Response to experimentally induced infection with bovine respiratory syncytial virus following intranasal vaccination of seropositive and seronegative calves

John A. Ellis Department of Veterinary Microbiology, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK S7N 5B4, Canada.

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Sheryl P. Gow Department of Large Animal Clinical Sciences, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK S7N 5B4, Canada.

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Noriko Goji Department of Veterinary Microbiology, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK S7N 5B4, Canada.

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Abstract

Objective—To determine whether a combination modified-live bovine respiratory syncytial virus (BRSV) vaccine can stimulate protective immunity in young BRSV-seropositive calves following intranasal (IN) administration.

Design—Controlled challenge study.

Animals—66 Holstein bull calves, 3 to 8 days old.

Procedures—In experiment 1, BRSV-seropositive and -seronegative calves were vaccinated IN with a commercially available combination modified-live virus vaccine formulated for SC administration; calves underwent BRSV challenge 4.5 months later. In experiment 2, BRSV-seronegative calves were vaccinated IN or SC (to examine the effect of route of administration) with the same combination vaccine that instead had a 1/100 dose of BRSV (to examine the effect of dose); calves underwent BRSV challenge 21 days later.

Results—In experiment 1, BRSV challenge resulted in severe respiratory tract disease with low arterial partial pressures of oxygen and lung lesions in most calves from all groups. Maximum change in rectal temperature was significantly greater in seropositive IN vaccinated calves, compared with seronegative IN vaccinated and seropositive control calves. Number of days of BRSV shedding was significantly lower in seronegative IN vaccinated calves than in seropositive IN vaccinated and seropositive control calves. In experiment 2, maximum change in rectal temperature was significantly greater in seronegative control calves, compared with seronegative IN and SC vaccinated calves. Shedding of BRSV was significantly reduced in seronegative IN and SC vaccinated calves, compared with control calves; also, lung lesions were reduced in seronegative IN and SC vaccinated calves.

Conclusions and Clinical Relevance—Maternal antibodies may inhibit priming of protective responses by IN delivered BRSV vaccines.

Abstract

Objective—To determine whether a combination modified-live bovine respiratory syncytial virus (BRSV) vaccine can stimulate protective immunity in young BRSV-seropositive calves following intranasal (IN) administration.

Design—Controlled challenge study.

Animals—66 Holstein bull calves, 3 to 8 days old.

Procedures—In experiment 1, BRSV-seropositive and -seronegative calves were vaccinated IN with a commercially available combination modified-live virus vaccine formulated for SC administration; calves underwent BRSV challenge 4.5 months later. In experiment 2, BRSV-seronegative calves were vaccinated IN or SC (to examine the effect of route of administration) with the same combination vaccine that instead had a 1/100 dose of BRSV (to examine the effect of dose); calves underwent BRSV challenge 21 days later.

Results—In experiment 1, BRSV challenge resulted in severe respiratory tract disease with low arterial partial pressures of oxygen and lung lesions in most calves from all groups. Maximum change in rectal temperature was significantly greater in seropositive IN vaccinated calves, compared with seronegative IN vaccinated and seropositive control calves. Number of days of BRSV shedding was significantly lower in seronegative IN vaccinated calves than in seropositive IN vaccinated and seropositive control calves. In experiment 2, maximum change in rectal temperature was significantly greater in seronegative control calves, compared with seronegative IN and SC vaccinated calves. Shedding of BRSV was significantly reduced in seronegative IN and SC vaccinated calves, compared with control calves; also, lung lesions were reduced in seronegative IN and SC vaccinated calves.

Conclusions and Clinical Relevance—Maternal antibodies may inhibit priming of protective responses by IN delivered BRSV vaccines.

Passive immunization by maternal antibodies is necessary for the survival of young vertebrates as a means to guard against pathogenic microbes, which are everywhere.1 Documenting this is the poor survivability of calves that have less than optimal, or complete, failure of passive transfer, many of which will succumb to diarrhea, respiratory tract disease, or septicemia early in postnatal life because of uncontrolled growth of pathogens.2 Maternal antibodies can be viewed as a double-edged sword, however, because most data indicate that they inhibit priming of protective responses following vaccination of young animals.1 These data are primarily derived from studies of parenteral vaccination. There is a dearth of similar studies that examine mucosally administered vaccines, although there is currently renewed interest in mucosal delivery of antigens as a means to override maternal antibodies and achieve successful priming of the acquired immune response in young seropositive animals.3

Bovine respiratory syncytial virus is a pneumovirus in the family Paramyxoviridae that infects cattle populations endemically worldwide4 and primarily clinically affects the young in recurrent seasonal outbreaks.4 Clinical disease in calves often occurs when passive immunity has waned and is characterized by pyrexia, coughing, and tachypnea, often progressing rapidly to dyspnea.4

The efficacy of parenteral administration of several commercially available modified-live and inactivated BRSV vaccines has been demonstrated in a BRSV-challenge model that produces typical signs and lesions of acute BRSV5–7; however, epidemiological features4 of BRSV infections in cattle indicate the need for immunoprophylactic interventions early in life when maternal immunity may affect successful parenteral immunization. To address this issue, a recent study8 with the aforementioned BRSV-challenge model has demonstrated the efficacy of IN administration of a combination viral vaccine in protecting young seronegative calves from virulent experimentally induced BRSV infection. Because most calves in the field will be variably seropositive for BRSV,4 the objective of the study reported here was to determine whether a combination modified-live BRSV vaccine could stimulate protective immunity in young BRSV-seropositive calves following IN administration.

Materials and Methods

Calves—Neonatal Holstein calves were obtained from local dairies. They were removed from their dams at birth and fed 1.5 L of either pooled frozen bovine colostrum that was previously screened by an ELISA for minimal antibodies to BRSV or reconstituted pooled spray-dried colostrum that had highly positive ELISA results for antibodies to BRSV.a Ingestion of this colostrum ensured that the calves had known antibody concentrations for BRSV at the time of vaccination and passive immunity to other commonly encountered bovine pathogens. Calves were then maintained on 2 L of milk replacer fed twice daily and, depending on their age, with ad libitum water, grass-legume hay, and commercial pelleted calf ration. They were reared outdoors in individual calf hutches and maintained throughout the studies according to the established guidelines of the Canadian Council on Animal Care.

