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Brief Communication |
Correspondence: 1Corresponding Author: Jason W Brooks, Animal Diagnostic Laboratory, The Pennsylvania State University, Orchard Road, University Park, PA 16802, e-mail: jwb21{at}psu.edu
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Abstract |
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Key Words: Bovine viral diarrhea virus • Odocoileus virginianus • PCR • persistent infection • white-tailed deer
Bovine viral diarrhea virus (BVDV) is an enveloped, single-stranded, positive-sense RNA virus of the genus Pestivirus of the family Flaviviridae.6 Other members of the genus include classical swine fever (hog cholera) virus, border disease virus, and tentative pestiviruses of the giraffe and pronghorn antelope.6,9,17 Currently, two BVDV genotypes are recognized: BVDV-1 and BVDV-2.6 Classically, pestiviruses were differentiated according to the species in which they were identified. It is now known that infections may occur across species; thus, genetic methods of classification are used.9 The genome of pestiviruses contains a single, large, open-reading frame that is flanked by 5' and 3' untranslated regions (UTRs).17 Because the 5' UTR represents the portion of the genome with the greatest sequence homology among pestivirus species, phylogenetic segregation is often based on analysis of this region.6,17
Bovine viral diarrhea virus most commonly infects cattle; however, sheep, goats, pigs, and various wild ruminants may also become infected.13,16,17 In cattle, infection with BVDV can result in immunosuppression, diarrhea, poor reproductive efficiency, abortion, congenital anomalies, mucosal disease, or birth of persistently infected carrier animals.12,13,15 Persistently infected cattle are the major source of BVDV as they continuously shed large quantities of virus into the environment; virus is also shed in lower amounts from acutely infected animals for several days after infection.10 Currently, there is one report of possible BVDV persistent infection in white-tailed deer (Chase CCL, Braun LJ, Leslie-Steen P, et al.: 2004, Evidence of bovine viral diarrhea virus persistent infection in two white-tailed deer in Southwestern South Dakota. AABP Proceedings, September 2004). It remains uncertain whether persistent infections occur in cervids and with what frequency.
In addition to domestic ruminants and swine, BVDV-like pestiviruses have been isolated from several species of deer and various wild and exotic ruminants.5,7,11,16,17 Pestiviruses have been detected by polymerase chain reaction (PCR) in 2 farm-raised Rocky Mountain elk (Cervus elaphus nelsoni) and a farm-raised white-tailed deer (Odocoileus virginianus) in Pennsylvania in 2004 (authors, unpublished data). Antibodies to BVDV have been detected in numerous wild and exotic ruminants in North America, Europe, and Africa.1,5,8,11,15 There are few reports of experimental infections of cervids with BVDV.13,14 In these reports minimal or no clinical disease was observed, although numerous animals became viremic, seroconverted, and/or shed virus. It has been suggested that cervids are susceptible to infection with BVDV but that they seldom develop disease.13,15 As BVDV and other pestiviruses are not strictly host specific, it is important to know whether populations of wild or farm-raised cervids are susceptible to infection with BVDV, are reservoirs of BVDV, and serve as sources of infection for domestic cattle.
According to the most recent survey of licensed deer farming operations, the deer farming industry is rapidly growing in Pennsylvania (Rosenberry C, Boyd R, Houghton G: June 2003, 2002 White-tailed deer and elk propagator inventory. Pennsylvania Game Commission). As of June 2003, this survey estimated that there were 25,600 captive cervids (largely white-tailed deer and elk) kept on 743 premises in Pennsylvania. It is important to the deer-farming industry to understand the significance of BVDV infections in deer populations. The objective of this study was to determine if BVDV was present in farm-raised Pennsylvania white-tailed deer submitted for necropsy. To the authors' knowledge, this is the first report of screening of necropsy specimens from farm-raised deer for BVDV.
