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The shedding of group A rotavirus antigen in a newly established closed specific pathogen-free swine herd
Ieterinary Microbiology 28 ( 1991 ) 2 1 3 - 2 2 9 Elsevier Sciencc Publishers B.V., A m s t e r d a m
213
The shedding of group A rotavirus antigen in a newly established closed specific pathogen-free swine herd Howard B. Gelberg a, Gerald N. Woode b, Timothy S. Kniffen ~, Michele Hardy b and William F. H a l f aDepartment ~[~l'eterinao, Pathobiology, College Qf Veterinary Medicine, Universio, ~flllinois. Urbana, 1L 61801, USA bDepartment Qf l'eterinary Microbiology and Parasitology, College Qf Veterinao, Medtcine, Texas A&M University, College Station, TX 77843, USA 'Department Q/ I'eterinatT Clinical Medicine, College Qf Veterinao' Medicine, University ~llllinois, Urbana, 1L 61801, USA (Accepted 7 February 1991 )
~BSTRACT Gelberg, H.B., Woode, G.N., Kniffen, T.S., Hardy, M. and Hall, W.F., 1991. The shedding of group A rotavirus antigen in a newly established closed specific pathogen-free swine herd. Vet. MicrobioL. 28:213-229. A longitudinal study was undertaken in a newly established specific pathogen-free (SPFI swine herd to determine the dynamics of rotavirus anligen shedding in a closed swine facility. Pregnant SPF gilts which populated the herd, and their offspring, were monitored weekly for lhree consecutive lactations. Fecal samples were assayed for lhe presence of group-specific viral antigen by a solid phase immunoassay (ELISA). Results indicate that in the week prior to farrow, 35% of samples from gilts/ sows contained rotavirus antigen. During nursing, 37% of the gilts'/sows' fecal samples also contained virus antigen. Over the course of three farrowings, every gilt/sow in the herd excreted virus antigen. Virus antigen was present in 25% of the samples tested from nursing pigs and in 70% of the samples tested from pigs in the postnursing period; 95% of the litters excreted virus antigen either while nursing or postweaning. Seasonal incidence in virus antigen excretion was noted with proportionally more suckling pigs virus antigen-positive in summer and proportionally more sows/gilts positive during winter. Diarrhea occurred only rarely in the sampled population. Although piglets shed rotavirus subctinically. ELISA positive feces from piglets of each lactation caused severe disease when fed to neonatal gnotobiotic pigs. Electropherotyping of these passaged viruses indicated minor variation in RNA banding patterns over time.
INTRODUCTION
Rotaviruses are among the most ubiquitous of infectious agents. Most, if not all, swine herds are infected with rotavirus ( D e b o u k et al., 1983). Rota-
0378-1135/91/$03.50
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Elsevier Science Publishers B.V.
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H.B. GELBERG ET AL.
viruses, alone or in concert with other enteropathogenic agents, are capable of causing illness and sometimes death in swine in commercial herds. Rotaviruses are extremely resistant to chemical decontamination, and only a few compounds are capable of effectively disinfecting a site (Sattar et al., 1983). This, in combination with the virus' resistance to heat and drying, make it extremely difficult to eliminate this infectious agent from a herd. The methods by which rotaviruses enter a herd and the means by which the virus is maintained in a population are unknown. Herein, a new specific pathogenfree (SPF) facility was populated with SPF gilts for the purpose of providing a clean and dependable source of pigs for research. Since the facility had not been previously occupied, a unique opportunity existed for documenting the dynamics of rotavirus antigen shedding in a newly established herd. In this study, 54 gilts and their offspring were sampled over the course of three consecutive lactations to document the shedding of rotavirus antigen in a swine facility. Further, the pathogenicity of selected viral antigen positive fecal samples was tested in gnotobiotic pigs and the RNA electropherotype of the viruses ascertained. MATERIALS AND METHODS
Animals Fifty-four seven-month-old bred gilts were obtained from a single SPF source. Abortions were induced in 47 animals with cloprostenol (Estrumate, Haver Lockhart, Shawnee, KS) or dinoprost t r o m e t h a m i n e (Lutalyse, Upjohn, Kalamazoo, MI ) for the purpose of estrus synchronization. All gilts were bred to one of six SPF boars. Gilts were maintained in 6.4 × 1.8 m metal units, one gilt to each pen. Each gilt had nose contact with a m a x i m u m of two other pigs but minimal contact with feces. Flooring was slatted fiberglass with an underlying water pit. Approximately one week prior to anticipated date of parturition, groups of 6-12 gilts/sows were moved through a series of outside walkways to a separate farrowing facility. Here they were housed in individual farrowing crates, six animals to a room. Flooring was slatted stainless steel or fiberglass and was later changed to plastic-coated expanded metal slats over a water pit. A heating pad and infrared lamp provided additional warmth for neonatal pigs. R o o m temperature was maintained at 18-21 °C, although it did become warmer in s u m m e r months. Feed contained 16% crude protein (ground corn and soybeans) fed twice daily at a rate necessary to maintain sow condition. After farrowing, 1.8 kg feed was given daily to each sow plus an additional 0.45 kg for each nursing pig. Individual units were scraped daily, and the entire farrowing facility was cleaned with high pressure cold water and disinfected with alkyl dimethyl benzyl a m m o n i u m chloride between farrowing groups.
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Pigs were regrouped at weaning (4-5 weeks) by weight and transferred to a temperature-controlled (32°C) nursery in separate rooms at the opposite end of the same building as the farrowing unit. Feed consisted of 20% crude protein pellets with whey and molasses introduced at 10-14 days old as a creep feed. At five weeks of age, the temperature in the nursery was lowered each day until it reached 26.6°C at seven weeks of age. Pigs were housed in galvanized wire pens approximately 4.1 × 3 m, 7-10 pigs to a pen, over water pits. Pigs in one pen had nose contact with pigs in a maximum of one other pen.
Sampling procedure Fecal samples were collected weekly from each gilt/sow in the farrowing facility beginning the week prior to farrowing and continuing until the litter was weaned. Twice daily observations of the pigs were made by the herdsman. The original 54 gilts were followed for three lactations. Individual gilts/ sows were removed from the sampling regimen if their litters were stillborn or small in number in which case the neonates were cross-fostered to other dams. Cross-fostered pigs were also removed from the sampling regimen. Fecal samples were transferred to sterile polypropylene or polystyrene tubes. Samples from adult pigs were either freshly voided or obtained per rectum. For suckling pigs, individual (or rarely composite) freshly voided samples were obtained weekly. After weaning, samples were obtained by manual expression from a single different animal each week from each litter. This continued for a period of 3-4 weeks until the animals were moved from the nursery. Thus, each litter had a single sample from a randomly chosen piglet tested for viral antigen weekly during the nursing and postweaning periods. Samples were transported to the laboratory and stored at - 20 ° C until assayed.
Enzyme-linked immunosorbent assay Samples were assayed for the presence ofrotavirus antigen by a solid phase enzyme-linked immunosorbent assay (ELISA Rotazyme II, Abbott, Chicago, IL) designed to detect the group-specific (VP-6) antigen. Polystyrene beads coated with guinea-pig anti-SA- 11 (simian rotavirus) served as capture antibody. It has been determined that SA-11 antibody cross reacts with porcine rotavirus antigens (Benfield et al., 1984). Rabbit anti-SA-11 horseradish peroxidase conjugate and o-phenylenediamine.2HC1 substrate was the indicator system. Absorbance values at 492 nm (Quantum II, Abbott, Chicago, IL) were used as an indicator of virus antigen presence. Positive and negative controls included with the kit were run with each sample and cut off points were determined by a microprocessor which added 0.075 to the optical density reading of the negative control. Specimens yielding equivocal absorbance values were run in duplicate.
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H.B. GELBERG ET AL.
Specificity testing Specificity of the ELISA procedure was ascertained by a blocking assay using unlabeled negative or positive rabbit anti-SA-I 1 antibody (Dakopatts, Santa Barbara, CA) on selected fecal samples (n = 42 ). Those samples which were blocked less than 50% as well as all ELISA-negative samples were tested by negative-contrast electron microscopy (Basgall et al., 1988 ). Electron microscopy was done with coded samples, that is, without knowledge of the ELISA results. As part of an ongoing study, over 400 additional fecal samples from gestating and nursing sows, piglets, and feeder pigs from the same farm were assayed both by commercial ELISA and two monoclonal antibody based microELISA systems to test for the occurrence of false positive reactions by rotazyme. For these latter systems, capture antibody was either polyclonal goat hyperimmune serum or anti-SA- 11. Secondary antibody was monoclonal antiVP6 (Zheng et al., 1989 ). Affinity-purified peroxidase labeled goat anti-mouse antibody (Kirkegaard and Perry Labs, Gaithersburg, MD) and o-phenylenediamine.2HCi was the indicator system. Known positive and negative fecal samples were assayed with each plate. Optical density was determined by spectrophotometry at 490 nm using a micro-ELISA autoreader (MR50, Dynatech Labs, Alexandria, VA). Net optical density (OD) readings greater than 0.2 were considered positive, while those below OD 0.1 were considered negative. Absorbance values between 0.10 and 0.20 were rerun a n d / o r rechecked by another method.
