Bovine viral diarrhea viruses are a group of positive single-stranded RNA viruses that belong to the Pestivirus genus of the Flaviviridae family.1 Bovine viral diarrhea virus is not host specific, and there is evidence that this virus not only infects cattle but also infects other domesticated ruminant species and wildlife. In fact, its distribution may extend to almost all species included in the order Artiodactila.2 Two types of infection are recognized following contact of a naïve animal with BVDV: acute and persistent. Acutely infected animals are believed to be inefficient BVDV transmitters because of the small amounts of virus shed for a short period of time, and these calves eventually clear the virus from their tissues.3 Conversely, PI animals shed a large amount of virus for long periods, making them the main source of secondary infections.4,5 Interestingly, results of a study6 suggest that BVDV circulates in cattle herds without PI animals. Despite many vaccines available and control programs primarily aimed at the detection and removal of PI animals, BVDV is still endemic in some parts of the world. The fact that BVDV still circulates in cattle populations despite all control measures and evidence that BVDV is present in wildlife warrant further investigation of the transmission at the wildlife-livestock interface.
The presence of BVDV in free-roaming or captive wild ruminants has been documented worldwide either by use of serologic surveys or virus isolation.7 In North America, the presence of BVDV in the wild ruminant population has been documented in wapiti (Cervus elaphus), mule deer (Odocoileus hemionus), WTD (Odocoileus virginianus), pronghorn antelope (Antilocapra americana), Canadian bison (Bison bison bison), and moose (Alces alces).8–20 The development of clinical signs caused by BVDV infection in wild ruminants is variable and follows the same course as in cattle.21 For instance, in 1 study,22 WTD fawns did not develop any clinical signs as a result of experimental infection with BVDV; however, they actively shed the virus in nasal excretions for up to 7 days after inoculation.
Only 2 studies23,24 have assessed interspecies transmission of BVDV, focusing on transmission by a PI animal. Both studies found that naïve animals became infected with BVDV by being in contact with PI animals. Furthermore, one of the studies23 found that pregnant WTD coming in contact with PI cattle can result in PI fawns. On the other hand, transmission by acutely infected animals is still a matter of controversy. Studies25–27 assessing transmission by acutely infected animals have focused on transmission in cattle or elk (C elaphus) with contradicting results, and there are no studies assessing the possibility of interspecies transmission by acutely infected animals. Additionally, WTD is the most abundant species of wild ruminants in the United States, and BVDV has been isolated from free-ranging WTD.13 The objective of the study reported here was to assess the feasibility of horizontal transmission of BVDV from acutely infected WTD to cattle by experimentally inoculating WTD fawns and commingling them with naïve calves.
Materials and Methods
Animals—Five 2- to 3-week-old female fawns were purchased from a commercial captive deer farm in Indiana. Fawns were fed a commercial doe milk replacer free of anti-BVDV antibodiesa; feeding intervals varied according to the fawns' ages. Water and creep feed were offered ad libitum starting at 21 days of age and continuing until the conclusion of the study.
Six colostrum-deprived Holstein bull calves were purchased from a large commercial dairy farm in Indiana. At the farm, calves were removed from the dams immediately at birth and placed in separate hutches to prevent colostrum ingestion. Additionally, ear tags were placed and navels were disinfected. Calves were brought to Purdue University Laboratory Animal Housing Facility within 12 hours after birth. For the first 48 hours after birth, calves were fed a human milk replacerb every 12 hours, and thereafter, a medicated commercial calf milk replacerc was given every 12 hours until the end of the study. In addition to the milk replacer, medicated creep feedd was offered ad libitum 7 days after birth until the end of the study.