Vaccines—A combination vaccineb containing modified-live BRSV, parainfluenza virus 3, bovine herpesvirus 1, types 1 and 2 bovine viral diarrhea virus, and avirulent live cultures of Pasteurella Multocida and Mannheimia hemolytica was obtained from the pharmacy at the Western College of Veterinary Medicine. The same vaccine with an approximate 1/100 dose (ie, the MID) of BRSV, compared with that found in the vaccine for commercial sale, was obtained directly from the manufacturer.b This so-called MID is used in federally regulated licensing trials and was therefore considered most relevant to examine a potential dose effect, as has been previously examined with other vaccines.6

BRSV challenge inoculum—The challenge inoculum consisted of lung wash obtained from a newborn calf infected with BRSV as previously described.9 The lung wash was confirmed negative for bacterial contamination, Mycoplasma sp, bovine herpes virus 1, parainfluenza virus 3, and bovine viral diarrhea viruses by use of standard diagnostic methods.

Clinical assessment—Calves were observed for clinical signs on days −1 and 0 prior to challenge and on days 1 through 8 after challenge with the BRSV inoculum. Clinical assessments were made at the same time each morning by a veterinarian unaware of treatment status of the calves. Clinical signs, including signs of depression, respiratory rate, dyspnea, and cough, were recorded and scored (Appendix). Rectal temperatures were also recorded. Calves were euthanized by barbituratec overdose 8 days after challenge if not found dead or euthanized earlier for humane reasons. Calves were euthanized prior to day 8 after challenge exposure if 2 clinical signs indicative of substantial respiratory tract disease, including moderate signs of depression, dull eyes, droopy ears, rough coat, gauntness, and moderate respiratory distress or dyspnea (> 80 breaths/min), were observed for 2 consecutive days. If calves were observed at any time with signs of severe respiratory distress, including pronounced open-mouthed, labored breathing (> 100 breaths/min), or were severely depressed and recumbent with total reluctance to rise or with a Pao2 < 45 mm Hg, they were euthanized immediately. These criteria were consistent with the Canadian Council of Animal Care guidelines and were approved by the Committee on Animal Care and Supply at the University of Saskatchewan.

Sample collection—Deep nasal swab specimens were taken from both nares prior to challenge and on days 2 through 8 after challenge for virus isolation.9 Swab specimens were placed in 1 mL of transport medium consisting of Dulbecco modified Eagle medium supplemented with 10% fetal bovine serum,d 0.5M MgSO4, 50mM HEPES, sodium penicillin G (200 U/mL), streptomycin sulfate (200 μg/mL), and amphotericin B (1 μg/mL) and frozen at −70°C. Serum samples were collected from blood obtained by jugular venipuncture. Arterial blood samples were collected from the caudal thoracic aorta,10 and Pao2 (mm Hg) measurements, corrected for rectal temperature, were performed by use of a gas analyzer.e

Antibody assays—The BRSV-specific IgG ELISAs for serum and colostral antibodies were performed as previously described.11 Briefly, the 96-well polystyrene microtiter platesf were coated overnight at 4°C with BRSV antigen11 diluted in carbonate buffer (pH, 9.6) in alternating rows with cell control (uninfected cell lysate), washed, and blocked with 0.2% gelating in carbonate buffer. Samples were added and incubated at 37°C for 1 hour. The plates were washed, followed by addition of a 1:5,000 dilution of horseradish peroxidase–conjugated protein G.h Single component 2,2'-azino-di(3-ethyl-benzthiazoline-6-sulfonate) was used as the enzyme substrate.i Sample and antibody dilutions were made in ELISA working buffer (0.01M phosphate buffer [pH, 7.2] with 0.75M NaCl and 0.05% Tween 20h) with 0.2% gelatin. Serum samples were diluted 1:50. Convalescent serum from an unvaccinated calf with naturally occurring BRSV infection was used as a positive standard. High OD values, relative to a fetal bovine serum negative control, were obtained with this serum. Optical density values that were measured by use of a microplate readerj were converted to ELISA units with a software program.k The net OD value of the sample serum was calculated by subtracting the mean sample OD values of the cell control–coated wells from the mean sample OD values of the BRSV-coated wells. Similarly, the net OD values were calculated for the positive and negative control sera. Final serum results were expressed in ELISA units as follows:

article image

where neg equals negative and pos equals positive. A BRSV-specific IgA ELISA on nasal secretions was performed as previously described.9 Briefly, 96-well plates were coated overnight at 4°C with BRSV antigen and cell control antigen in carbonate buffer (pH, 9.6), washed, and blocked with 0.2% gelatin in carbonate buffer for 30 minutes at 37°C. Fluid from nasal swab specimens was diluted 1:2 in ELISA working buffer containing 0.2% gelatin and added to the 96-well plate in duplicate wells containing BRSV antigen and cell control antigen. Plates were washed and samples were added and allowed to incubate for 1 hour at 37°C. After washing the plates, the conjugate was added and allowed to incubate for 1 hour at 37°C. Bound IgA antibody was detected with horseradish peroxidase–conjugated rabbit anti-bovine IgAl at 1:4,000. The assay colorimetric conversion was detected with 2,2'-azino-di(3-ethyl-benzthiazoline-6-sulfonate) and stopped with 1% SDS. A pool of nasal secretions from 5 naturally exposed cattle was used as the standard for these assays and was tested at a 1:2 dilution. All steps in the ELISAs contained 100-μL volumes. After each step, plates were washed 4 times in 0.01M PBS solution (pH, 7.2) containing 0.05% Tween 20. Positive control samples comprised pooled supernatants from nasal swab specimens from vaccinated cattle obtained after BRSV-challenge exposure, and negative control samples comprised pooled supernatants from nasal swab specimens from clinically normal cattle seronegative for BRSV. The ELISA OD units were calculated as already described by use of the relevant control samples.