Transmission studies For transmission studies, pairs of caesarean derived pigs were maintained in gnotobiotic isolators and fed a commercial milk replacer (SPF LAC, Borden Inc. ). Each pair of pigs, at 24 h of age, was orally inoculated with 0.5-2 ml of clarified, filtered (0.45/~) feces from a single EEISA positive piglet. There were two fecal samples each from the first, second and third parities. A total of 12 pigs were therefore utilized for the transmission studies. One of each pair of pigs was euthanized at 24 h postinoculation (p.i.) and the other at 48 h p.i. Fecal samples from the gnotobiotic pigs were assayed for group A specific antigen by micro-EEISA. Intestinal samples were fixed in formalin t'or immunohistochemical detection of group A rotavirus antigen in tissue sections.
Immunohistochemistry Tissue sections from seven equidistant regions of small intestine (McAdaragh et al., 1980) were fixed in neutral buffered formalin, processed in low melt paraffin (Peel-A-Way, American Scientific Products, McGaw Park, IL), and sectioned to 5 ,urn thick. They were then processed for immunohis-
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tochemical localization of group A specific rotavirus antigen (Gelberg et al., 1990).
RNA electropherotyping RNA extracted from feces of the virus infected gnotobiotic piglets was electropherotyped as described previously (Theft et al., 1981; Gaul et al., 1982; Zheng et al., 1989 ).
Statistical analyses Qualitative data (presence or absence of virus antigen) was compared by Chi squared analysis with a P~< 0.05 denoting significance. Statistical analyses were run on SAS (Statistical Analysis System, Raleigh, NC ) using a Proc. GLM (general linear models procedure). RESULTS Forty-two samples were checked by ELISA, blocking ELISA a n d / o r EM. Twenty-three were confirmed positive for rotavirus antigen. Eighteen samples were negative for rotavirus antigen by ELISA, blocking ELISA and EM. One sample was positive by ELISA and negative by blocking ELISA and EM. We did not have sufficient feces to retest this sample. Overall agreement between Rotazyme and one or both of the validation assays was therefore 97.6% on the 42 samples tested by the three assays. Of the Rotazyme-positive samples, 23 of 24 were confirmed positive by another test (95.8%). Of the 400 samples tested both by commercial ELISA and monoclonal micro-ELISA, only four were reactive by commercial ELISA. These same four samples were positive by monoclonal micro-ELISA. All four of these samples contained small numbers of rotavirus-like particles on EM examination. Diarrhea was rarely observed in the herd under study, was transient, and occurred in one or several nursing animals in a litter. The presence of diarrhea was not correlated with detection of group A specific antigen in feces. During the week prior to parturition, 17 of 44 (38.6%) samples from gilts, 5 of 34 (14.7%) samples from second litter sows, and 19 of 38 (50.0%) samples from third litter sows contained rotavirus antigen (Fig. 1 ). Second lactation sows excreted virus antigen at a significantly ( P = 0 . 0 2 2 ) lower rate than gilts or third lactation sows. Taken as a single farrowing unit, 41 of 116 (35.3%) fecal samples from gilts/sows were positive for rotavirus antigen in the week prior to farrowing (Table 1 ). During the nursing period, gilts shed virus antigen in 49 of 153 (32.0%) samples (Fig. 1 ). During the second lactation, sows shed virus antigen in 45 of 136 (33.1%) samples while nursing and third lactation sows shed virus antigen in 54 of 112 (48.2%) samples while nursing. Second lactation sows excreted virus antigen significantly ( P = 0.004) less often than the other two
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H.B. GELBERG ET AL.
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Gilts/sows A B C Pigs (nursing) A B C Pigs (weaned) A B C
Weeks postfarrow
Weeks postweaning 1
-1
1
2
3
4
41 116 35.3
32 109 29.4
40 111 36.0
42 101 41.6
34 80 42.5
189 517 36.6
21 103 20.4
22 116 19.0
25 108 23.1
34 80 42.5
t02 407 25.1 29 41 70.7
2
Totals
58 86 67.4
3
67 92 72.8
154 219 70.3
ICombined data from three consecutive parities. A: N u m b e r o f samples ELISA positive for group A specific rotavirus antigen. B: N u m b e r of samples assayed. C: Percent antigen positive samples.
groups during week one of lactation and gilts significantly ( P = 0 . 0 0 1 ) less often during week three. Taking the three parities as a single unit, lactating gilts/sows shed virus antigen in 32 of 109 samples (29.4%) during week one, in 40 of 111 (36.0%) samples during week two, in 42 of 101 (41.6%) samples during week three, and in 34 of 80 (42.5%) samples during week four (Table 1 ). Overall, 148 of 401 (36.9%) samples were virus antigen positive.
SHEDDING
OF ROTAVIRUS
ANTIGEN
2 19
Taken as a whole, 38 of 54 (70.4%) gilts excreted virus antigen at some time during the sampling period. For second litter sows, 25 of 49 (51.0%) excreted virus antigen, and 32 of 38 (84.2%) third litter sows excreted virus antigen. Overall, for all farrowing pigs, 95 of 141 (67.4%) demonstrated virus antigen on at least one occasion for each parity. All 37 of the gilts/sows followed for three lactations excreted virus antigen sometime during the study. Pigs nursing gilts excreted virus antigen in 40 of 150 (26.7%) samples (Fig. 2). After weaning, 39 of 69 (56.5%) pig fecal samples from gilts contained virus antigen. Among first lactation pigs, 26 of 53 (49.1%) litters excreted virus antigen sometime during the nursing period, 32 of 41 (78.0%) litters excreted virus antigen during the postweaning period, and 37 of 41 (90.2%) litters excreted virus antigen at least once during the study. Nursing pigs from second litter sows excreted virus antigen in 45 of 138 (32.6%) samples (Fig. 2). Weaned pigs from second lactation sows excreted virus antigen in 53 of 79 ( 59.6% ) samples. Among second litter pigs, 30 of 44 (68.2%) litters excreted virus antigen sometime during nursing 36 of 42 ( 85.7% ) sometime during the postweaning period, and 41 of 42 (97.6%) litters excreted virus antigen sometime during the study. Nursing pigs of third litter sows excreted virus antigen in 17 of 119 (14.3%) samples (Fig. 2). Weaned pigs from third litter sows contained rotavirus antigen in their feces in 62 of 71 (87.3%) samples. Among third lactation pigs, 18 of 38 (47.4%) litters excreted virus antigen sometime during nursing, and 31 of 35 (88.6%) during the postweaning period; 34 of 35 (97.1%) litters excreted virus antigen sometime during the study.
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Fig. 2. Virus antigen shedding by pigs during nursing and post-weaning. N = Number of samples assayed.
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H.B. GELBERGETAL
During the first postweaning week, pigs of gilts excreted virus antigen significantly (P=0.029) less often than the other two groups. Over three parities, pigs while nursing excreted virus antigen in 21 of 103 (20.4%) samples during week one, 22 of 116 (19.0%) samples during week two, 25 of 108 (23.1%) samples during week three, and 34 of 80 (42.5%) samples during week four (Table 1 ). As a single unit, over three parities, suckling pigs excreted virus antigen in 102 of 407 (25.1%) samples. Seventy-four of 135 (54.8%) litters excreted virus antigen sometime during nursing. Taken as a single farrowing unit regardless of the dam's age, weaned pigs excreted virus antigen in 29 of 41 (70.7%) samples during week one, in 58 of 86 (67.4%) samples during week two, and in 67 of 92 (72.8%) samples during week three (Table 1 ). Overall, feces from weaned pigs contained virus antigen in 154 of 219 (70.3%) samples; 99 of 118 (83.4%) litters excreted virus antigen during the postweaning period. Combining data from the three parities into a single farrowing unit, 112 of 118 (94.9%) litters demonstrated virus antigen in their feces sometime during the study. Among all piglet samples, 256 of 626 (40.9%) were virus antigen positive. Weaned pigs excreted virus antigen more often than suckling pigs and there was a steady increase in the percent of pigs excreting antigen with increasing age. A seasonal peak in numbers of gilts/sows excreting virus antigen was observed during the winter months (December through March, P < 0.005 ) and a seasonal trough in late summer and early fall (July through September) (Fig. 3). Nursing pigs excreted virus antigen most often in winter and sum-u- Nursing Pigs + Post-weaning pigs -~- Gilts/Sows 100
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Fig. _3. Seasonal rotavirus shedding by gilts/sows and piglets.