Virus inoculum—The noncytopathic BVDV-1a strain 544 WTD was used (GenBank accession No. EU597009). This strain was isolated from free-ranging WTD during the Indiana firearm hunting season.14 Virus propagation was performed as described.14
Experimental design—All procedures were approved by the Purdue Animal Care and Use Committee. The number of calves and fawns was mainly restricted by budget limitations. Fawns and calves were housed in the same isolation room under biosafety level 2 at the Purdue University Laboratory Animal Housing Facility. Animals were allowed to acclimate for up to 9 days before the beginning of the study. During this period, blood samples for buffy coat and serum collection were obtained from all animals to test for BVDV types 1 and 2 by means of RT-PCR and VN assays. On arrival, fawns and calves were housed in separate pens. Prior to commingling, all personnel in contact with fawns and calves changed protective clothing, gloves, and boots between handling each species.
As part of the study protocol, starting on arrival day, all calves underwent prophylactic antimicrobial treatment with enrofloxacine (5 mg/kg, SC, q 24 h for 5 days). Additionally, calves received a single dose of 5 g of probiotics POf and a single dose of 3 mL of selenium and vitamin Eg SC.
Following the acclimation period, all fawns were intranasally inoculated (day 0 of study) with 2 mL of noncytopathic BVDV-1a virus suspension (third passage in cell culture) with a titer of 106.7 TCID50/mL. Two days after inoculation, fawns and calves were commingled until the end of the study. Animals were allocated randomly to 5 groups: 1 group included 1 fawn and 2 calves, and 4 groups included 1 fawn and 1 calf. During this period, animals in the same pen shared feed and water sources.
Clinical examinations performed daily in calves and fawns included measurement of rectal temperature and evaluation of attitude, fecal consistency, and the presence of abnormal respiratory tract signs. A clinical scoring system was used to assign numeric values to daily observations as described.28 The following 4 categories were assessed: lethargy, hemorrhage, respiratory tract signs, and diarrhea. At the time of the clinical evaluations, study personnel were unaware of the diagnostic results.
Blood and serum samples were obtained on days −6, 0 (immediately before inoculation), 7, 14, and 21 for buffy coat samples for RT-PCR and VN assays and BVDV-specific antibody ELISA.h Nasal, rectal, and saliva swab specimens were collected on days 0 (immediately before inoculation), 3, 7, 14, 17, and 21 for RT-PCR assay. By 21 days after inoculation, all animals were euthanized by IV administration of a euthanasia solutioni according to label instructions. Following euthanasia, animals were necropsied at the Indiana Animal Disease Diagnostic Laboratory. During postmortem examination, the following samples were collected for histologic evaluation, immunohistochemical analysis, and virus isolation: lymphoid organs (tonsils; retro-pharyngeal, mandibular, and mesenteric lymph nodes; spleen; and thymus), digestive tract (esophagus, rumen, duodenum, jejunum, and Peyer's patches in the jejunum, ileum, colon, and rectum), respiratory tract (trachea and lung), heart, skin, and bone marrow. Two samples were collected from each tissue; the first was fixed in neutral-buffered 10% formalin for histologic evaluation, and the second was frozen at −80°C for virus isolation and possible RT-PCR assay.
Virus isolation—Madin-Darby bovine kidney epithelial cells were prepared in 48-well plates grown in 5% (vol/vol) horse serum,j 20mM l-glutamine,k and an antimicrobial-antimycotic mixture consisting of penicillin (100 U/mL), streptomycin (10 μg/mL), and gentamicin (50 μg/mL). Samples (0.25 mL/well) were inoculated in duplicate on cell suspensions and left for 24 hours before culture medium was removed and replaced with new medium. On day 2 after inoculation, cells in duplicate 48-well plates were fixed after cell culture medium was removed by immersing them in cold 80% aqueous acetone for 10 minutes; cells were then evaluated via immunofluorescence microscopy by use of fluorescein isothiocyanate–labeled antibodiesl–n specific for BVDV.