Quantitative virus isolation—Virus shedding was quantitatively determined by use of a microisolation plaque assay with bovine embryonic lung fibroblasts.12 This assay has a maximum calculated sensitivity of 10 pfu/mL, as previously described.12

Postmortem analysis—On necropsy, the respiratory tract of each calf was harvested and analyzed for the percentage of pneumonic tissue as previously described,9 with minor modifications. Dorsal and ventral surfaces of the lungs were photographed with a digital camera. Tracings were made on acetate from color prints of the digital images of the dorsal and ventral surfaces, outlining the total lung area and coloring in the dark-red, obviously pneumonic areas as well as emphysematous areas. The percentage area of the lungs affected for each tracing was determined by use of a software program.m

Experimental design—Two separate experiments were conducted: one experiment with the commercially available combination BRSV vaccine,b and a second experiment with an MID of the BRSV component in the otherwise same vaccineb given IN or SC to address the possible effects of dose and route of administration on the response.

Experiment 1

Eleven 3- to 8-day-old BRSV-seropositive (ELISA units range, 47 to 74) calves (group A) and ten 3- to 8-day-old BRSV-seronegative (ELISA units range, 2 to 15) calves (group B) were vaccinated IN with 1 mL (0.5 mL/nostril) of the combination vaccineb containing modified-live BRSV. Eleven 3- to 8-day-old BRSV-seropositive (ELISA units range, 44 to 70) control calves (group C) were administered 1 mL of the vaccine diluent (distilled water) IN. To prevent cross-exposure to shedding of vaccine viruses, each group of calves was maintained in separated individual calf hutches in separate (approx 60 × 60-feet) outdoor pens prior to vaccination and for approximately 60 days after vaccination, when they were commingled in 1 large (approx 60 × 120-feet) outdoor pen with a 3-sided shelter. On the day of challenge, approximately 4.5 months after vaccination, calves were transported 60 miles to the Western College of Veterinary Medicine and were challenged by aerosol delivery of BRSV into the enclosed, approximately 24 × 8 × 8-feet transport (stock) trailer (1,024 cubic feet of air space). For aerosol delivery, 30 mL of in vivo–passaged BRSV inoculum (103.4 pfu/mL) was placed in each of 3 ultrasonic nebulizers,n which were placed equidistant approximately 6 feet off the floor of the trailer. After approximately 40 minutes in the sealed trailer, calves were removed from the trailer and maintained as a single group in 1 large covered pen with access to a large uncovered open-air corral. Calves were monitored for clinical signs for 8 days after challenge, after which they were euthanized and necropsied, unless euthanized earlier because of signs of severe respiratory tract disease.

Serum samples were obtained prior to vaccination, prior to commingling, prior to challenge, on day 8 after challenge, or at the time of euthanasia and analyzed for BRSV-specific IgG by use of an ELISA. Arterial blood samples for analysis of Pao2 were collected on day 6 after challenge. Nasal swab specimens were collected prior to vaccination, prior to challenge, and on days 2 to 8 after challenge for analysis of BRSV shedding.

Experiment 2

Eleven 3- to 8-day-old BRSV-seronegative calves (group D) were vaccinated IN with 1 mL (0.5 mL/nostril; intended recommended dose for this vaccine) of the combination vaccine containing an MID of modified-live BRSVb; twelve 3- to 8-day-old BRSV-seronegative calves (group E) were vaccinated SC with 2 mL (label recommended dose) of the combination vaccine containing an MID of modified-live BRSV,b and eleven 3- to 8-day-old BRSV-seronegative control calves (group F) were administered 1 mL of the vaccine diluent (distilled water) IN. To prevent cross-exposure to shedding of vaccine viruses, each group of calves was maintained in separated individual calf hutches in separate (approx 60 × 60 feet) outdoor pens prior to vaccination and for 21 days, after which they were commingled on the day of challenge and transported 60 miles to the Western College of Veterinary Medicine. They were challenged as in experiment 1 with the same amount and lot of in vivo–passaged BRSV and then removed from the trailer and maintained as a single group in 1 large covered pen with access to a large uncovered open-air corral (approx 60 × 60 feet). Calves were monitored for clinical signs for 8 days after challenge, after which they were euthanized and necropsied, unless euthanized earlier because of severe respiratory tract disease.

Serum samples were obtained prior to vaccination, prior to challenge, and on day 8 after challenge or at the time of euthanasia and analyzed for BRSV-specific IgG by ELISA. Arterial blood samples for analysis of Pao2 were collected on day 6 after challenge. Nasal swab specimens were collected prior to vaccination, prior to challenge, and on days 2 to 8 after challenge for analysis of BRSV shedding. Concentrations of BRSV-specific IgA were assayed in the same nasal samples collected on the day of challenge and on the day of euthanasia.

Data analysis—Repeated observations were summarized to reflect clinically important outcomes. These calculated variables were then used for data analysis. Outcomes that were determined for each calf included the maximum rectal temperature, maximum change in rectal temperature, and number of days the rectal temperature was > 39.6°C (103.3°F). Maximum change in rectal temperatures after challenge was also determined for each calf by subtracting the values after challenge, days 1 to 8, from the baseline. Baseline was calculated for each calf by use of the mean value for 2 days prior to challenge. The maximum concentration of BRSV particles shed per individual nasal swab specimen prior to euthanasia and the number of days that virus particles were shed were also calculated for each calf. Maximum mean clinical score and the number of days a calf had a mean clinical score of ≥ 1 were also determined. Arterial blood oxygenation and percentage of the lungs affected with pneumonic lesions were also assessed. Descriptive statistics were performed, and data were assessed for normality. Appropriate nonparametric tests were used to compare the differences between treatment groups for the summarized outcomes of interest.8 All of the variables were compared between the treatment and control groups by use of the Kruskall-Wallis test, and all differences with a value of P < 0.05 were considered significant. If the P value was < 0.05 on the Kruskall-Wallis test, post hoc pairwise comparisons were completed by use of the Mann-Whitney U test to identify where the differences existed between the groups. For experiment 2, impact of the change in antibody concentrations after vaccination and after challenge on oxygen saturation and in the percentage of lungs affected with pneumonic lesions was assessed by use of the Spearman rank correlation (ρ). All analyses were performed by use of a commercially available software program.o