Between January 1, 2005, and December 31, 2005, the carcasses of 61 farm-raised white-tailed deer were submitted for necropsy to the Animal Diagnostic Laboratory of The Pennsylvania State University. Ages of the animals ranged from 3 days to 12 years, with a mean of 2.1 years and a median of 1.3 years (SEM = 0.32 years). Of the animals for which sex was recorded (n = 56), 31 (55.4%) were male and 25 (44.6%) were female. Deer originated from 16 counties within the state of Pennsylvania (39°43'–42°N, 74°43'–80°31'W). All animals were submitted because of natural death of unknown cause or following euthanasia because of severe illness.
Necropsy was routinely performed on each carcass. Representative sections of all major organs—including brain, heart, lung, liver, spleen, kidney, small intestine, large intestine, mesenteric lymph node, and skeletal muscle—were fixed in 10% neutral-buffered formalin, processed by standard methods, stained with hematoxylin and eosin, and examined by light microscopy for microscopic changes.
Single-tube real-time reverse transcriptase PCR (real-time RT-PCR) for the simultaneous detection of BVDV-1 and BVDV-2 was performed using the Rotor-Gene 3000 Q PCR Thermocycler.a,2 Briefly, an organ pool containing fresh lung, spleen, kidney, and mesenteric lymph node and one pool of ileum and spiral colon from each carcass was processed for RNA extraction using TRIzol.b Each 50 µL reaction was composed of 10X Taq Gold PCR buffer, 4 mM MgCl2, 250 mM of a blend of deoxyribonucleotide triphosphates with 2'-deoxyuridine 5'-triphosphate, 1 unit AmpliTaq Gold Polymerase, 10 units MultiScribe Reverse Transcriptase, and 20 units RNAse inhibitor.c Two hundred nM of each of the following PCR primers were used in the reactions: BVDV-1 and BVDV-2 forward 5'/TAGCCATGCCCTTAGTAGGAC/3', BVDV-1 reverse 5'/GACGACTACCCTGTACTCAGG/3', and BVDV-2 reverse 5'/GACGAGTCCCCTGTACTCAGG/3'. One hundred nM each of 2 dual-labeled probes were included in the reaction for the typing of BVDV-1 and BVDV-2. Probe sequences labeled with distinct fluorescent reporter and quencher dyes were BVDV-1 5'/FAM/AACAGTGGTGAGTTCGTTGGATGGCTT/BHQ1/3' and BVDV-2 5'/HEX/TAGCAGTGAGTCCATTGGATGGCCGA/BHQ1/3'. Reverse transcription was performed for 30 minutes at 48°C followed by 10 minutes at 95°C. Forty-five cycles of 25 seconds at 95°C and 60 seconds at 60°C were performed immediately after reverse transcription.
Tissues for virus isolation were collected from each carcass and frozen at –80°C. A virus isolation procedure was performed on one pool of fresh frozen lung, spleen, kidney, and mesenteric lymph node and one pool of fresh frozen ileum and spiral colon from samples with partial amplification by real-time RT-PCR. The tissues were homogenized in viral transport medium containing antibiotics and centrifuged at 2060 x g for 10 minutes. The supernatant was filtered using 0.2 µm membrane filter and inoculated onto 25 cm2 tissue culture flasks and 4-chambered tissue culture slides of Madin-Darby bovine kidney cells (MDBK) cells. Maintenance medium used was 2% fetal bovine serum (BVDV-free based on virus culture, immunoperoxidase staining of cell monolayers, and real-time RT-PCR) in minimal essential medium (MEM) with antibiotics (kanamycin, gentamicin, penicillin, streptomycin, amphoterocin).4 Positive cell-culture controls consisted of MDBK cells inoculated with noncytopathic BVDV-NY-1. Cell culture flasks were incubated in 5% CO2 at 37°C and observed for cytopathic effects for 6 days. The 4-chambered tissue culture slides were fixed in acetone after 3 days of incubation and stained with fluorescent antibody (FA) conjugated to detect the presence of BVDV. Two serial cell-culture passages were performed in both the 25 cm2 flasks and the 4-chambered tissue culture slides. After both the first and second passages, the cells were examined by fluorescent microscopy in an indirect immunofluorescent antibody method.