BVDV RT-PCR assay and sequence analysis—Quantitative real-time PCR assay was performed on serum, nasal, saliva, and rectal swab specimens as described.29 Viral RNA was extracted from appropriate samples by use of a viral RNA extraction kito as recommended by the manufacturer. Real-time PCR assay was performed on clinical samples as described by targeting the 5′-untranslated region of the viral genome. Realtime PCR assay was performed with a RT-PCR kitp in a reaction volume of 25 μL by use of 5 μL of extracted template. Primers were added at a final concentration of 0.4μM each; the probe was used at a final concentration of 0.2μM. For quantification, a 1:10 serial dilution of BVDV type 1 preparations of a known virus titer were used to generate a standard curve. The set of standards was included in each run with clinical samples to determine the validity, relative amount, and reproducibility of the assay. The amount of BVDV in each sample was calculated by converting contact time (ie, Ct) value to virus titer by use of the standard curve.
The viral RNA extracted from tissue samples collected at necropsy that were positive by means of virus isolation was then analyzed to verify the degree of homology to the strain used in this study. The set of primers used in the RT-PCR reaction were 103/326. The amplified PCR products were purified by use of a commercial purification kito according to the manufacturer's protocol. These products were sequenced by use of an automated sequencer at the Purdue University genomic core facilities and analyzed, and their homology to the strain used in the present study and other BVDV strains was determined on the basis of published sequence information and reference control viruses. This analysis was performed by use of computer software.q
Histologic and immunohistochemical evaluation—Tissues were fixed by immersion in neutral-buffered 10% formalin immediately after collection. Fixed tissues were processed, embedded, and sectioned at 5 μm; each section was stained with H&E.
Immunohistochemical analysis for BVDV in tissues was performed by use of BVDV-specific monoclonal antibodies at the Cornell University Diagnostic Laboratory. In brief, tissue sections (thickness, 5 μm) were deparaffinized, rehydrated, and treated with proteinase K. Each section was incubated with optimally diluted BVDV-specific monoclonal antibodies. Antigen-antibody complexes were stained by use of a biotin-strepavidin-diaminobenzidine system. Sections were counterstained with Gill hematoxylin.
VN—Virus neutralization titers against BVDV were determined as follows: test sera were diluted in a 2-fold series from an initial dilution of 1:4 to a maximum dilution of 1:1,024. Next, 50 μL of inoculum containing 200 TCID50 of BVDV types 1 and 2 was added to each well, and the plates were incubated at 37°C for 2 hours. Fifty microliters of suspension of Madin-Darby bovine kidney epithelial cells at 3 × 105 cells/mL was added to each well, and plates were incubated for 3 days at 37°C. Titer was determined via microscopic examination of the monolayer of cells for cytopathic effect. Results were expressed as the reciprocal of the serum 2-fold dilution at which 50% neutralization of virus occured.
Results
Prior to inoculation, 4 fawns and 1 calf became sick from causes other than BVDV infection. One fawn developed oral abscesses that resolved with antimicrobial treatment (20 mg of oxytetracycliner/kg, SC, q 48 h); 3 fawns developed diarrhea, which completely resolved in 2 of them. The fawn with intermittent diarrhea throughout the study was euthanized and necropsied on day 16 because of severe physical deterioration. No parasites were detected in fecal samples collected from the sick fawns. However, fecal swab specimens submitted for bacteriologic testing were positive for Escherichia coli. The sick calf developed septicemia as determined on the basis of hyperfibrinogenimia, neutropenia with left shift, and hypopyon observed in the left eye. Results from samples submitted for bacteriologic and parasitological evaluation were negative for this calf. The clinical score was not considered given the fact that some animals were sick before inoculation day and continued to be sick throughout the study period, making it difficult to associate the clinical score with BVDV infection.
All fawns and calves tested negative for anti-BVDV antibodies in VN and ELISA and buffy coat RT-PCR assays prior to the first day of the study. All fawns had evidence of BVDV infection as early as 3 days after inoculation and shed the virus for up to 18 days as determined on the basis of buffy coat RT-PCR assay and nasal, saliva, and rectal swab specimens (Table 1).