SHEDDING OF ROTAVIRUS ANTIGEN
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met and least often in spring (P<0.005) while weaned pigs excreted virus antigen more evenly through the year with a significant (P<0.005) peak in early spring and with troughs in April, May, and October (Fig. 3 ). As the rotavirus positive fecal samples had been obtained from subclinically infected animals, the following experiment was conducted to determine the virulence, if any, of the rotavirus present in the population. Two separate fecal samples for each of the three parities was passaged in gnotobiotic piglets. By 48 h p.i., all inoculated animals had watery diarrhea and positive fecal micro-ELISA values. Immunohistochemical evaluation of small intestine showed marked villous epithelial loss and blunting with abundant brown immunohistochemical reaction product in the cytoplasm of surviving enterocytes (Fig. 4 ). These results were similar to previous reports of the pathogenesis of rotavirus infection in pigs (Theil et al., 1978; McAdargh et al., 1980). Negative tissues and negative controls lacked cytoplasmic reaction product. Preincubation of virus with antibody completely blocked specific staining of rotavirus antigen in tissue sections. Viral RNA was extracted from feces of experimentally infected pigs and the electrophoretic banding patterns (Fig. 5) were shown to be similar among samples from the three parities. One sample, from a pig inoculated with feces from parity one that died prior to sacrifice and was severely autolyzed, was micro-ELISA positive and electrophoretically negative. One sample from
Fig. 4. Mid-jejunum from gnotobiotic pig 48 h p.i. with filtered feces from a parity one piglet. Villi are short and blunt. Dark cells at tips of villi are immunoperoxidase positive for group A rotavirus antigen. Hematoxylin counterstain.
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H . B . G E L B E R G E T AL.
A
B
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E
F
G
1 ~
2-...3/4 ~--
m
6m
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10-11--
Fig. 5. Electrophoretic migration patterns of viral RNA extracted from feces of newborn, gnolobiotic pigs experimentally inoculated with feces from parity one (lane B), parity two (lanes C and D), and parity three (lanes E and F) pigs. Reference OSU (lanes A and G). Lane B contains a mixture of group A minor variants. Lane D contains, in addition to group A, group C rotavirus RNA. p a r i t y two d e m o n s t r a t e d , in a d d i t i o n to g r o u p A r o t a v i r u s , the presence o f an atypical r o t a v i r u s g e n o t y p e . DISCUSSION In spite o f a large b o d y o f literature d o c u m e n t i n g different aspects o f rotavirus infection a n d disease in m a n y species o f d o m e s t i c animals, little has
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been published about the natural history of rotavirus antigen @edding over time under field conditions. We had opportunity to document; in a newly established closed SPF swine herd, the dynamics of rotavirus antigen shedding both in individual animals as they aged, and in the herd in general. Rotavirus disease is generally observed in the 1-4-week-old age group (k~,ohl et al., 1978) but severe disease has also been reported in animals weaned~at 6-8 weeks of age (Woode, 1986). We were curious to determine whether a newly established closed herd, in newly constructed facilities, would develop enzootic rotavirus infections when populated entirely by adult animals presumably resistant to rotavirus disease. Assuming that virus antigen was introduced into the herd under these conditions, we were also interested in learning how virus antigen spreads through the herd over successive parities, whether there is an electrophoretic change in virus over time, whether the gilts/sows excreted virus antigen more or less often with increasing age and whether offspring of older sows were more or less at risk of infection. The assay used for the detection of virus antigen in this study was a commercial ELISA designed to detect the group specific (VP-6) antigen c o m m o n to all group A rotaviruses. Studies in various species have compared the results of the commercial ELISA used in this study (Rotazyme II) with other means of rotavirus detection (Cornell et al., 1982; Rubenstein and Miller, 1982; Benfield et al., 1984; Morinet et al., 1984; Chernesky et al., 1985; Miotti et al., 1985). Two studies specifically tested pig feces (Cornell et al., 1982; Benfield et al., 1984). In one study the authors found when comparing negative contrast EM with commercial ELISA, that of 157 samples tested, sensitivity of the ELISA was 96% (Cornell et al., 1982). ELISA positive EM negative samples were shown to contain rotavirus antigen by a blocking assay. The results from our limited (42 samples) specificity testing indicated that 23 of 24 positive samples by ELISA were confirmed positive by either EM or blocking ELISA (95.8% ). In the other study involving rotavirus antigen detection in samples from swine, agreement between EM and ELISA was 72% with EM detecting more positive samples than did commercial ELISA (Benfield et al., 1984). In that study, positive cut off values for commercial ELISA were an optical density of >~0.1 at 490 nm. In our study cut off values varied depending on the positive and negative samples included with the kit. They ranged from an OD of 0.073 to 0.144. A total of 22 of 446 ( < 5%) of the positive samples had OD values < 0.1. While EM might have increased the number of rotavirus positive samples from our swine population, it would not determine if ELISAnegative, EM positive samples contained group A rotavirus or morphologically identical, but antigenically distinct, group B or C rotaviruses. Studies have shown that atypical rotaviruses are not detected by Rotazyme (Cornell et al., 1982 ). In a Belgian study, pararotavirus infections were seen in a large proportion of pigs from three different herds (Debouk et al., 1983 ).
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Besides suggesting the possible presence of atypical rotavirus in ELISA negative EM positive samples, the authors of the paper reporting 72% agreement between commercial ELISA and EM postulate that the presence of inhibitors in fecal material that reduces binding of rotavirus to antibody, intestinal luminal antirotavirus antibodies and immune complexes may create false negatives in the enzyme immunoassay as compared with EM (Benfield et al., 1984). False positive reactions were not reported in either of the swine studies, or in our continuing studies assaying 400 sow and piglet samples both by commercial ELISA and confirmatory monoclonal-based micro-ELISAs. These data contrast with a recent study comparing commercial rotavirus assays on human fecal specimens. That study found that while the commercial ELISA used in our studies correctly identified all positive samples, it also produced false positive reactions (Dennehy et al., 1988 ). The reasons for the apparent difference in specificity of commercial ELISA on human versus porcine specimens are not clear but rheumatoid factor-like substances in human stool may be responsible for lowered specificity in that species (Yolken and Stopa, 1979). In a study which monitored rotavirus excretion in experimentally infected gnotobiotic pigs, EM detected virus excretion for a longer period than commercial ELISA (Benfield et al., 1988). In addition, commercial ELISA is directed against SA-11 which is subgroup 1 specific. Thus, while all group A rotaviruses should be detected by this method, subgroup 1 viruses, such as porcine OSU may react more strongly than subgroup 2 porcine rotaviruses accounting for a discrepancy between EM and Rotazyme. This longitudinal study has revealed much about the natural history of rotavirus infections in a well managed SPF swine herd. It has reaffirmed the notion that rotaviruses are ubiquitous, suggests that subclinical infections are commonplace in pigs and that age resistance to rotavirus infection may not occur. Importantly, gilts/sows often shed virus antigen prior to farrowing and during nursing and that it is therefore unrealistic to expect to be able to eliminate rotavirus from an infected swine breeding unit. We believe, based on the fact that all gilts/sows excreted virus antigen sometime during the study, that our results are not spurious and that rotavirus shedding may be a common event even in well-managed swine operations. We agree with others who suggest that sows may play an important role as the source of rotavirus infection of pigs (Benfield et al., 1982). It is postulated that the dam sheds virus at a time when the neonate has large amounts of passive immunity and can best cope with infection (Bohl, 1980). This would explain the subclinical nature of the infection in suckled piglets. Later infections occur in presumably immune pigs, resulting in a lack of clinically detectable disease. In another study (Benfield et al., 1982), 12 sows were sampled every four days beginning five days prior to farrow to 14 days postpartum. Virus shed-
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ding was determined by EM a n d / o r virus isolation after concentration of the fecal specimens. Five animals (42%) shed virus. Four of 12 litters developed diarrhea but only two litters were from sows who shed virus. In a different study, 33% of postpartum sows in one herd and 10% of postpartum sows from a second herd excreted virus antigen as detected by ELISA (Grom et al., 1984). It appears that the gilts/sows either harbor the organism in a latent form which is activated by hormonal or other changes occurring near parturition; are continuously infected and shed virus antigen throughout their lives (Lecce and King, 1980), or are reinfected. Since serum antirotavirus IgG antibody levels in gilts/sows increase during lactation and decrease between farrowings in the animals under study (Gelberg et al., 1991 ), we suspect that active infections occur in those gilts/sows that excrete antigen. Our infectivity studies indicate that virulent virus was present in the herd, and the results contrast with another study which concluded that reinfection of seropositive sows seemed unlikely (Benfield et al., 1988 ). Although we cannot exclude the possibility that some of the animals shed virus passively following ingestion, but without active infection, we think that this is unlikely. The gnotobiotic piglets experimentally infected with ELISA positive feces became ill and demonstrated histologic and immunohistochemical evidence of virus infection. From these data, we can conclude that the rotaviruses present were virulent strain (s). Viral RNA extracted from the experimental pigs' feces showed minor variations in banding patterns which suggests that they arose from similar viral parents, and suggests that in this closed herd, new rotavirus group A types were not introduced, a n d / o r several strains were cocirculating. Our study suggests the virus to be most prevalent in older (third parity) dams and least prevalent in their nursing pigs. In other studies (Svensmark, 1983; Fu and Hampson, 1987 ) rotavirus infection was most common in pigs born to gilts. Our evidence contradicts other studies suggesting that weaned pigs rarely excrete virus (Debouck et al., 1983 ) and sows never excrete virus (Debouk et al., 1983; Fu and Hampson, 1989). Pigs were weaned at a later age in the former study perhaps accounting for the difference in data. Generally, more litters excreted virus antigen during the postweaning period than during nursing but there was a nearly linear increase in virus antigen excretion with age (Fig. 2). While it is tempting to attribute this increase in antigen excretion to loss of passive immunity, in agammaglobulinemic human patients, human milk caused disappearance of rotavirus from feces with reappearance while the patient was still receiving milk (Saulsbury et al., 1980 ). Likewise, in agammaglobulinemic pigs, age dependent resistance to rotavirus is noted to occur (Kirstein et al., 1985). In children, in addition to the state of immunity, the outcome of rotavirus infection may depend on chronologic age and state of maturity of the gastrointestinal tract which contributes to a