Results

Experiment 1—Four calves (1 from group A, 2 from group B, and 1 from group C) died or were euthanized as a result of severe diarrhea that occurred at least 12 days after vaccination and before commingling of groups. No clinical signs of respiratory tract disease or pulmonary lesions were observed in these calves. After challenge at approximately 4.5 months of age, calves in all groups developed signs of respiratory tract disease, including pyrexia (Figure 1), that were characteristic of BRSV infection. There were no significant differences in the maximum rectal temperature reached (P = 0.08) and the number of days a calf had a rectal temperature > 39.6°C (103.3°F; P = 0.5) among groups. There was a difference among groups in the maximum change in rectal temperature from baseline (defined as the mean rectal temperature recording for 2 days immediately prior to challenge). Seropositive IN vaccinated calves in group A had a significantly (P = 0.002) greater maximum change in rectal temperature (median, 2.7°C [4.9°F]; range, 2.1° to 3.5°C [3.7° to 6.3°F]) from baseline readings, compared with seronegative IN vaccinated calves in group B (median, 1.9°C [3.4°F]; range, 1.4° to 2.5°C [2.5° to 4.5°F]) or seropositive control calves in group C (P = 0.03; median, 2.2°C [3.9°F]; range, 2.9° to 3.2°C [3.3° to 5.8°F]). No significant (P = 0.08) differences were detected between seronegative IN vaccinated calves (group B) and seropositive control calves (group C). No significant difference was detected among groups in maximum mean clinical score (P = 0.8) or the number of days a calf had a mean clinical score ≥ 1 (P = 0.4).

Figure 1—
Figure 1—

Experiment 1, median rectal temperatures before and after BRSV challenge in approximately 4.5-month-old calves that had been previously vaccinated IN at 3 to 8 days of age with a combination vaccineb containing modified-live BRSV (group A, 10 BRSV-seropositive calves; group B, 8 BRSV-seronegative calves) or with vaccine diluent (group C, 10 BRSV-seropositive calves). Error bars represent minimum and maximum rectal temperatures recorded for each day.

Citation: Journal of the American Veterinary Medical Association 236, 9; 10.2460/javma.236.9.991

On day 6 after challenge, 1 calf in group B was found dead and 2 calves from each group required euthanasia because of signs of severe respiratory tract disease. On day 7 after challenge, 1 calf in group C was found dead and 3 calves from group A, 2 calves from group B, and 2 calves from group C required euthanasia because of signs of severe respiratory tract disease. There was no significant (P > 0.7) difference in mortality rate (as defined by death or required euthanasia) prior to termination of the study on day 8 after challenge among the treatment groups.

Nasal shedding of BRSV

No shedding of BRSV was detected in any calf prior to challenge in any of the groups (Figure 2). Calves in group B (seronegative and vaccinated) shed virus for significantly fewer days (median, 2 days; range, 0 to 4 days) than calves in groups A (P = 0.05; median, 3 days; range, 0 to 5 days) and C (P = 0.02; median, 4 days; range, 0 to 5 days). There were no significant (P = 0.5) differences in the maximum amount of BRSV shedding among groups.

Figure 2—
Figure 2—

Experiment 1, nasal shedding of BRSV after BRSV challenge in approximately 4.5-month-old calves that had been previously vaccinated IN at 3 to 8 days of age with a combination vaccineb containing modified-live BRSV (group A, 10 BRSV-seropositive calves; group B, 8 BRSV-seronegative calves) or with vaccine diluent (group C, 10 BRSV-seropositive calves). Bars indicate the percentage of calves shedding detectable BRSV on each day. Absence of a bar indicates that no calves shed virus that day.

Citation: Journal of the American Veterinary Medical Association 236, 9; 10.2460/javma.236.9.991

BRSV-specific antibody responses

At the time of challenge, all calves in all groups had minimal BRSV-specific IgG (< 14 ELISA units). After challenge, only 4 calves in group B had anamnestic (> 3 times increase in ELISA units) BRSV IgG responses at the time of death or euthanasia; none of the other calves in any group had increased IgG responses after challenge.

Pneumonic lesions and Pao2

Calves in all groups had pneumonic lesions typical of acute BRSV infection (Figure 3). Although calves in group A had more extensive lesions (median, 47%; range, 27% to 57% pneumonic lung) than calves in group B (median, 30%; range, 5% to 53% pneumonic lung) or C (median, 38%; range, 3% to 50% pneumonic lung), there were no significant differences among groups. There were no significant (P = 0.08) differences in the aortic blood oxygen concentrations in surviving calves on day 7 among the 3 treatment groups.

Figure 3—
Figure 3—

Experiment 1, scatterplot of the percentage of lungs affected with pneumonic lesions after BRSV challenge in approximately 4.5-month-old calves that had been previously vaccinated IN at 3 to 8 days of age with a combination vaccineb containing modified-live BRSV (group A, 10 BRSV-seropositive calves; group B, 8 BRSV-seronegative calves) or with vaccine diluent (group C, 10 BRSV-seropositive calves).

Citation: Journal of the American Veterinary Medical Association 236, 9; 10.2460/javma.236.9.991

Experiment 2—Three calves (1 from group E and 2 from group F) died or were euthanized because of severe diarrhea (coronavirus [n = 2 calves] and coccidia [1]) between the time of vaccination and challenge. No clinical signs of respiratory tract disease or pulmonary lesions were observed in these calves. Retrospective analyses of serum IgG titers indicated that 6 calves were considered seropositive (ELISA units range, 25 to 86) prior to vaccination as a result of apparent unnoticed ingestion of maternal colostrum. These calves were removed from further groupwise analyses. After challenge at approximately 21 days after vaccination, calves in all groups developed signs of respiratory tract disease, including pyrexia (Figure 4), that were characteristic of BRSV infection. There were no significant differences among groups in maximum rectal temperatures (P = 0.1) and the number of days the rectal temperature was > 39.6°C (103.3°F [P = 0.1]). There was a significant difference in the maximum change in rectal temperature from baseline between the groups. Calves in group F (seronegative and control) had significantly greater changes in rectal temperature (median, 2.5°C [4.5°F]; range, 1.7° to 3.0°C [3.1° to 5.5°F]) from baseline, compared with group D (seronegative and IN vaccinated; P = 0.03; median, 1.9°C [3.4°F]; range, 0.9° to 2.4°C [1.7° to 4.4°F]) and group E (seronegative and SC vaccinated; P = 0.04; median, 2.0°C [3.7°F]; range, 1.6° to 2.4°C [2.9° to 4.4°F]). No significant (P = 0.3) differences were detected in maximum rectal temperature change from baseline between groups D and E. There were no significant differences in the maximum mean clinical score (P = 0.3) or the number of days a calf had a clinical score of ≥ 1 (P = 0.4) among groups. On day 5 after challenge, 1 calf in group E required euthanasia because of signs of severe respiratory tract disease. On day 6 after challenge, 1 calf in group D was found dead. On day 7 after challenge, 1 calf in group E and 2 calves from group F required euthanasia because of signs of severe respiratory tract disease. There was no significant (P > 0.5) difference in mortality rate (as defined by death or required euthanasia) prior to termination of the study on day 8 after challenge among the treatment groups.