Summary of BVDV RT-PCR assay results in WTD (Odocoileus virginianus) fawns intranasally inoculated with noncytopathic BVDV-1a and commingled with colostrum-deprived Holstein calves.
Days after inoculation (days of cohabitation) | |||||||||||||||||||||||
---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
0 (−2) | 3 (1) | 7 (5) | 14 (12) | 17 (15) | 21 (19) | ||||||||||||||||||
Sample group | Animal | B | N | O | R | N | O | R | B | N | O | R | B | N | O | R | N | O | R | B | N | O | R |
1 | F63* | − | − | − | − | + | − | + | + | + | + | − | + | + | + | − | + | + | − | + | n | n | n |
C53 | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | + | − | − | − | |
C91 | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | |
2 | F1504 | − | − | − | − | + | − | − | + | + | − | + | + | − | − | + | − | − | − | − | − | − | − |
C51 | − | − | − | − | − | − | − | − | − | − | − | + | − | − | − | + | − | − | + | − | − | − | |
3 | F56 | − | − | − | − | + | + | − | + | + | − | + | − | − | − | − | − | − | − | − | − | − | − |
C50 | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | |
4 | F1526 | − | − | − | − | + | − | − | + | + | − | + | + | + | − | + | + | + | + | + | + | + | − |
C92 | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | − | + | − | + | − | |
5 | F61 | − | − | − | − | + | − | + | + | + | − | + | + | − | + | − | − | − | − | − | − | − | − |
C52 | − | − | − | − | − | − | − | + | − | − | − | + | − | − | − | − | − | − | + | − | − | − |
Euthanized by day 16 after inoculation. Samples collected at necropsy included tissues, buffy coat, and nasal, oral, and rectal swab specimens.
− = Negative result. + = Positive result. B = Buffy coat. C = Calf. F = Fawn. n = Not done. N = Nasal swab specimen. O = Oral swab specimen. R = Rectal swab specimen.
Following cohabitation, calves and fawns were commonly seen sharing the same pen area. Four of 6 calves had positive results of buffy coat RT-PCR assay for BVDV. Virus was detected in the buffy coat in one of the calves as early as 5 days and as late as 18 days after cohabitation (Table 1). Only one calf had evidence of the virus in a nasal swab specimen, and another calf had evidence of the virus in 1 oral swab specimen.
Four of the 5 fawns had positive results of VN assays and positive results of the BVDV-specific antibody ELISA. One fawn had positive results of ELISA by day 14 but was seronegative via VN testing. One fawn that was euthanized on day 16 had positive results of ELISA but was seronegative via VN testing. By 21 days after inoculation, the remaining 3 fawns developed VN titers ranging from 1:4 to 1:8 and had positive results of ELISA. Only the calf that had positive results of PCR assay for BVDV after 5 days of cohabitation developed antibodies against BVDV as determined on the basis of positive results of ELISA.
Virus was isolated from the intestines, lungs, or pooled lymph nodes in 3 of the 5 fawns and in 4 calves 21 days after inoculation (19 days after cohabitation; Table 2). The RNA from these BVDV-positive tissues from the fawns and calves was 99.9% homologous to the strain used in this study as determined on the basis of analysis of the 5′-untranslated region.
Summary of virus isolation results in the same animals as in Table 1.
Group | Animal | Intestine | Pooled lymph nodes | Lung |
---|---|---|---|---|
1 | F63 | + | − | − |
C53 | + | + | − | |
C91 | − | − | − | |
2 | F1504 | − | − | − |
C51 | + | − | − | |
3 | F56 | − | − | − |
C50 | − | − | − | |
4 | F1526 | − | + | + |
C92 | + | + | + | |
5 | F61 | − | + | − |
C52 | + | − | + |
See Table 1 for key.