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state of transient virus-host equilibrium, which results in carriage, asymptomatic infection or disease (Champsaur et al., 1984a). In pigs, environmental, social, and physiologic stresses (Kirstein et al., 1985) as well as changes in brush border enzyme activity (Hampson and Kidder, 1986) occurring at weaning may also be contributory to the development of disease a n d / o r an increase in the rate of enterocyte infection and viral antigen shedding. At weaning, villous height decreases and the comparatively short crypts of the pig's distal small intestine elongate providing a large, less mature enterocyte population theoretically more susceptible to malabsorptive diarrheas (Hampson, 1986). Our results question the value of viral diagnosis from feces since the presence of virus antigen in the feces was not correlated with the presence of disease. Our results are similar to a study of Danish swine herds in which 30% of fecal samples from normal animals contained rotaviral antigen (Svensmark, 1983). Likewise, in a study of all children under 2 years of age admitted to a pediatric ward there was a high incidence of asymptomatic rotavirus shedding and no correlation between most virus shedding and diarrhea (Champsaur et al., 1984b). Collectively, gilts/sows shed virus antigen in 189 of 517 (36.6%) fecal samples regardless of the stage of lactation including the week prior to parturition. Thus, virus antigen apparently is distributed throughout the nursery at all times, although the incidence of shedding by gilts/sows is somewhat higher in winter. The reasons for this are not clear. Reports ofrotavirus outbreaks in humans suggest an increased incidence of rotavirus gastroenteritis in cold seasons versus warm seasons (Kapikian et al., 1976; Konno et al., 1978). Seasonal increases in numbers of suckling pigs excreting virus antigen were noted with peak numbers in summer and to a lesser extent in winter as opposed to the gilts/sows whose excretion data peaked in winter. More weaned pigs had virus antigen in their feces in April, May, and October, although peaks and troughs were relatively flat. In a Venezuelan study, seasonal variation in rotavirus excretion by pigs was not seen; rotavirus was not detected in the feces of pigs less than one week of age (we detected virus antigen at three days of age, we did not sample many animals earlier than this) and peaked at 3-4 weeks of age (pigs were followed to 4-6 weeks of age) (Utrera et al., 1984). Virus was detected in 21.3 percent of all diarrheic fecal samples. No virus was detected from control animals. The Venezuelan study was in a relatively stable climatic environment perhaps mimicked best by our postweaning experimental group. In this latter group, the number of samples containing virus antigen was relatively constant. Others have suggested that temperature is not the important variable in virus excretion but relative humidity; dry environments therefore allowing for dust-borne spread of virus (Brandt et al., 1982; Paul and Erinle, 1982). In conclusion, the presence of virus antigen excretion by the gilts/sows and
SHEDDING OF ROTAV1RUS ANTIGEN
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the presence of virus antigen in the feces of young pigs suggests that elimination of rotavirus antigen from a swine facility is not an easily attainable goal. In addition, there are at least 4 serotypes of porcine rotavirus (Paul et al., 1988 ), and cross-protection between serotypes is unlikely (Bohl et al., 1984 ). A separate study (Gelberg et al., 1991 ) documents active antibody response and passive transfer of immunity in these experiments. It is possible, based on this study, field investigations, and the work of others (Lecce et al., 1978; McNulty and Logan, 1982), that rotavirus diarrhea develops in those herds with detectable sow immunity because of build-up of rotavirus in the environment permitting the infection to overcome passive immunity. The development of disease versus infection may thus be a function of virus challenge dose. ACKNOWLEDGEMENTS
The authors thank Dr. C. Almgren, Dr. E. Basgall, Dr. E. Cohen, Ms. A. Gast, Mr. E. Parr, and Dr. J. Patterson for technical assistance. This work was supported in part by a grant from the Illinois Department of Agriculture.
REFERENCES Basgall, E.J., Scherba, G. and Gelberg, H.B., 1988. Diagnostic virology in veterinary pathology: techniques for negative staining. Proc. Electron Microscopy Society of America, pp. 366367. Benfield, D.A., Stotz, I., Moore, R. and McAdaragh, J.P., 1982. Shedding of rotavirus in feces of sows before and after farrowing. J. Clin. Microbiol., 16: 186-190. Benfield, D.A., Stotz, I.J., Nelson, E.A. and Groon, K.S., 1984. Comparison of a commercial enzyme-linked immunosorbent assay with electron microscopy, fluorescent antibody, and virus isolation for the detection of bovine and porcine rotavirus. Am. J. Vet. Res., 45:19982002. Benfield, D.A., Francis, D.H., McAdaragh, J.P., Johnson, D.D., Bergeland, M.E., Rossow, K. and Moore, R., 1988. Combined rotavirus and K99 Escherichia coli infection in gnotobiotic pigs. Am. J. Vet. Res., 49: 330-337. Bohl, E.H., 1980. Enteric viral infections as related to diarrhea in swine. In: S.D. Acres (Editor). Proc. Third Int. Symp. Neonatal Diarrhea. October 6-8, Saskatoon, Saskatchewan, Canada, Vet. Inf. Dis. Org., pp. 1-19. Bohl, E.H., Kohler, E.M., Saif, L.J., Cross, R.F., Agnes, A.G. and Theft, K.W., 1978. Rotavirus as a cause of diarrhea in pigs. J. Am. Vet. Med. Assoc., 172: 458-463. Bohl, E.H., Theil, K.W. and Saif, L.J., 1984. Isolation and serotyping of porcine rotaviruses and antigenic comparisons with other rotaviruses. J. Clin. Microbiol., 19:105-111. Brandt, C.D., Kim, H.W., Rodriguez, W.J., Arrobio, J.O., Jeffries, B.C. and Parrott, R.H., 1982. Rotavirus gastroenteritis and weather. J. Clin. Microbiol., 16: 478-482. Champsaur, H., Henry-Amar, H., Goldszmidt, D., Prevot, J., Bourjouane, M., Questiaux, E. and Bach, C., 1984a. Rotavirus carriage, asymptomatic infection, and disease in the first two years of life. It. Serological response. J. Infect. Dis., 149: 675-682. Champsaur, H., Questiaux, E., Prevot, J., Henry-Amar, M., Goldszmidt, D. and Bourjouane,
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M., 1984b. Rotavirus carriage, asymptomatic infection, and disease in the first two years of life. I. Virus shedding, J. Infect. Dis., 149: 667-674. Chcrnesky, M., Castriciano, S., Mahony, J. and DeLong, D., 1985. Examination of the Rotazyme II Enzyme Immunoassay for the diagnosis of rotavirus gastroentcritis. J. Clin. Microbiol., 22: 462-464. Cornell, W.D., Whitaker, H.K., Craft, J. and Alderson. C., 1982. A comparison of negative contrast electron microscopy and the enzyme-linked immunosorbent assay in the detection of rotavirus infection in pigs and calves. Proc. Am. Assoc. Vet. Lab. Diag. 25th Ann. Meet., November 7-9, pp. 209-220. Debouck, P., Callebaut, P. and Pensaert, M., 1983. The pattern of rotavirus and pararotavirus excretions in pigs on closed swine herds. In: S.D. Acres (Editor). Proc. Fourth Int. Syrup. Neonatal Diarrhea. Saskatoon, Canada, October 3-5, Vet. Inf. Dis. Org.. pp. 77-87. Dennchy, P.H.. Gauntlett, D.R. and Tente, W.E., 1988. Comparison of nine commercial immunoassays for the detection of rotavirus in fecal specimens. J. Clin. Microbiol., 26: 16301634. Fu, Z.F. and Hampson, D.J., 1987. Group A rotavirus excretion patterns in naturally infected pigs. Rcs. Vet. Sci., 43: 297-300. Fu, Z.F. and Hampson, D.J., 1989. Natural transmission of group A rotavirus within a pig population. Res. Vet. Sci., 46:312-317. Gaul, S.K., Simpson, T.F., Woode, G.N. and Fulton, R.W., 1982. Antigenic relationship among some animal rotaviruses: Virus neutralization in vitro and cross-protection in piglets. J. Clin. Microbiol., 16: 495-503. Gelberg. H.B., Hall, W.F., Woode, G.N., Basgall, E.J. and Scherba. G,, 1990. Multinucleate enterocytcs associated with experimental group A porcine rotavirus infection. Vet. Pathol.. 27: 453-454. Gelberg, H.B., Patterson, J.S. and Woode, G.N. A longitudinal study ofrotavirus antibody tilers in swine in a closed specific pathogen-free herd. Vet. Microbiol., 28:231-242. Groin, J., Bernard, S., Valencak, Z. and Zeleznik, Z., t984. Characteristic of rotavirus inli~clions m industrial production of pigs. Proc. Int. Pig Vet. Soc. Congress, Ghent. Belgium, p. 58. Hampson, D.J., 1986. Alterations in piglet small intestinal structure al weaning. Res. Vet. Sci., 40: 32-40. Hampson, D.J. and Kidder, D.E., 1986. Influence of creep feeding and weaning on brush border enzyme activities in the piglet small intestine. Res. Vet. Sci., 40:24-31. Kapikian, A.Z.. Kim. H.W., Wyalt, R.G., Cline, W.L., Arrobio, J.D., Brand, C.D., Rodriquez, W.J., Sack. D.A., Chanock, R.M. and Parrott, R.H., 1976. Human reovirus-like agent as a major pathogen associated with "winter" gastroenteritis in hospitalized infants and young children. New Eng. J. Med., 294: 965-972. Kirstein, C.G.. Clare, D.A. and Lecce, J.G., 1985. Development of resistance of cnterocytes to rotavirus in neonatal, agammaglobulinemic piglets. J. Virol., 55: 567-573. Konno. T., Suzuki, H., lmai, A., Kutsuzawa, T., lshida, N., Katsushima, N., Sakamoto, M., Kitaoka, S., Tsuboi, R. and Adachi, M., 1978. A long-term survey of rotavirus infection in Japanese children with acute gastroenteritis. J. lnfec. Dis., 138: 569-576. Lccce. J. and King, M.W., 1980. Persistent rotaviral infection producing multiple episodes of diarrhea in weanling pigs reared in isolation. In: S.D. Acres (Editor), Proc. Third Int. Syrup. Neonatal Diarrhea. October 6-8, Saskatoon, Saskatchewan, Canada, Vet. Inf. Dis. Org., pp. 21-34. Leccc, J.W., King, M.W. and Dorsey, W.E., 1978. Rearing regimen producing piglet diarrhea ( rotavirus ) and its relevance to acute infantile diarrhea. Science, 199: 776-778. McAdaragh, J.P., Bergeland, M.E., Meyer, R.C., Johnshoy, M.W., Stotz, l.J., Benfield, D.A. and Hammer, R., 1980. Pathogenesis of rotaviral enteritis in gnotobiotic pigs: A microscopic study. Am. J. Vet. Res., 39: 1223-1228. McNult,~, M.S. and Logan, E.F., 1992. Longitudinal survey ofrotavirus infection in calves. Vet. Rec.. 113: 333-335.
