly detected in the majority of both the group A and B pigs. Only 3 of 25 animals in groups A and B did not develop detectable viremia within the first week post-inoculation or exposure (Supplemental Table 3). No detectable viremia was present in any of the samples from the group C pigs (Fig. 3C).
Viral nucleic acid was easily detected in oral fluids; levels were greater than nasal swabs but lower than levels detected in fecal swabs (Fig. 4). Oral fluids pooled from the pen housing groups A and B were PCR positive at 2 dpi and remained positive until 28 dpi. Oral fluids from group C appeared to be positive at the time of the first successful collection point (4 dpi) and maintained positivity until 9 dpi. Subsequent oral fluids collected from group C were PEDV negative at 14 dpi and a weak Cq (>37) was observed on 21 and 28 dpi.
PCR detection of Porcine epidemic diarrhea virus (PEDV) in pooled oral fluid samples after PEDV challenge. Data are represented by real-time PCR quantification cycles (Cq). Pen samples from groups A and B (●) and group C (○) are shown. A Cq value of ≥38 was considered negative.
Environmental PEDV
Viral nucleic acid was present on the V-troughs, pens, and food bins on both the groups A and B and group C sides of the animal room (Fig. 5). Differences between the 2 sides of the room were evident by the nucleic acid on the walls closest to the 2 pens; a Cq value of 40.0 (negative) on the group C wall and a Cq value of 31.7 on the groups A and B wall.
Porcine epidemic diarrhea virus (PEDV) detection in environmental samples collected on 14 dpi. Swabs were collected from both the groups A and B side and the group C side of the common animal room. Data are represented by real-time PCR quantification cycles (Cq). A Cq value of ≥38 was considered negative.
PEDV tissue tropism
Of the tissues tested by IHC (Supplemental Table 4, http://vdi.sagepub.com/content/by/supplemental-data), only small intestines and mesenteric lymph nodes were positive. All other tissues were IHC negative. However, small amounts of viral nucleic acid were detected by PCR in several additional tissues, including nasal turbinates, tonsil, thymus, and spleen (Supplemental Table 5, http://vdi.sagepub.com/content/by/supplemental-data).
Small intestinal sections and mesenteric lymph nodes from group A pigs had the greatest amount of nucleic acid present between 4 and 7 dpi (Supplemental Table 5). Most group A pigs (7 of 10) maintained positive Cq values in the small intestine until the end of the study at 42 dpi; whereas, all mesenteric lymph node samples had negative Cq values at this time. Interestingly, all group B pigs had negative Cq values in the small intestine at 42 dpi.
Antibody response
Serologic analysis of serum samples was performed using an IFAT and serum neutralization assays. All pre-inoculation samples were negative for antibody, and there was significant seroconversion in the group A and B animals (Fig. 6). Seroconversion at 14 dpi in these groups correlated well with resolution of viremia (compare Figs. 3 and 6). There was no evidence of seroconversion in group C, despite clear demonstration of PEDV nucleic acid in nasal and oral fluid samples. Interestingly, the group B animals reached lower peak antibody titers and appear to have more rapid antibody decay based on the IFAT (Fig. 6A). In addition, group B animals reached peak neutralization titers 1 week after group A, at 21 dpi (Fig. 6B).
Antibody response after Porcine epidemic diarrhea virus challenge in all 3 experimental groups. Data are represented by the group geometric mean indirect fluorescent antibody titer (A) and serum neutralization titer (B) during each week post-inoculation. Inoculated (▲), contact transmission (■), and aerosol transmission (○) groups are shown.
Discussion
Both contact and aerosol routes of transmission were investigated in our study. Rapid infection was clearly demonstrated in group B animals, with 4 of 5 pigs shedding PEDV in feces 2 days after exposure to the group A animals. Transmission to group B animals was likely the result of fecal-oral direct contact with group A animals; however, indirect contact with contaminated fomites, such as pens or feed bins, cannot be eliminated. Although viral shedding patterns and clinical signs were virtually identical between groups A and B, 2 important differences should be highlighted. First, intestinal samples from 7 of 10 group A pigs were PCR positive at 42 dpi, whereas none of the group B intestinal samples were positive at this time (p = 0.0256, Fisher exact test). Second, antibody titers in the group B pigs were typically lower and had more rapid decay compared to group A pigs. These differences are likely the result of the group B pigs being infected with a lower PEDV dose than group A pigs.
Productive aerosol transmission was not demonstrated in our study, despite repeated PEDV nucleic acid detection in the nares of all 5 of the aerosol transmission pigs. The dose of PEDV transmitted via aerosolization may have been too low. The Cq values detected in nasal swabs of the aerosol transmission pigs were relatively low, with the greatest quantity detected at a Cq of 31.45. This may have been because of the small amount of virus aerosolization or the BSL-3 housing conditions, where air exchange occurred almost 15 times each hour. The circulating PEDV detected in nasal swabs may have been inactivated by chemicals used to disinfect the room. The lack of productive transmission may have been the result of the number and age of the indicator animals. This level of nucleic acid may have been infectious only to highly susceptible 1-day-old piglets.