On necropsy, no gross lesions were identified in any of the animals. Histologically, all BVDV-infected fawns and calves had marked lymphoid atrophy in the Peyer's patches. No other lesions characteristic of BVDV were observed. Because of budget restrictions, immunohistochemical analysis was performed on an ileum specimen from only one of the BVDV-infected calves. There was positive labeling of BVDV antigen as evidenced by low numbers of scattered positive-staining cells in areas of lymphoid depletion and necrosis and in the lamina propia of villi.
Discussion
Neither fawns nor calves developed any clinical signs related to the infection; this is consistent with results obtained in our laboratory by use of the same BVDV strain.22,30 However, 4 fawns were sick within 24 to 48 hours after arrival: 1 fawn developed oral lesions, and 3 fawns developed diarrhea, which resolved in 2 of them. The diarrhea may have been caused by stress from transportation. Escherichia coli was cultured from fecal swab specimens of affected fawns. The calf that was sick 1 week after arrival most likely had septicemia as a result of colostrum deprivation. However, parasitological and bacteriologic results were negative (fecal swab specimens), likely because of antimicrobial use prior to culturing.
All fawns were successfully infected with BVDV following inoculation and were actively shedding the virus as early as 3 days after inoculation and for as long as 18 days in feces, nasal secretions, or oral secretions. The 2 fawns that were sick throughout the study had more severe viremia and shed higher quantities of virus for a longer period of time (data not shown). Concurrent infections in 2 of these fawns may have potentiated the effects of BVDV infection, enhancing the duration of the viremia and amount of virus excreted. In a previous study,31 BVDV has acted synergistically with bovine rotavirus, worsening the clinical signs in dual-infected calves and resulting in increased BVDV replication. Under natural conditions, concurrent infections may have an effect on BVDV shedding, as in the present study.
Four of the 6 calves were infected with BVDV as a result of direct contact with the fawns as evidenced by the presence of the virus in the buffy coat and in tissues collected at necropsy. Studies assessing BVDV transmission from acutely infected animals to in-contact animals are few. One study27 found that elk in contact with acutely infected elk actively shedding BVDV resulted in infection of the in-contact animals. Similar to our results, the in-contact elk developed viremia; however, there was no evidence of shedding of the virus following infection. Conversely, 2 BVDV transmission studies26,32 between acutely infected calves and naïve calves failed to reveal successful transmission. In both studies, evidence of infection was based on seroconversion and not virus detection.
The presence of antibodies in 4 fawns coincided with clearance of the virus and the inability to isolate the virus from tissues at necropsy. Only the calf with evidence of infection by 5 days after cohabitating with the fawns developed antibody titers against BVDV by 21 days after inoculation as determined on the basis of ELISA results. There is a possibility that the infected calves did not have enough time to develop antibodies, although seroconversion usually occurs within 14 to 30 days after infection.33 Microscopic findings agreed with previous studies22,27,34,35 in which the primary histologic lesions observed were lymphoid depletion of Peyer's patches and thymic atrophy. Compared with previous studies34,36,37 that used virulent BVDV strains, the lack of clinical signs and the paucity of lesions in the present study might be attributable to the low virulence of the strain used.
Colostrum-deprived calves were used in the present study for various reasons. In studies in which animals are challenge inoculated with infectious agents, the susceptibility to infection of those animals should be the same. Because of budget constraints and the fact that approximately 75% of dairy farmers in the United States vaccinate their cattle against BVDV,38 it was almost impossible to obtain calves that were free of anti-BVDV antibodies and that were given colostrum.
The authors are aware that this experimental setting did not mimic field conditions. In previous studies, one regional39 and another national,40 authors reported that approximately 50% of farmers observed either direct wildlife contact or wildlife contact with cattle feed sources, which gives external validity to the present study. Furthermore, the degree of contact between cattle and wildlife varies on the basis of feed and water availability, management systems, and animal density.41 Therefore, epidemiologically, the importance of these findings stem from the detection of virus shedding into the environment by both species, which potentially can lead to the infection of a pregnant animal and hence the spread and perpetuation of the virus among both populations. Nevertheless, it should be noted that the latter largely depends on the amount and duration of virus shedding, the infectious dose, duration of virus survival in the environment, population density, and contact frequency between livestock and wildlife.