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Miotti, P.G., Eiden, J. and Yolken, R.H., 1985. Comparative efficiency of commercial immunoassays for the diagnosis of rotavirus gastroenteritis during the course of infection. J. Clin. Microbiol., 22: 693-698. Morinet, F., Ferchal, F., Colimon, R. and Perol, Y., 1984. Comparison of six methods for detecting human rotavirus in stools. Eur. J, Clin. Microbiol., 3:136-140. Paul, M.O. and Erinle, E.A.~ 1982. Influence of humidity on rotavirus prevalence among Nigerian infants and young children with gastroenteritis. J. Clin. Microbiol., 15: 212-215. Paul, P.S., l~yoo, Y.S., Andrews, J.J. and Hill, H.T., 1988. Isolation of two new serotypes of porcine rotavirus from pigs with diarrhea. Arch. V irol., 100: 139-143. Rubenstein, A.S. and Miller, M.F., 1982. Comparison of an enzyme immunoassay with electron microscopic procedures for detecting rotavirus. J. Clin. Microbiol., 15: 938-944. Saltar, S.A., Raphael. R.A. and Springthorpe, V.S., 1983. Rotaviruses and chemical disinfectants. In: S.D. Acres (Editor). Proc. Fourth Int. Symp. Neonatal Diarrhea. October 3-5, Saskatoon, Canada, Vet. Inf. Dis. Org., pp. 90-98. Saulsbury, F.T., Winkelstein, J.A. and Yolken, R.H., 1980. Chronic rotavirus infection in immunodeficiency. J. Pediatr., 97: 61-65. Svensmark~ B., 1983. Prevalence rate of porcine rotavirus in Danish swine herds. Ann. Rech. Vet.. 14: 433-436. Theil, K.W., Bohl, E.H., Cross, R.F., Kohler, E.M. and Agnes, A.G., 1978. Pathogenesis of porcine rotaviral infection in experimentally inoculated gnotobiolic pigs. Am. J. Vet. Res., 39: 213-220. Theil, K.W., McCloskey, C.M,, Saif, L,.J., Redman, D.R., Bohl, E.H., Hancock, D.D., Kohler, E.M. and Moorhead, P.D., 1981. Rapid, simple method of preparing double-stranded ribonucleic acid for analysis by polyacry|amide gel electrophoresis. J. Clin. Microbiol., 14: 273280. Utrera, V., Mazzali de llja, R., Gorziglia, M. and Esparza. J., 1984. Epidemiological aspects of porcine rotavirus infection in Venezuela. Res. Vet. Sci., 86:310-315. Woode, G.N., 1986. Porcine rotavirus infection. In: A.D. Eeman, B. Straw, R.D. Glock, W.L. Mengeling, R.H.C. Penny, and E, Scholl (Editors), Diseases of swine, 6th edn. Iowa State University Press, Ames~ Iowa, pp. 368-382. Yolken, R.H. and Stopa, P.J., 1979. Analysis of nonspecific reactions in enzyme-linked immunosorbent assay testing for human rotaviruses. J. Clin. Microbiol., 10: 703-707. Zhcng, S., Woode, G.N, Melendy, D.R. and Ramig, R.F., 1989. Comparative studies of the antigenic polypeptide species VP4, VP6 and VP7 of three strains of bovine rotavirus. J. Clin. Microbiol., 27: 1939-1945.
213
The shedding of group A rotavirus antigen in a newly established closed specific pathogen-free swine herd Howard B. Gelberg a, Gerald N. Woode b, Timothy S. Kniffen ~, Michele Hardy b and William F. H a l f aDepartment ~[~l'eterinao, Pathobiology, College Qf Veterinary Medicine, Universio, ~flllinois. Urbana, 1L 61801, USA bDepartment Qf l'eterinary Microbiology and Parasitology, College Qf Veterinao, Medtcine, Texas A&M University, College Station, TX 77843, USA 'Department Q/ I'eterinatT Clinical Medicine, College Qf Veterinao' Medicine, University ~llllinois, Urbana, 1L 61801, USA (Accepted 7 February 1991 )
~BSTRACT Gelberg, H.B., Woode, G.N., Kniffen, T.S., Hardy, M. and Hall, W.F., 1991. The shedding of group A rotavirus antigen in a newly established closed specific pathogen-free swine herd. Vet. MicrobioL. 28:213-229. A longitudinal study was undertaken in a newly established specific pathogen-free (SPFI swine herd to determine the dynamics of rotavirus anligen shedding in a closed swine facility. Pregnant SPF gilts which populated the herd, and their offspring, were monitored weekly for lhree consecutive lactations. Fecal samples were assayed for lhe presence of group-specific viral antigen by a solid phase immunoassay (ELISA). Results indicate that in the week prior to farrow, 35% of samples from gilts/ sows contained rotavirus antigen. During nursing, 37% of the gilts'/sows' fecal samples also contained virus antigen. Over the course of three farrowings, every gilt/sow in the herd excreted virus antigen. Virus antigen was present in 25% of the samples tested from nursing pigs and in 70% of the samples tested from pigs in the postnursing period; 95% of the litters excreted virus antigen either while nursing or postweaning. Seasonal incidence in virus antigen excretion was noted with proportionally more suckling pigs virus antigen-positive in summer and proportionally more sows/gilts positive during winter. Diarrhea occurred only rarely in the sampled population. Although piglets shed rotavirus subctinically. ELISA positive feces from piglets of each lactation caused severe disease when fed to neonatal gnotobiotic pigs. Electropherotyping of these passaged viruses indicated minor variation in RNA banding patterns over time.
INTRODUCTION
Rotaviruses are among the most ubiquitous of infectious agents. Most, if not all, swine herds are infected with rotavirus ( D e b o u k et al., 1983). Rota-
0378-1135/91/$03.50
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Elsevier Science Publishers B.V.