Figure 4—
Figure 4—

Experiment 2, median rectal temperatures before and after BRSV challenge in calves that had been vaccinated 21 days ago (at 3 to 8 days of age) with a combination vaccineb containing a minimum immunizing (1/100) dose of modified-live BRSV (group D: IN vaccination, 8 BRSV-seronegative calves; group E: SC vaccination, 9 BRSV-seronegative calves) or with vaccine diluent (group F: IN vaccination, 7 BRSV-seronegative calves). See Figure 1 for remainder of key.

Citation: Journal of the American Veterinary Medical Association 236, 9; 10.2460/javma.236.9.991

Nasal shedding of viruses

No shedding of BRSV was detected in any calf prior to challenge in any of the groups (Figure 5). No differences were found between groups for the number of days BRSV was shed (P = 0.07). Vaccinated calves in groups D (IN; P = 0.02; median, 440 pfu/mL; range, 60 to 2,040 pfu/mL) and E (SC; P = 0.02; median, 540 pfu/mL; range, 80 to 1,000 pfu/mL) shed significantly less virus than calves in the control group F (median, 1,680 pfu/mL; range, 550 to 2,480 pfu/mL).

Figure 5—
Figure 5—

Experiment 2, nasal shedding of BRSV after BRSV challenge in calves that had been vaccinated 21 days ago (at 3 to 8 days of age) with a combination vaccineb containing a minimum immunizing (1/100) dose of modified-live BRSV (group D: IN vaccination, 8 BRSV-seronegative calves; group E: SC vaccination, 9 BRSV-seronegative calves) or with vaccine diluent (group F: IN vaccination, 7 BRSV-seronegative calves). See Figure 2 for remainder of key.

Citation: Journal of the American Veterinary Medical Association 236, 9; 10.2460/javma.236.9.991

Pneumonic lesions and Pao2

Calves in all groups had pneumonic lesions typical of acute BRSV infection (Figure 6). Unvaccinated calves in group F had significantly more severe lung lesions (median, 38%; range, 23% to 48% pneumonic lung) than in IN vaccinated calves in group D (P = 0.05; median, 20%; range, 15% to 61% pneumonic lung) and SC vaccinated calves in group E (P = 0.02; median, 17%; range, 10% to 54% pneumonic lung). Similarly, although the unvaccinated calves in group F had lower Pao2 measurements (median, 59 mm Hg; range, 51 to 100 mm Hg) than IN vaccinated calves in group D (median, 80 mm Hg; range, 47 to 94 mm Hg) and SC vaccinated calves in group E (median, 86 mm Hg; range, 53 to 98 mm Hg), these differences were not significant (P = 0.1). Four of the 6 calves that were inadvertently seropositive at the time of vaccination still had what would be considered potentially protective concentrations of BRSV-specific IgG at the time of challenge (ELISA units range, 29 to 74). All 6 of the calves had minimal or less severe lung lesions (pneumonic lung range, 6.5% to 14%) than all of the seronegative calves.

Figure 6—
Figure 6—

Experiment 2, scatterplot of the percentage of lungs affected with pneumonic lesions after BRSV challenge in calves that had been vaccinated 21 days ago (at 3 to 8 days of age) with a combination vaccineb containing a minimum immunizing (1/100) dose of modified-live BRSV (group D: IN vaccination, 8 BRSV-seronegative calves; group E: SC vaccination, 9 BRSV-seronegative calves) or with vaccine diluent (group F: IN vaccination, 7 BRSV-seronegative calves).

Citation: Journal of the American Veterinary Medical Association 236, 9; 10.2460/javma.236.9.991

BRSV-specific antibody responses

At the time of challenge, all calves (that were not excluded) in all groups had minimal BRSV-specific IgG (< 14 ELISA units). After challenge, only 1 calf in group D had an anamnestic (> 3X increase in ELISA units) BRSV IgG response at the time of euthanasia on day 8 after challenge; none of the other calves in any group seroconverted.

Evaluation of BRSV-specific IgA antibodies in nasal secretions from the calves indicated no significant (P = 0.3) differences among groups on the day of challenge (group D: median, 12 ELISA units; range, 0 to 41 ELISA units; group E: median, 1 ELISA unit; range, 0 to 10 ELISA units; and group F: median, 5 ELISA units; range, 1 to 12 ELISA units). After challenge, nasal IgA concentrations were not significantly (P = 0.9) different among the groups (group D: median, 11 ELISA units; range, 8 to 219 ELISA units; group E: median, 13 ELISA units; range, 4 to 101 ELISA units; and group F: median, 11 ELISA units; range, 9 to 18 ELISA units).

The range of ELISA units in groups D and E could be misleading. One calf in group D had much higher IgA concentrations than the other calves in the group both on the day of challenge (41 ELISA units) and on day 8 after challenge (219 ELISA units); on the day of challenge, the next highest concentration detected in this group was 13 ELISA units. On day 8 after challenge, the next highest concentration detected in the group was 24 ELISA units. Similarly, 1 calf in group E had IgA concentrations of 101 ELISA units on day 8 after challenge, whereas the next highest ELISA unit in that group on that day was 44 ELISA units.

Correlation analyses

Nasal IgA production in response to vaccination did not correlate with either percentage of lung lesions (P = 0.2) or the blood oxygen saturation (P = 0.8).

Discussion

Studies5–7 of vaccine efficacy for licensing and other regulatory purposes are generally conducted in seronegative animals that are challenged shortly after vaccination, usually < 1 month. Although such studies5–8 can demonstrate the efficacy of vaccine-stimulated, disease-sparing immune responses, as has been previously reported for parenteral and IN vaccination for BRSV by use of the challenge BRSV-challenge model described herein, they do not address other cofactors such as passive immunity and longevity of response that can affect overall vaccine efficacy. The purpose of this investigation was to address these cofactors. The poor efficacy reported here, compared with previous studies5–8 on this BRSV-challenge model, following IN vaccination of seropositive and seronegative calves and BRSV challenge approximately 140 days later could be the result of several factors.