To the best of our knowledge, this is the first study on transmission of BVDV from acutely infected deer to livestock. In this study, BVDV-infected WTD infected naïve calves with BVDV-1a when commingled together for 21 days. On the basis of these findings, wildlife acutely infected with BVDV may be a potential source of infection for susceptible cattle. Field investigations to determine the extent that wild animals contribute to the propagation of this disease would be informative.
ABBREVIATIONS
BVDV | Bovine viral diarrhea virus |
PI | Persistently infected |
RT | Reverse transcriptase |
VN | Virus neutralization |
WTD | White-tailed deer |
Zoologic Doe Milk Replacer, Pet Ag, Hampshire, Ill.
Parent's Choice, PBM Nutritionals, Georgia, Vt.
Nurse Chow 100, Purina, St Louis, Mo.
Calf Startena, Purina, St Louis, Mo.
Baytril, Bayer Health Care LLC, Shawnee Mission, Kan.
Probios, Bomac Vet Plus, Knapp, Wis.
BO-SE, Intervet/Shering-Plough Animal Health, Millsboro, Del.
IDEXX HerdCheck BVDV antibody ELISA, IDEXX, Westbrook, Me.
Fatal-Plus, Vortech Pharmaceuticals, Dearborn, Mich.
Sigma Chemical Co, St Louis, Mo.
Gibco/BRL Life Science, Grand Island, NY.
National Veterinary Services Laboratory, Ames, Iowa.
American Bioresearch Inc, Seymour, Tenn.
VMRD Inc, Pullman, Wash.
QIAamp viral RNA extraction kit, Qiagen Inc, Santa Clarita, Calif.
QuantiTect Probe RT-PCR kit, Qiagen Inc, Santa Clarita, Calif.
DNASTAR software, DNA Star Inc, Madison, Wis.
LA-200, Pfizer Animal Health, Exton, Pa.
References
- 1.↑
Thiel HJ, Collet MS, Gould EA, et al. Flaviviridae. In: Fauquet CM, Mayo MA, Maniloff J, et al, eds. Virus taxonomy: VIIIth report of the International Committee on Taxonomy of Viruses. San Diego: Elsevier Academic Press, 2005;981–998.
- 2.↑
Becher P, Orlich M, Kosmidou A, et al. Genetic diversity of pestiviruses: identification of novel groups and implications for classification. Virology 1999; 262:64–71.
- 3.↑
Lindberg A, Houe H. Characteristics in the epidemiology of bovine viral diarrhea virus (BVDV) of relevance to control. Prev Vet Med 2005; 72:55–73.
- 4.
Brock KV, Redman DR, Vickers ML, et al. Quantitation of bovine viral diarrhea virus in embryo transfer flush fluids collected from a persistently infected heifer. J Vet Diagn Invest 1991; 3:99–100.
- 5.
Kirkland PD, Mackintosh SG, Moyle A. The outcome of widespread use of semen from a bull persistently infected with pestivirus. Vet Rec 1994; 135:527–529.
- 6.↑
Moen A, Sol J, Sampimon O. Indication of transmission of BVDV in the absence of persistently infected (PI) animals. Prev Vet Med 2005; 72:93–98.
- 7.↑
Van Campen H, Frölich K, Hofmann M. Pestivirus infection. In: Williams ES, Barker IK, eds. Infectious diseases of wild mammals. 3rd ed. Ames, Iowa: Blackwell Publishing, 2001;232–244.
- 8.
Aguirre A, Hansen DE, Starkey EE, et al. Serologic survey of wild cervids for potential disease agents in selected national parks in the United States. Prev Vet Med 1995; 21:313–322.
- 9.
Chase CCL, Braun LJ, Leslie-Steen P, et al. Bovine viral diarrhea virus multiorgan infection in two white-tailed deer in southeastern South Dakota. J Wildl Dis 2008; 44:753–759.