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viruses, alone or in concert with other enteropathogenic agents, are capable of causing illness and sometimes death in swine in commercial herds. Rotaviruses are extremely resistant to chemical decontamination, and only a few compounds are capable of effectively disinfecting a site (Sattar et al., 1983). This, in combination with the virus' resistance to heat and drying, make it extremely difficult to eliminate this infectious agent from a herd. The methods by which rotaviruses enter a herd and the means by which the virus is maintained in a population are unknown. Herein, a new specific pathogenfree (SPF) facility was populated with SPF gilts for the purpose of providing a clean and dependable source of pigs for research. Since the facility had not been previously occupied, a unique opportunity existed for documenting the dynamics of rotavirus antigen shedding in a newly established herd. In this study, 54 gilts and their offspring were sampled over the course of three consecutive lactations to document the shedding of rotavirus antigen in a swine facility. Further, the pathogenicity of selected viral antigen positive fecal samples was tested in gnotobiotic pigs and the RNA electropherotype of the viruses ascertained. MATERIALS AND METHODS
Animals Fifty-four seven-month-old bred gilts were obtained from a single SPF source. Abortions were induced in 47 animals with cloprostenol (Estrumate, Haver Lockhart, Shawnee, KS) or dinoprost t r o m e t h a m i n e (Lutalyse, Upjohn, Kalamazoo, MI ) for the purpose of estrus synchronization. All gilts were bred to one of six SPF boars. Gilts were maintained in 6.4 × 1.8 m metal units, one gilt to each pen. Each gilt had nose contact with a m a x i m u m of two other pigs but minimal contact with feces. Flooring was slatted fiberglass with an underlying water pit. Approximately one week prior to anticipated date of parturition, groups of 6-12 gilts/sows were moved through a series of outside walkways to a separate farrowing facility. Here they were housed in individual farrowing crates, six animals to a room. Flooring was slatted stainless steel or fiberglass and was later changed to plastic-coated expanded metal slats over a water pit. A heating pad and infrared lamp provided additional warmth for neonatal pigs. R o o m temperature was maintained at 18-21 °C, although it did become warmer in s u m m e r months. Feed contained 16% crude protein (ground corn and soybeans) fed twice daily at a rate necessary to maintain sow condition. After farrowing, 1.8 kg feed was given daily to each sow plus an additional 0.45 kg for each nursing pig. Individual units were scraped daily, and the entire farrowing facility was cleaned with high pressure cold water and disinfected with alkyl dimethyl benzyl a m m o n i u m chloride between farrowing groups.
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Pigs were regrouped at weaning (4-5 weeks) by weight and transferred to a temperature-controlled (32°C) nursery in separate rooms at the opposite end of the same building as the farrowing unit. Feed consisted of 20% crude protein pellets with whey and molasses introduced at 10-14 days old as a creep feed. At five weeks of age, the temperature in the nursery was lowered each day until it reached 26.6°C at seven weeks of age. Pigs were housed in galvanized wire pens approximately 4.1 × 3 m, 7-10 pigs to a pen, over water pits. Pigs in one pen had nose contact with pigs in a maximum of one other pen.
Sampling procedure Fecal samples were collected weekly from each gilt/sow in the farrowing facility beginning the week prior to farrowing and continuing until the litter was weaned. Twice daily observations of the pigs were made by the herdsman. The original 54 gilts were followed for three lactations. Individual gilts/ sows were removed from the sampling regimen if their litters were stillborn or small in number in which case the neonates were cross-fostered to other dams. Cross-fostered pigs were also removed from the sampling regimen. Fecal samples were transferred to sterile polypropylene or polystyrene tubes. Samples from adult pigs were either freshly voided or obtained per rectum. For suckling pigs, individual (or rarely composite) freshly voided samples were obtained weekly. After weaning, samples were obtained by manual expression from a single different animal each week from each litter. This continued for a period of 3-4 weeks until the animals were moved from the nursery. Thus, each litter had a single sample from a randomly chosen piglet tested for viral antigen weekly during the nursing and postweaning periods. Samples were transported to the laboratory and stored at - 20 ° C until assayed.
Enzyme-linked immunosorbent assay Samples were assayed for the presence ofrotavirus antigen by a solid phase enzyme-linked immunosorbent assay (ELISA Rotazyme II, Abbott, Chicago, IL) designed to detect the group-specific (VP-6) antigen. Polystyrene beads coated with guinea-pig anti-SA- 11 (simian rotavirus) served as capture antibody. It has been determined that SA-11 antibody cross reacts with porcine rotavirus antigens (Benfield et al., 1984). Rabbit anti-SA-11 horseradish peroxidase conjugate and o-phenylenediamine.2HC1 substrate was the indicator system. Absorbance values at 492 nm (Quantum II, Abbott, Chicago, IL) were used as an indicator of virus antigen presence. Positive and negative controls included with the kit were run with each sample and cut off points were determined by a microprocessor which added 0.075 to the optical density reading of the negative control. Specimens yielding equivocal absorbance values were run in duplicate.
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H.B. GELBERG ET AL.
Specificity testing Specificity of the ELISA procedure was ascertained by a blocking assay using unlabeled negative or positive rabbit anti-SA-I 1 antibody (Dakopatts, Santa Barbara, CA) on selected fecal samples (n = 42 ). Those samples which were blocked less than 50% as well as all ELISA-negative samples were tested by negative-contrast electron microscopy (Basgall et al., 1988 ). Electron microscopy was done with coded samples, that is, without knowledge of the ELISA results. As part of an ongoing study, over 400 additional fecal samples from gestating and nursing sows, piglets, and feeder pigs from the same farm were assayed both by commercial ELISA and two monoclonal antibody based microELISA systems to test for the occurrence of false positive reactions by rotazyme. For these latter systems, capture antibody was either polyclonal goat hyperimmune serum or anti-SA- 11. Secondary antibody was monoclonal antiVP6 (Zheng et al., 1989 ). Affinity-purified peroxidase labeled goat anti-mouse antibody (Kirkegaard and Perry Labs, Gaithersburg, MD) and o-phenylenediamine.2HCi was the indicator system. Known positive and negative fecal samples were assayed with each plate. Optical density was determined by spectrophotometry at 490 nm using a micro-ELISA autoreader (MR50, Dynatech Labs, Alexandria, VA). Net optical density (OD) readings greater than 0.2 were considered positive, while those below OD 0.1 were considered negative. Absorbance values between 0.10 and 0.20 were rerun a n d / o r rechecked by another method.
Transmission studies For transmission studies, pairs of caesarean derived pigs were maintained in gnotobiotic isolators and fed a commercial milk replacer (SPF LAC, Borden Inc. ). Each pair of pigs, at 24 h of age, was orally inoculated with 0.5-2 ml of clarified, filtered (0.45/~) feces from a single EEISA positive piglet. There were two fecal samples each from the first, second and third parities. A total of 12 pigs were therefore utilized for the transmission studies. One of each pair of pigs was euthanized at 24 h postinoculation (p.i.) and the other at 48 h p.i. Fecal samples from the gnotobiotic pigs were assayed for group A specific antigen by micro-EEISA. Intestinal samples were fixed in formalin t'or immunohistochemical detection of group A rotavirus antigen in tissue sections.
Immunohistochemistry Tissue sections from seven equidistant regions of small intestine (McAdaragh et al., 1980) were fixed in neutral buffered formalin, processed in low melt paraffin (Peel-A-Way, American Scientific Products, McGaw Park, IL), and sectioned to 5 ,urn thick. They were then processed for immunohis-
SHEDDING OF ROTAVIRUS ANTIGEN
217
tochemical localization of group A specific rotavirus antigen (Gelberg et al., 1990).
RNA electropherotyping RNA extracted from feces of the virus infected gnotobiotic piglets was electropherotyped as described previously (Theft et al., 1981; Gaul et al., 1982; Zheng et al., 1989 ).
Statistical analyses Qualitative data (presence or absence of virus antigen) was compared by Chi squared analysis with a P~< 0.05 denoting significance. Statistical analyses were run on SAS (Statistical Analysis System, Raleigh, NC ) using a Proc. GLM (general linear models procedure). RESULTS Forty-two samples were checked by ELISA, blocking ELISA a n d / o r EM. Twenty-three were confirmed positive for rotavirus antigen. Eighteen samples were negative for rotavirus antigen by ELISA, blocking ELISA and EM. One sample was positive by ELISA and negative by blocking ELISA and EM. We did not have sufficient feces to retest this sample. Overall agreement between Rotazyme and one or both of the validation assays was therefore 97.6% on the 42 samples tested by the three assays. Of the Rotazyme-positive samples, 23 of 24 were confirmed positive by another test (95.8%). Of the 400 samples tested both by commercial ELISA and monoclonal micro-ELISA, only four were reactive by commercial ELISA. These same four samples were positive by monoclonal micro-ELISA. All four of these samples contained small numbers of rotavirus-like particles on EM examination. Diarrhea was rarely observed in the herd under study, was transient, and occurred in one or several nursing animals in a litter. The presence of diarrhea was not correlated with detection of group A specific antigen in feces. During the week prior to parturition, 17 of 44 (38.6%) samples from gilts, 5 of 34 (14.7%) samples from second litter sows, and 19 of 38 (50.0%) samples from third litter sows contained rotavirus antigen (Fig. 1 ). Second lactation sows excreted virus antigen at a significantly ( P = 0 . 0 2 2 ) lower rate than gilts or third lactation sows. Taken as a single farrowing unit, 41 of 116 (35.3%) fecal samples from gilts/sows were positive for rotavirus antigen in the week prior to farrowing (Table 1 ). During the nursing period, gilts shed virus antigen in 49 of 153 (32.0%) samples (Fig. 1 ). During the second lactation, sows shed virus antigen in 45 of 136 (33.1%) samples while nursing and third lactation sows shed virus antigen in 54 of 112 (48.2%) samples while nursing. Second lactation sows excreted virus antigen significantly ( P = 0.004) less often than the other two
218
H.B. GELBERG ET AL.