It is generally accepted that passive immunity from maternal antibodies not only protects young animals from infectious disease but can also inhibit the response to vaccination early in life.1 Epidemiological data and experimentally induced infection studies have shown that although maternal antibodies do not prevent BRSV infection, they do reduce signs of clinical disease subsequent to infection.4 Reduced pneumonic lesions in the seropositive calves in experiment 2 that were excluded from the analysis of the sero-negative calves further confirm this disease-sparing effect in our BRSV-challenge model of severe acute BRSV infection. This disease-sparing effect could be the result of the transfer and resecretion of maternal IgG1 to the respiratory mucosae, as has been demonstrated in the gastrointestinal tract13 or the exudation of antibodies from the microvasculature in response to the inflammation4 that occurs subsequent to BRSV infection. Relatedly, maternal IgG1 resecreted onto the mucosae of the nose and pharynx could affect the inhibitory signaling related to priming of B-cell responses14 or neutralize vaccine virus, thereby affecting the apparent lack of efficacy observed in the calves that were seropositive at the time of IN vaccination.

There are conflicting data concerning the inhibitory activity of colostral antibodies on the priming of BRSV-specific responses. Consistent with the lack of disease-sparing effect of vaccination in the seropositive vaccinated calves in experiment 1 of this study, others have reported the inhibitory effect of maternal antibodies on vaccine-induced antibody responses to BRSV following parenteral vaccination.15–17 In contrast, the priming of BRSV-specific antibody18 and cellular immune19 responses following parenteral vaccination of seropositive calves has been reported. As well, more recent challenge studies20–22 report reduction of BRSV shedding following challenge of sero-positive calves that were vaccinated parenterally or IN; however, in those studies, no or mild clinical disease was produced following the experimentally induced infections and pulmonary pathological findings were not reported, making it difficult to assess the disease-sparing effect of those protocols. These apparent inconsistencies in the ability of vaccines to override maternal antibodies may be the result of the probability that overriding of maternal antibodies by vaccines is not an all or none phenomenon, and is likely affected by the concentration of maternal antibodies at the time of vaccination as well as vaccine formulation, including antigen mass, degree of attenuation of vaccinal BRSV, and effects of adjuvants.

Reinfection and some degree of clinical disease are well-documented features of the respiratory syncytial virus infections in both cattle and humans,4 suggesting that the duration of clinical immunity can be short. Consistent with these epidemiological data was the poor efficacy of IN vaccination in protecting the seronegative calves when they were challenged > 4 months after a single vaccination. Following challenge, most of these calves developed moderate to severe clinical disease and pulmonary lesions, compared with those calves that were vaccinated and challenged within 30 days in this and previously reported studies.5–8 This apparent short duration of immunity for BRSV contrasts with what has been reported following parenteral vaccination of seronegative calves for bovine viral diarrhea virus and bovine herpes virus-1.23,24 In contrast to BRSV that replicates only locally in the epithelia of the respiratory tract,25 both vaccinal and field isolates of bovine viral diarrhea virus and bovine herpes virus-1 replicate systemically,26,27 which may stimulate different immune effector mechanisms or provide for more long-term systemic memory responses. The apparent short duration of immunity for BRSV4 is consistent with the idea that it is difficult to stimulate long-term memory responses on mucosal surfaces28 and may be a caveat to assuming that immune responses to all agents in commonly used combination vaccines have a similar duration of immunity.

It is possible that the dose or degree of attenuation of BRSV or the vaccine formulation, which contained live bacterial cultures, contributed to the poor efficacy observed in experiment 1. Arguing against these possibilities are the data from experiment 2, which constituted a more conventional efficacy study for vaccine licensing and demonstrated disease-sparing effects (decreased rectal temperature change, decreased shed of BRSV, and decreased severity of pulmonary lesions) in seronegative calves that were vaccinated with an MID of BRSV and challenged 3 weeks later. Although this disease-sparing effect was not as great as has been observed following IN administration of another combination BRSV vaccine at commercial release dose8 or an MID by one of our authors (JAE), it was associated with an anamnestic mucosal IgA response in several of the calves and a minimal systemic IgG response in most of the calves after challenge, as in previous studies.5,17 Interestingly, similar to previous studies,17 parenteral administration of this vaccine also apparently primed for an anamnestic IgA response subsequent to challenge in several calves.

These results raise additional questions about the management of vaccines for BRSV in young calves, but could, to some extent, be an artifact of the experimental conditions. To determine the potential inhibitory effect of maternal antibodies, young calves need to be vaccinated neonatally and held in isolation (to prevent natural exposure to BRSV) until maternal antibodies decay. This period (≥ 3 months) that is required for decay of colostral antibodies to nonprotective concentrations may be approximately the extent of duration of immunity for respiratory syncytial virus infections,4 making it difficult, if not impossible, to determine experimentally which factor, inhibition by maternal antibodies or short duration of immunity, may have been more contributory to the observed poor efficacy of this vaccine following IN administration to young calves. Nevertheless, given the documented complete or partial failure of passive transfer in many calf populations,2 and resultant variability in the window of susceptibility to clinical BRSV infection, neonatal vaccination is still a viable tool to reduce calfhood pneumonia or effectively prime young animals for protective acquired immunity. As well, given the endemicity of BRSV,4 it is likely that in many, if not most, calf-rearing situations, calves that are iatrogenically primed by vaccination neonatally would be naturally and repeatedly boosted by circulating BRSV, thereby developing effective acquired immunity. This latter likelihood is another phenomenon that is difficult to model experimentally but is probably important in developing and maintaining clinical immunity to BRSV and other respiratory pathogens in the context of variable passive transfer. Further studies are required to determine the timing of primary and secondary immunizations and also to determine which routes and times of vaccine administration will provide the optimal combination to engender protective acquired clinical immunity against BRSV in young calves.

ABBREVIATIONS

BRSV

Bovine respiratory syncytial virus

IN

Intranasally

MID

Minimum immunizing dose

OD

Optical density

pfu

Plaque forming unit

a.

Headstart Gold, Saskatoon Colostrum Co, Saskatoon, SK, Canada.

b.