- 10.
Davidson WR, Crow CB. Parasites, diseases, and health status of sympatric populations of Sika deer and white-tailed deer in Maryland and Virginia. J Wildl Dis 1983; 19:345–348.
- 11.
Duncan C, Van Campen H, Soto S, et al. Persistent bovine viral diarrhea virus infection in wild cervids of Colorado. J Vet Diagn Invest 2008; 20:650–653.
- 12.
Kocan AA, Franzmann AW, Waldrup KA, et al. Serologic studies of select infectious diseases of moose (Alces alces L.) from Alaska. J Wildl Dis 1986; 22:418–420.
- 13.↑
Passler T, Walz PH, Ditchkoff SS, et al. Evaluation of hunter-harvested white-tailed deer for evidence of bovine viral diarrhea virus infection in Alabama. J Vet Diagn Invest 2008; 20:79–82.
- 14.↑
Pogranichniy RM, Raizman E, Thacker HL, et al. Prevalence and characterization of bovine viral diarrhea virus in the white-tailed deer population in Indiana. J Vet Diagn Invest 2008; 20:71–74.
- 15.
Stauber EH, Autenrieth R, Markham OD, et al. A seroepide-miologic survey of three pronghorn (Antilocapra americana) populations in southeastern Idaho, 1975–1977. J Wildl Dis 1980; 16:109–115.
- 16.
Stauber EH, Nellis CH, Magonigle RA, et al. Prevalence of reactors to selected livestock pathogens in Idaho mule deer. J Wildl Manage 1977; 41:515–519.
- 17.
Van Campen H, Ridpath J, Williams E, et al. Isolation of bovine viral diarrhea virus from a free-ranging mule deer in Wyoming. J Wildl Dis 2001; 37:306–311.
- 18.
Vanleeuwen JA, Keefe GP, Tremblay R, et al. Seroprevalence of infection with Mycobacterium avium subspecies paratuberculosis, bovine leukemia virus, and bovine viral diarrhea virus in maritime Canada dairy cattle. Can Vet J 2001; 42:193–198.
- 19.
Wolf KN, DePerno CS, Jenks JA, et al. Selenium status and antibodies to selected pathogens in white-tailed deer (Odocoileus virginianus) in southern Minnesota. J Wildl Dis 2008; 44:181–187.
- 20.
Deregt D, Tessaro SV, Baxi MK, et al. Isolation of bovine viral diarrhoea viruses from bison. Vet Rec 2005; 157:448–450.
- 21.↑
Ridpath JF, Driskell EA, Chase CCL, et al. Reproductive tract disease associated with inoculation of pregnant white-tailed deer with bovine viral diarrhea virus. Am J Vet Res 2008; 69:1630–1636.
- 22.↑
Raizman EA, Pogranichniy R, Levy M, et al. Experimental infection of white-tailed deer fawns (Odocoileus virginianus) with bovine viral diarrhea virus type-1 isolated from free-ranging white-tailed deer. J Wildl Dis 2009; 45:653–660.
- 23.↑
Passler T, Walz PH, Ditchkoff SS, et al. Cohabitation of pregnant white-tailed deer and cattle persistently infected with diarrhea virus results in persistently infected fawns. Vet Microbiol 2009; 134:362–367.
- 24.
Uttenthal Å, Hoyer MJ, Grøndahl C, et al. Vertical transmission of bovine viral diarrhoea virus (BVDV) in mousedeer (Tragulus javanicus) and spread to domestic cattle. Arch Virol 2006; 151:2377–2387.
- 25.
Niskanen R, Lindberg A, Larsson B, et al. Lack of virus transmission from bovine viral diarrhoea virus infected calves to susceptible peers. Acta Vet Scand 2000; 41:93–99.
- 26.
Niskanen R, Lindberg A, Tråvén M. Failure to spread bovine virus diarrhoea virus infection from primarily infected calves despite concurrent infection with bovine coronavirus. Vet J 2002; 163:251–259.