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Gilts/sows A B C Pigs (nursing) A B C Pigs (weaned) A B C
Weeks postfarrow
Weeks postweaning 1
-1
1
2
3
4
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32 109 29.4
40 111 36.0
42 101 41.6
34 80 42.5
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21 103 20.4
22 116 19.0
25 108 23.1
34 80 42.5
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2
Totals
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3
67 92 72.8
154 219 70.3
ICombined data from three consecutive parities. A: N u m b e r o f samples ELISA positive for group A specific rotavirus antigen. B: N u m b e r of samples assayed. C: Percent antigen positive samples.
groups during week one of lactation and gilts significantly ( P = 0 . 0 0 1 ) less often during week three. Taking the three parities as a single unit, lactating gilts/sows shed virus antigen in 32 of 109 samples (29.4%) during week one, in 40 of 111 (36.0%) samples during week two, in 42 of 101 (41.6%) samples during week three, and in 34 of 80 (42.5%) samples during week four (Table 1 ). Overall, 148 of 401 (36.9%) samples were virus antigen positive.
SHEDDING
OF ROTAVIRUS
ANTIGEN
2 19
Taken as a whole, 38 of 54 (70.4%) gilts excreted virus antigen at some time during the sampling period. For second litter sows, 25 of 49 (51.0%) excreted virus antigen, and 32 of 38 (84.2%) third litter sows excreted virus antigen. Overall, for all farrowing pigs, 95 of 141 (67.4%) demonstrated virus antigen on at least one occasion for each parity. All 37 of the gilts/sows followed for three lactations excreted virus antigen sometime during the study. Pigs nursing gilts excreted virus antigen in 40 of 150 (26.7%) samples (Fig. 2). After weaning, 39 of 69 (56.5%) pig fecal samples from gilts contained virus antigen. Among first lactation pigs, 26 of 53 (49.1%) litters excreted virus antigen sometime during the nursing period, 32 of 41 (78.0%) litters excreted virus antigen during the postweaning period, and 37 of 41 (90.2%) litters excreted virus antigen at least once during the study. Nursing pigs from second litter sows excreted virus antigen in 45 of 138 (32.6%) samples (Fig. 2). Weaned pigs from second lactation sows excreted virus antigen in 53 of 79 ( 59.6% ) samples. Among second litter pigs, 30 of 44 (68.2%) litters excreted virus antigen sometime during nursing 36 of 42 ( 85.7% ) sometime during the postweaning period, and 41 of 42 (97.6%) litters excreted virus antigen sometime during the study. Nursing pigs of third litter sows excreted virus antigen in 17 of 119 (14.3%) samples (Fig. 2). Weaned pigs from third litter sows contained rotavirus antigen in their feces in 62 of 71 (87.3%) samples. Among third lactation pigs, 18 of 38 (47.4%) litters excreted virus antigen sometime during nursing, and 31 of 35 (88.6%) during the postweaning period; 34 of 35 (97.1%) litters excreted virus antigen sometime during the study.
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220
H.B. GELBERGETAL
During the first postweaning week, pigs of gilts excreted virus antigen significantly (P=0.029) less often than the other two groups. Over three parities, pigs while nursing excreted virus antigen in 21 of 103 (20.4%) samples during week one, 22 of 116 (19.0%) samples during week two, 25 of 108 (23.1%) samples during week three, and 34 of 80 (42.5%) samples during week four (Table 1 ). As a single unit, over three parities, suckling pigs excreted virus antigen in 102 of 407 (25.1%) samples. Seventy-four of 135 (54.8%) litters excreted virus antigen sometime during nursing. Taken as a single farrowing unit regardless of the dam's age, weaned pigs excreted virus antigen in 29 of 41 (70.7%) samples during week one, in 58 of 86 (67.4%) samples during week two, and in 67 of 92 (72.8%) samples during week three (Table 1 ). Overall, feces from weaned pigs contained virus antigen in 154 of 219 (70.3%) samples; 99 of 118 (83.4%) litters excreted virus antigen during the postweaning period. Combining data from the three parities into a single farrowing unit, 112 of 118 (94.9%) litters demonstrated virus antigen in their feces sometime during the study. Among all piglet samples, 256 of 626 (40.9%) were virus antigen positive. Weaned pigs excreted virus antigen more often than suckling pigs and there was a steady increase in the percent of pigs excreting antigen with increasing age. A seasonal peak in numbers of gilts/sows excreting virus antigen was observed during the winter months (December through March, P < 0.005 ) and a seasonal trough in late summer and early fall (July through September) (Fig. 3). Nursing pigs excreted virus antigen most often in winter and sum-u- Nursing Pigs + Post-weaning pigs -~- Gilts/Sows 100
80
60
40
20
0
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Fig. _3. Seasonal rotavirus shedding by gilts/sows and piglets.
SHEDDING OF ROTAVIRUS ANTIGEN
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met and least often in spring (P<0.005) while weaned pigs excreted virus antigen more evenly through the year with a significant (P<0.005) peak in early spring and with troughs in April, May, and October (Fig. 3 ). As the rotavirus positive fecal samples had been obtained from subclinically infected animals, the following experiment was conducted to determine the virulence, if any, of the rotavirus present in the population. Two separate fecal samples for each of the three parities was passaged in gnotobiotic piglets. By 48 h p.i., all inoculated animals had watery diarrhea and positive fecal micro-ELISA values. Immunohistochemical evaluation of small intestine showed marked villous epithelial loss and blunting with abundant brown immunohistochemical reaction product in the cytoplasm of surviving enterocytes (Fig. 4 ). These results were similar to previous reports of the pathogenesis of rotavirus infection in pigs (Theil et al., 1978; McAdargh et al., 1980). Negative tissues and negative controls lacked cytoplasmic reaction product. Preincubation of virus with antibody completely blocked specific staining of rotavirus antigen in tissue sections. Viral RNA was extracted from feces of experimentally infected pigs and the electrophoretic banding patterns (Fig. 5) were shown to be similar among samples from the three parities. One sample, from a pig inoculated with feces from parity one that died prior to sacrifice and was severely autolyzed, was micro-ELISA positive and electrophoretically negative. One sample from
Fig. 4. Mid-jejunum from gnotobiotic pig 48 h p.i. with filtered feces from a parity one piglet. Villi are short and blunt. Dark cells at tips of villi are immunoperoxidase positive for group A rotavirus antigen. Hematoxylin counterstain.
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H . B . G E L B E R G E T AL.
A
B
C D
E
F
G
1 ~
2-...3/4 ~--
m
6m
,
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Fig. 5. Electrophoretic migration patterns of viral RNA extracted from feces of newborn, gnolobiotic pigs experimentally inoculated with feces from parity one (lane B), parity two (lanes C and D), and parity three (lanes E and F) pigs. Reference OSU (lanes A and G). Lane B contains a mixture of group A minor variants. Lane D contains, in addition to group A, group C rotavirus RNA. p a r i t y two d e m o n s t r a t e d , in a d d i t i o n to g r o u p A r o t a v i r u s , the presence o f an atypical r o t a v i r u s g e n o t y p e . DISCUSSION In spite o f a large b o d y o f literature d o c u m e n t i n g different aspects o f rotavirus infection a n d disease in m a n y species o f d o m e s t i c animals, little has
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been published about the natural history of rotavirus antigen @edding over time under field conditions. We had opportunity to document; in a newly established closed SPF swine herd, the dynamics of rotavirus antigen shedding both in individual animals as they aged, and in the herd in general. Rotavirus disease is generally observed in the 1-4-week-old age group (k~,ohl et al., 1978) but severe disease has also been reported in animals weaned~at 6-8 weeks of age (Woode, 1986). We were curious to determine whether a newly established closed herd, in newly constructed facilities, would develop enzootic rotavirus infections when populated entirely by adult animals presumably resistant to rotavirus disease. Assuming that virus antigen was introduced into the herd under these conditions, we were also interested in learning how virus antigen spreads through the herd over successive parities, whether there is an electrophoretic change in virus over time, whether the gilts/sows excreted virus antigen more or less often with increasing age and whether offspring of older sows were more or less at risk of infection. The assay used for the detection of virus antigen in this study was a commercial ELISA designed to detect the group specific (VP-6) antigen c o m m o n to all group A rotaviruses. Studies in various species have compared the results of the commercial ELISA used in this study (Rotazyme II) with other means of rotavirus detection (Cornell et al., 1982; Rubenstein and Miller, 1982; Benfield et al., 1984; Morinet et al., 1984; Chernesky et al., 1985; Miotti et al., 1985). Two studies specifically tested pig feces (Cornell et al., 1982; Benfield et al., 1984). In one study the authors found when comparing negative contrast EM with commercial ELISA, that of 157 samples tested, sensitivity of the ELISA was 96% (Cornell et al., 1982). ELISA positive EM negative samples were shown to contain rotavirus antigen by a blocking assay. The results from our limited (42 samples) specificity testing indicated that 23 of 24 positive samples by ELISA were confirmed positive by either EM or blocking ELISA (95.8% ). In the other study involving rotavirus antigen detection in samples from swine, agreement between EM and ELISA was 72% with EM detecting more positive samples than did commercial ELISA (Benfield et al., 1984). In that study, positive cut off values for commercial ELISA were an optical density of >~0.1 at 490 nm. In our study cut off values varied depending on the positive and negative samples included with the kit. They ranged from an OD of 0.073 to 0.144. A total of 22 of 446 ( < 5%) of the positive samples had OD values < 0.1. While EM might have increased the number of rotavirus positive samples from our swine population, it would not determine if ELISAnegative, EM positive samples contained group A rotavirus or morphologically identical, but antigenically distinct, group B or C rotaviruses. Studies have shown that atypical rotaviruses are not detected by Rotazyme (Cornell et al., 1982 ). In a Belgian study, pararotavirus infections were seen in a large proportion of pigs from three different herds (Debouk et al., 1983 ).