A 1 cm by 2 cm full-thickness section of pinna was collected from each carcass and placed in 10% neutral-buffered formalin for immunohistochemical (IHC) staining procedure to identify BVDV antigen. Tissues were processed as for routine histopathology and 3-µm sections of tissue were floated onto a 42°C water bath, mounted on charged slides, and allowed to dry overnight. After deparaffinization, slides were rinsed 3 times with water, incubated for 15 minutes at 37°C with 0.05% protease,e and rinsed 3 times with water. Immunohistochemical analysis was performed on a DAKO Autostainer Universal Staining Systemf using the Zymed Histostain Plus Mouse Primary kitg and mouse origin primary monoclonal antibody 15C5h diluted to 1:200 in phosphate-buffered saline–Brij buffer at pH 7.5. Slides were preserved with Crystal/Mounti and were examined by light microscopy for the presence of BVDV antigen. Positive IHC controls consisted of bovine ear-notch tissue positive for BVDV by IHC.
Real-time RT-PCR assay was performed on 122 tissue pools from the 61 animals tested. All tissue samples (n = 92) from 46 of the 61 deer were negative for both BVDV-1 and BVDV-2 RNA by real-time RT-PCR. At least one sample from each of the 15 remaining animals showed evidence of minimal amplification; these were considered for further evaluation. Frozen tissues from these 15 deer and from 10 of the 46 PCR-negative deer were subjected to virus isolation as described. After 2 serial passages and evaluation by FA, all samples were found to be negative for BVDV. Thus, the PCR results from the 15 remaining deer were considered to be attributable to nonspecific amplification. All animals were classified as negative for BVDV-1 and BVDV-2.
Evaluation of skin biopsies stained for BVDV by IHC was performed on 56 of the 61 carcasses. All skin samples tested negative for BVDV antigen. Tissues from the remaining 5 animals were not available for testing. No gross or histologic lesions suggestive of BVDV infection, as reported for experimental infections with BVD virus of deer or cattle,13,14,18 were detected in any of the 61 animals.
Bovine viral diarrhea virus was not detected in any deer in this study, suggesting that fatal acute and persistent BVDV infections are uncommon in farm-raised white-tailed deer in Pennsylvania. The samples in this study were collected from deer that died of unknown causes, frequently after a period of illness. Based on the previous detection of BVDV by PCR in farm-raised elk and deer, it was hypothesized that some cases may have been the result of acute or persistent BVDV infections. With a sample size of 61 animals out of an estimated population of 25,000, by using this real-time RT PCR assay the authors can conclude with 95% confidence that the prevalence of BVDV in the necropsy population of captive white-tailed deer is not greater than 1.0%.j,3 Assuming that the presence of BVDV is positively associated with the risk of mortality in captive deer, infected animals would more likely be detected in our necropsy population than in animals randomly selected from the captive population. These findings suggest that although cervids are susceptible to infection with BVDV, as demonstrated in previous studies,13,14,15 BVDV is not a common cause of mortality in Pennsylvania farm-raised white-tailed deer, and that the prevalence of infection in the general population is less than 1%.
The prevalence of BVDV exposure and clearance of the infection in farm-raised cervids can be determined by measuring serum antibody levels. As this study was conducted on a necropsied population, testable serum samples were not available. Further evaluation of samples collected from live animals would be required to determine whether natural infection with BVDV is associated with an increased risk of clinical or subclinical disease.
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Acknowledgments |
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TOPSources and manufacturersAbstractReferences |
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a. Corbett Life Science, Sydney, Australia.
c. Applied Biosystems, Foster City, CA.
d. American BioResearch, Sevierville, TN.
e. Sigma-Aldrich, St. Louis, MO.
f. DakoCytomation, Carpinteria, CA.
g. Zymed Laboratories Inc., San Francisco, CA.
h. Cornell University, Ithaca, NY.
j. FreeCalc Version 2, AusVet Animal Health Services, Wentworth Falls, NSW, Australia.
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