- 27.↑
Tessaro SV, Carman PS, Deregt D. Viremia and virus shedding in elk infected with type 1 and virulent type 2 bovine viral diarrhea virus. J Wildl Dis 1999; 35:671–677.
- 28.↑
Cortese VS, West KH, Hassard LE, et al. Clinical and immunologic responses of vaccinated and unvaccinated calves to infection with a virulent type-II isolate of bovine viral diarrhea virus. J Am Vet Med Assoc 1998; 213:1312–1319.
- 29.↑
Hoffmann B, Depner K, Schirrmeier H, et al. A universal heterologous internal control system for duplex real-time RT-PCR assays used in a detection system for pestiviruses. J Virol Methods 2006; 136:200–209.
- 30.
Raizman EA, Pogranichniy R, Lévy M, et al. Experimental infection of colostrum deprived calves with bovine viral diarrhea virus type 1a isolated from free-ranging white-tailed deer (Odocoileus virginianus). Can J Vet Res 2011; 75:65–68.
- 31.↑
Kelling CL, Steffen DJ, Cooper VL, et al. Effect of infection with bovine viral diarrhea virus alone, bovine rotavirus alone, or concurrent infection with both on enteric disease in gnotobiotic neonatal calves. Am J Vet Res 2002; 63:1179–1186.
- 32.
Niskanen R, Lindberg A, Larsson B, et al. Lack of virus transmission from bovine viral diarrhoea virus infected calves to susceptible peers. Acta Vet Scand 2000; 41:93–99.
- 33.↑
Kelling CL, Hunsaker BD, Steffen DJ, et al. Characterization of protection from systemic infection and disease by use of a modified-live noncytopathic bovine viral diarrhea virus type 1 vaccine in experimentally infected calves. Am J Vet Res 2005; 66:1785–1791.
- 34.
Kelling CL, Steffen DJ, Topliff CL, et al. Comparative virulence of isolates of bovine viral diarrhea virus type II in experimentally inoculated six- to nine-month-old calves. Am J Vet Res 2002; 63:1379–1384.
- 35.
Pedrera M, Gomez-Villamandos JC, Romero-Trevejo JL, et al. Apoptosis in lymphoid tissues of calves inoculated with noncytopathic bovine viral diarrhea virus genotype-1: activation of effector caspase-3 and role of macrophages. J Gen Virol 2009; 90:2650–2659.
- 36.
Liebler-Tenorio EM, Ridpath JE, Neill JD. Distribution of viral antigen and development of lesions after experimental infection with highly virulent bovine viral diarrhea virus type 2 in calves. Am J Vet Res 2002; 63:1575–1584.
- 37.
Odeon AC, Kelling CL, Marshall DJ, et al. Experimental infection of calves with bovine viral diarrhea virus genotype II (NY-93). J Vet Diagn Invest 1999; 11:221–228.
- 38.↑
USDA APHIS. NAHMS Dairy 2007 info sheet: bovine viral diarrhea (BVD) management practices and detection in bulk tank milk in the United States, 2007. Available at: nahms.aphis.usda.gov/dairy/dairy07/Dairy07_is_BVD.pdf. Accessed Apr 20, 2010.
- 39.↑
USDA APHIS. NAHMS Dairy 2007 Part I: reference of dairy cattle health and management practices in the United States. Available at: nahms.aphis.usda.gov/dairy/dairy07/Dairy2007_Part_I.pdf. Accessed Aug 15, 2009.
- 40.↑
Raizman EA, Wells SJ, Jordan PA, et al. Mycobacterium avium subsp. paratuberculosis from free-ranging deer and rabbits surrounding Minnesota dairy herds. Can J Vet Res 2005; 69:32–38.
- 41.↑
Van Campen H, Rhyan J. The role of wildlife in diseases of cattle. Vet Clin North Am Food Anim Pract 2010; 26:147–161.