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Besides suggesting the possible presence of atypical rotavirus in ELISA negative EM positive samples, the authors of the paper reporting 72% agreement between commercial ELISA and EM postulate that the presence of inhibitors in fecal material that reduces binding of rotavirus to antibody, intestinal luminal antirotavirus antibodies and immune complexes may create false negatives in the enzyme immunoassay as compared with EM (Benfield et al., 1984). False positive reactions were not reported in either of the swine studies, or in our continuing studies assaying 400 sow and piglet samples both by commercial ELISA and confirmatory monoclonal-based micro-ELISAs. These data contrast with a recent study comparing commercial rotavirus assays on human fecal specimens. That study found that while the commercial ELISA used in our studies correctly identified all positive samples, it also produced false positive reactions (Dennehy et al., 1988 ). The reasons for the apparent difference in specificity of commercial ELISA on human versus porcine specimens are not clear but rheumatoid factor-like substances in human stool may be responsible for lowered specificity in that species (Yolken and Stopa, 1979). In a study which monitored rotavirus excretion in experimentally infected gnotobiotic pigs, EM detected virus excretion for a longer period than commercial ELISA (Benfield et al., 1988). In addition, commercial ELISA is directed against SA-11 which is subgroup 1 specific. Thus, while all group A rotaviruses should be detected by this method, subgroup 1 viruses, such as porcine OSU may react more strongly than subgroup 2 porcine rotaviruses accounting for a discrepancy between EM and Rotazyme. This longitudinal study has revealed much about the natural history of rotavirus infections in a well managed SPF swine herd. It has reaffirmed the notion that rotaviruses are ubiquitous, suggests that subclinical infections are commonplace in pigs and that age resistance to rotavirus infection may not occur. Importantly, gilts/sows often shed virus antigen prior to farrowing and during nursing and that it is therefore unrealistic to expect to be able to eliminate rotavirus from an infected swine breeding unit. We believe, based on the fact that all gilts/sows excreted virus antigen sometime during the study, that our results are not spurious and that rotavirus shedding may be a common event even in well-managed swine operations. We agree with others who suggest that sows may play an important role as the source of rotavirus infection of pigs (Benfield et al., 1982). It is postulated that the dam sheds virus at a time when the neonate has large amounts of passive immunity and can best cope with infection (Bohl, 1980). This would explain the subclinical nature of the infection in suckled piglets. Later infections occur in presumably immune pigs, resulting in a lack of clinically detectable disease. In another study (Benfield et al., 1982), 12 sows were sampled every four days beginning five days prior to farrow to 14 days postpartum. Virus shed-
SHEDDING OF ROTAVIRUS ANTIGEN
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ding was determined by EM a n d / o r virus isolation after concentration of the fecal specimens. Five animals (42%) shed virus. Four of 12 litters developed diarrhea but only two litters were from sows who shed virus. In a different study, 33% of postpartum sows in one herd and 10% of postpartum sows from a second herd excreted virus antigen as detected by ELISA (Grom et al., 1984). It appears that the gilts/sows either harbor the organism in a latent form which is activated by hormonal or other changes occurring near parturition; are continuously infected and shed virus antigen throughout their lives (Lecce and King, 1980), or are reinfected. Since serum antirotavirus IgG antibody levels in gilts/sows increase during lactation and decrease between farrowings in the animals under study (Gelberg et al., 1991 ), we suspect that active infections occur in those gilts/sows that excrete antigen. Our infectivity studies indicate that virulent virus was present in the herd, and the results contrast with another study which concluded that reinfection of seropositive sows seemed unlikely (Benfield et al., 1988 ). Although we cannot exclude the possibility that some of the animals shed virus passively following ingestion, but without active infection, we think that this is unlikely. The gnotobiotic piglets experimentally infected with ELISA positive feces became ill and demonstrated histologic and immunohistochemical evidence of virus infection. From these data, we can conclude that the rotaviruses present were virulent strain (s). Viral RNA extracted from the experimental pigs' feces showed minor variations in banding patterns which suggests that they arose from similar viral parents, and suggests that in this closed herd, new rotavirus group A types were not introduced, a n d / o r several strains were cocirculating. Our study suggests the virus to be most prevalent in older (third parity) dams and least prevalent in their nursing pigs. In other studies (Svensmark, 1983; Fu and Hampson, 1987 ) rotavirus infection was most common in pigs born to gilts. Our evidence contradicts other studies suggesting that weaned pigs rarely excrete virus (Debouck et al., 1983 ) and sows never excrete virus (Debouk et al., 1983; Fu and Hampson, 1989). Pigs were weaned at a later age in the former study perhaps accounting for the difference in data. Generally, more litters excreted virus antigen during the postweaning period than during nursing but there was a nearly linear increase in virus antigen excretion with age (Fig. 2). While it is tempting to attribute this increase in antigen excretion to loss of passive immunity, in agammaglobulinemic human patients, human milk caused disappearance of rotavirus from feces with reappearance while the patient was still receiving milk (Saulsbury et al., 1980 ). Likewise, in agammaglobulinemic pigs, age dependent resistance to rotavirus is noted to occur (Kirstein et al., 1985). In children, in addition to the state of immunity, the outcome of rotavirus infection may depend on chronologic age and state of maturity of the gastrointestinal tract which contributes to a
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state of transient virus-host equilibrium, which results in carriage, asymptomatic infection or disease (Champsaur et al., 1984a). In pigs, environmental, social, and physiologic stresses (Kirstein et al., 1985) as well as changes in brush border enzyme activity (Hampson and Kidder, 1986) occurring at weaning may also be contributory to the development of disease a n d / o r an increase in the rate of enterocyte infection and viral antigen shedding. At weaning, villous height decreases and the comparatively short crypts of the pig's distal small intestine elongate providing a large, less mature enterocyte population theoretically more susceptible to malabsorptive diarrheas (Hampson, 1986). Our results question the value of viral diagnosis from feces since the presence of virus antigen in the feces was not correlated with the presence of disease. Our results are similar to a study of Danish swine herds in which 30% of fecal samples from normal animals contained rotaviral antigen (Svensmark, 1983). Likewise, in a study of all children under 2 years of age admitted to a pediatric ward there was a high incidence of asymptomatic rotavirus shedding and no correlation between most virus shedding and diarrhea (Champsaur et al., 1984b). Collectively, gilts/sows shed virus antigen in 189 of 517 (36.6%) fecal samples regardless of the stage of lactation including the week prior to parturition. Thus, virus antigen apparently is distributed throughout the nursery at all times, although the incidence of shedding by gilts/sows is somewhat higher in winter. The reasons for this are not clear. Reports ofrotavirus outbreaks in humans suggest an increased incidence of rotavirus gastroenteritis in cold seasons versus warm seasons (Kapikian et al., 1976; Konno et al., 1978). Seasonal increases in numbers of suckling pigs excreting virus antigen were noted with peak numbers in summer and to a lesser extent in winter as opposed to the gilts/sows whose excretion data peaked in winter. More weaned pigs had virus antigen in their feces in April, May, and October, although peaks and troughs were relatively flat. In a Venezuelan study, seasonal variation in rotavirus excretion by pigs was not seen; rotavirus was not detected in the feces of pigs less than one week of age (we detected virus antigen at three days of age, we did not sample many animals earlier than this) and peaked at 3-4 weeks of age (pigs were followed to 4-6 weeks of age) (Utrera et al., 1984). Virus was detected in 21.3 percent of all diarrheic fecal samples. No virus was detected from control animals. The Venezuelan study was in a relatively stable climatic environment perhaps mimicked best by our postweaning experimental group. In this latter group, the number of samples containing virus antigen was relatively constant. Others have suggested that temperature is not the important variable in virus excretion but relative humidity; dry environments therefore allowing for dust-borne spread of virus (Brandt et al., 1982; Paul and Erinle, 1982). In conclusion, the presence of virus antigen excretion by the gilts/sows and
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the presence of virus antigen in the feces of young pigs suggests that elimination of rotavirus antigen from a swine facility is not an easily attainable goal. In addition, there are at least 4 serotypes of porcine rotavirus (Paul et al., 1988 ), and cross-protection between serotypes is unlikely (Bohl et al., 1984 ). A separate study (Gelberg et al., 1991 ) documents active antibody response and passive transfer of immunity in these experiments. It is possible, based on this study, field investigations, and the work of others (Lecce et al., 1978; McNulty and Logan, 1982), that rotavirus diarrhea develops in those herds with detectable sow immunity because of build-up of rotavirus in the environment permitting the infection to overcome passive immunity. The development of disease versus infection may thus be a function of virus challenge dose. ACKNOWLEDGEMENTS
The authors thank Dr. C. Almgren, Dr. E. Basgall, Dr. E. Cohen, Ms. A. Gast, Mr. E. Parr, and Dr. J. Patterson for technical assistance. This work was supported in part by a grant from the Illinois Department of Agriculture.
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