Vista Once SQ, Intervet Canada, Whitby, ON, Canada.

c.

Euthanyl Forte, MTC Pharmaceuticals, Cambridge, ON, Canada.

d.

Invitrogen, Burlington, ON, Canada.

e.

Model 288, Ciba-Corning, Medfield, Mass.

f.

Immulon 4HBX, Thermo Electron Corp, Milford, Mass.

g.

Sigma Chemical Co, St Louis, Mo.

h.

Zymed, San Francisco, Calif.

i.

Kirkegaard & Perry Laboratories Inc, Gaithersburg, Md.

j.

Benchmark microplate reader, Bio-Rad Laboratories Inc, Mississauga, ON, Canada.

k.

Microplate Manager, version 5.0.1, Bio-Rad Laboratories Inc, Mississauga, ON, Canada.

l.

Bethyl Laboratories Inc, Montgomery, Tex.

m.

Image 1, Universal Imaging Corp, West Chester, Pa.

n.

Ultra-Neb 99, Devilbiss, Somerset, Pa.

o.

SPPS 17 for Windows, Analytical Software, Chicago, Ill.

References

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    Tizard I. Immunity fetus and newborn. In: Veterinary immunology, an introduction. Philadelphia: WB Saunders Co, 2009;228236.

  • 2.

    National Animal Health Monitoring System. Dairy 1996: national dairy health evaluation project. Dairy heifer morbidity, mortality, and health management focusing on preweaned heifers. Fort Collins, Colo: USDA, APHIS Veterinary Services, 1996.

    • Search Google Scholar
    • Export Citation
  • 3.

    Griebel PJ. Mucosal vaccination of the newborn: an unrealized opportunity. Expert Rev Vaccines 2009;8:13.

  • 4.

    Baker JC, Ellis JA, Clark EG. Bovine respiratory syncytial virus. Vet Clin North Am Food Anim Pract 1997;13:425454.

  • 5.

    West K, Petrie L & Konoby C, et al. The efficacy of modified-live bovine respiratory syncytial virus vaccines in experimentally infected calves. Vaccine 1999;18:907919.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 6.

    Ellis J, West K & Konoby C, et al. Efficacy of an inactivated respiratory syncytial virus vaccine in calves. J Am Vet Med Assoc 2001;218:19731980.

  • 7.

    Ellis J, West KH & Waldner C, et al. Efficacy of a saponin-adjuvanted inactivated respiratory syncytial virus vaccine in calves. Can Vet J 2005;46:155162.

    • Search Google Scholar
    • Export Citation
  • 8.

    Ellis J, Gow S & West K, et al. Response of calves to challenge exposure with virulent bovine respiratory syncytial virus following intranasal administration of vaccines formulated for parenteral administration. J Am Vet Med Assoc 2007;230:233243.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 9.

    West K, Petrie L & Haines DM, et al. The effect of formalin-inactivated vaccine on respiratory disease associated with bovine respiratory syncytial virus infection in calves. Vaccine 1999;17:809820.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 10.

    Will JA, Bisgard GE. Cardiac catheterization of unanesthetized large domestic animals. J Appl Physiol 1972;33:400401.

  • 11.

    West K, Ellis J. Functional analysis of antibody responses of feedlot cattle to bovine respiratory syncytial virus following vaccination with mixed vaccines. Can J Vet Res 1997;61:2833.

    • Search Google Scholar
    • Export Citation
  • 12.

    West K, Bogdan J & Hamel A, et al. A comparison of diagnostic methods for the detection of bovine respiratory syncytial virus in experimental clinical specimens. Can J Vet Res 1998;62:245250.

    • Search Google Scholar
    • Export Citation
  • 13.

    Besser TE, McGuire TC & Gay CC, et al. Transfer of functional immunoglobulin G (IgG) antibody into the gastrointestinal tract accounts for IgG clearance in calves. J Virol 1988;62:22342237.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 14.

    Tizard IR. Regulation of acquired immunity. In: Veterinary immunology: an introduction. Philadelphia: WB Saunders Co, 2009;216.

  • 15.

    Fulton RW, Briggs RE & Payton ME, et al. Maternally derived humoral immunity to bovine viral diarrhea virus (BVDV) 1a, BVDV1b, BVDV2, bovine herpesvirus-1, parainfluenza-3 virus bovine respiratory syncytial virus, Mannheimia haemolytica and Pasteurella multocida in beef calves, antibody decline by half-life studies and effect on response to vaccination. Vaccine 2004;22:643649.

    • Search Google Scholar
    • Export Citation
  • 16.

    O'Neill RG, Woolliams JA & Glass EJ, et al. Quantitative evaluation of genetic and environmental parameters determining antibody response induced by vaccination against bovine respiratory syncytial virus. Vaccine 2006;24:40074016.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 17.

    O'Neill RG, Fitzpatrick JL & Glass EJ, et al. Optimisation of the response to respiratory virus vaccines in cattle. Vet Rec 2007;161:269270.

  • 18.

    Kimman TG, Westenbrink F, Straver PJ. Priming for local and systemic antibody memory responses to bovine respiratory syncytial virus: effect of amount of virus, virus replication, route of administration and maternal antibodies. Vet Immunol Immunopathol 1989;22:145160.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 19.

    Ellis JA, Hassard LE & Cortese VS, et al. The effect of perinatal vaccination on humoral and cellular immune responses in cows and young calves. J Am Vet Med Assoc 1996;208:393400.

    • Search Google Scholar
    • Export Citation
  • 20.

    Mawhinney IC, Burrows MR. Protection against bovine respiratory syncytial virus challenge following a single dose of vaccine in young calves with maternal antibody. Vet Rec 2005;156:139143.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 21.

    Harmeyer SS, Murray J & Imrie C, et al. Efficacy of a live bovine respiratory syncytial virus vaccine in seropositive calves. Vet Rec 2006;159:456457.

  • 22.

    Vangeel I, Antonis AF & Fluess M, et al. Efficacy of a modified live intranasal bovine respiratory syncytial virus vaccine in 3-week-old calves experimentally challenged with BRSV. Vet J 2007;174:627635.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 23.

    Ellis J, West K & Cortese V, et al. Effect of maternal antibodies on induction and persistence of vaccine-induced immune responses against bovine viral diarrhea virus type II in young calves. J Am Vet Med Assoc 2001;219:351356.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 24.