Detection of PEDV nucleic acid in fecal swabs, nasal swabs, and oral fluids was relatively consistent between 2 and 14 dpi in this age group. However, detection of PEDV viral antigen in intestinal sections was highly variable among pigs euthanized on the same day postinfection, and inconsistencies were present within different intestinal sections collected from the same pig. Of the pigs that were negative on IHC for PEDV antigen between 2 and 9 dpi, all 4 pigs were positive for PEDV nucleic acid in fecal swabs. Specifically, one of the pigs euthanized on 7 dpi was negative on IHC but was shedding large amounts of PEDV nucleic acid in feces collected on the same day (Cq = 21.52). In addition, all pigs euthanized after 2 dpi had high levels of PEDV nucleic acid detected in small intestinal samples. Discrepancies between IHC and PCR results were also reported in 3-week-old pigs after infection with a different PEDV strain. 10 These IHC inconsistencies warrant collection of several intestinal sections from multiple pigs to ensure initial diagnosis and complete resolution of PEDV carriage after an acute outbreak.
Mild to moderate clinical disease was documented for ~1 week after PEDV inoculation of the group A pigs. However, PEDV shedding in fecal and nasal swabs were detected up to 28 dpi, ~3 weeks after clinical disease had abated. In addition, PEDV nucleic acid was detected in intestinal samples from 7 of 10 group A pigs at the conclusion of the study. Long-term virus carriage and shedding in an age group with transient and mild clinical signs could be easily overlooked in a large production system. However, we do not know the infectious transmissibility of the PEDV detected in fecal swabs or intestinal samples. Moreover, each of the 7 pigs that tested positive for PEDV nucleic acid in the intestine at 42 dpi had negative fecal swabs collected on the same day, indicating the presence of intestinal PEDV with a lack of viral shedding. In a previous study, 3 pigs naturally exposed to PEDV had intermittent viral shedding through 42 dpi; however, naive pigs only became infected when introduced at 7 and 14 dpi, but not at later times. 3 Taken together, this suggests that the presence of virus in the feces and intestine several weeks after inoculation may not pose a risk for productive transmission. However, postweaning pigs should still be considered possible reservoirs for PEDV and may support subclinical infections long after resolution of clinical signs.
Footnotes
Acknowledgements
We thank Andrew Suddith, Curtis Concannon, Alexandra Fuller, and Ranjni Chand for their assistance with sample collection and laboratory testing.
Authors’ contributions
MC Niederwerder, JC Nietfeld, and J Bai contributed to design of the study; contributed to acquisition, analysis, and interpretation of data; and drafted the manuscript. L Peddireddi, J Anderson, B An, RD Oberst, DM Madson, and GA Anderson contributed to acquisition of data. B Breazeale contributed to acquisition and analysis of data, and drafted the manuscript. MA Kerrigan contributed to design of the study and contributed to acquisition of data. K Crawford and KM Lager contributed to analysis and interpretation of data. RRR Rowland contributed to analysis and interpretation of data, and drafted the manuscript. RA Hesse contributed to conception and design of the study; contributed to acquisition, analysis, and interpretation of data; and drafted the manuscript. All authors critically revised the manuscript; gave final approval; and agreed to be accountable for all aspects of the work in ensuring that questions relating to the accuracy or integrity of any part of the work are appropriately investigated and resolved.
a.
Virkon, DuPont, Wilmington, DE.
b.
Kindly provided by Dr. Matt Ackerman, Swine Veterinary Services, Greensburg, IN.
c.
MagMAX-96 viral RNA isolation kit, Life Technologies, Grand Island, NY.
d.
Kingfisher 96 magnetic particle processor, Fisher Scientific, Pittsburgh, PA.
e.
Stomacher 80 Biomaster, ThermoScientific, Swedesboro, NJ.
f.
Path-ID multiplex one-step kit, Life Technologies, Grand Island, NY.
g.
CFX96 Touch real-time PCR detection system, Bio-Rad Laboratories, Hercules, CA.
h.
Bond-Max autostainer, Leica Biosystems Inc., Buffalo Grove, IL.
i.
Polymer refine detection kit, Leica Biosystems Inc., Buffalo Grove, IL.
j.
Kindly provided by Dr. Eric Nelson, South Dakota State University, Brookings, SD.
k.
Bond primary antibody diluent, Leica Biosystems Inc., Buffalo Grove, IL.
l.
Powervision poly-HRP α-mouse polymer, Leica Biosystems Inc., Buffalo Grove, IL.
m.
Kindly provided by Dr. Sabrina Swenson, Diagnostic Virology Laboratory, National Veterinary Services Laboratory, Ames, IA.
Declaration of conflicting interests
The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.
Funding
The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work was supported by the National Pork Board award number 13-228.