    Ellis JA, Waldner C & Rhodes C, et al. Longevity of protective immunity to experimental bovine herpesvirus-1 infection following inoculation with a combination modified-live virus vaccine in beef calves. J Am Vet Med Assoc 2005;227:123128.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 25.

    Viuff B, Tjørnehøj K & Larsen L, et al. Replication and clearance of respiratory syncytial virus: apoptosis is an important pathway of virus clearance after experimental infection with bovine respiratory syncytial virus. Am J Pathol 2002;161:21952207.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 26.

    Lemaire M, Meyer G & Baranowski E, et al. Production of bovine herpesvirus type 1-seronegative latent carriers by administration of a live-attenuated vaccine in passively immunized calves. J Clin Microbiol 2000;38:42334238.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 27.

    Kleiboeker SB, Lee SM & Jones CA, et al. Evaluation of shedding of bovine herpesvirus 1, bovine viral diarrhea virus 1, and bovine viral diarrhea virus 2 after vaccination of calves with a multivalent modified-live virus vaccine. J Am Vet Med Assoc 2003;222:13991403.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 28.

    Tizard I. Immunity at body surfaces. In: Veterinary immunology: an introduction. Philadelphia: WB Saunders Co, 2009;253254.

Appendix

Clinical scoring system used to evaluate calves.

VariableClinical scoring
Depression0 = Normal
1 = Mild; moves slowly, head down
2 = Moderate; tends to lie down, staggers
3 = Severe; recumbent or stands with difficulty
Respiratory rate0 = ≤ 44 breaths/min
1 = 45–64 breaths/min
2 = 65–80 breaths/min
3 = ≥ 81 breaths/min
Dyspnea0 = Normal
1 = Mild; short and rapid
2 = Moderate; labored, abdominal
3 = Severe; very labored, grunting
Cough*0 = < 3 episodes
1 = 3+ episodes

Cough scores were assigned to calves with spontaneous coughing during clinical examination period (approx 1 h/d).

  • Figure 1—

    Experiment 1, median rectal temperatures before and after BRSV challenge in approximately 4.5-month-old calves that had been previously vaccinated IN at 3 to 8 days of age with a combination vaccineb containing modified-live BRSV (group A, 10 BRSV-seropositive calves; group B, 8 BRSV-seronegative calves) or with vaccine diluent (group C, 10 BRSV-seropositive calves). Error bars represent minimum and maximum rectal temperatures recorded for each day.

  • Figure 2—

    Experiment 1, nasal shedding of BRSV after BRSV challenge in approximately 4.5-month-old calves that had been previously vaccinated IN at 3 to 8 days of age with a combination vaccineb containing modified-live BRSV (group A, 10 BRSV-seropositive calves; group B, 8 BRSV-seronegative calves) or with vaccine diluent (group C, 10 BRSV-seropositive calves). Bars indicate the percentage of calves shedding detectable BRSV on each day. Absence of a bar indicates that no calves shed virus that day.

  • Figure 3—

    Experiment 1, scatterplot of the percentage of lungs affected with pneumonic lesions after BRSV challenge in approximately 4.5-month-old calves that had been previously vaccinated IN at 3 to 8 days of age with a combination vaccineb containing modified-live BRSV (group A, 10 BRSV-seropositive calves; group B, 8 BRSV-seronegative calves) or with vaccine diluent (group C, 10 BRSV-seropositive calves).

  • Figure 4—

    Experiment 2, median rectal temperatures before and after BRSV challenge in calves that had been vaccinated 21 days ago (at 3 to 8 days of age) with a combination vaccineb containing a minimum immunizing (1/100) dose of modified-live BRSV (group D: IN vaccination, 8 BRSV-seronegative calves; group E: SC vaccination, 9 BRSV-seronegative calves) or with vaccine diluent (group F: IN vaccination, 7 BRSV-seronegative calves). See Figure 1 for remainder of key.

  • Figure 5—

    Experiment 2, nasal shedding of BRSV after BRSV challenge in calves that had been vaccinated 21 days ago (at 3 to 8 days of age) with a combination vaccineb containing a minimum immunizing (1/100) dose of modified-live BRSV (group D: IN vaccination, 8 BRSV-seronegative calves; group E: SC vaccination, 9 BRSV-seronegative calves) or with vaccine diluent (group F: IN vaccination, 7 BRSV-seronegative calves). See Figure 2 for remainder of key.

  • Figure 6—

    Experiment 2, scatterplot of the percentage of lungs affected with pneumonic lesions after BRSV challenge in calves that had been vaccinated 21 days ago (at 3 to 8 days of age) with a combination vaccineb containing a minimum immunizing (1/100) dose of modified-live BRSV (group D: IN vaccination, 8 BRSV-seronegative calves; group E: SC vaccination, 9 BRSV-seronegative calves) or with vaccine diluent (group F: IN vaccination, 7 BRSV-seronegative calves).

  • 1.

    Tizard I. Immunity fetus and newborn. In: Veterinary immunology, an introduction. Philadelphia: WB Saunders Co, 2009;228236.

  • 2.

    National Animal Health Monitoring System. Dairy 1996: national dairy health evaluation project. Dairy heifer morbidity, mortality, and health management focusing on preweaned heifers. Fort Collins, Colo: USDA, APHIS Veterinary Services, 1996.

    • Search Google Scholar
    • Export Citation
  • 3.

    Griebel PJ. Mucosal vaccination of the newborn: an unrealized opportunity. Expert Rev Vaccines 2009;8:13.

  • 4.

    Baker JC, Ellis JA, Clark EG. Bovine respiratory syncytial virus. Vet Clin North Am Food Anim Pract 1997;13:425454.

  • 5.

    West K, Petrie L & Konoby C, et al. The efficacy of modified-live bovine respiratory syncytial virus vaccines in experimentally infected calves. Vaccine 1999;18:907919.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 6.

    Ellis J, West K & Konoby C, et al. Efficacy of an inactivated respiratory syncytial virus vaccine in calves. J Am Vet Med Assoc 2001;218:19731980.

  • 7.

    Ellis J, West KH & Waldner C, et al. Efficacy of a saponin-adjuvanted inactivated respiratory syncytial virus vaccine in calves. Can Vet J 2005;46:155162.

    • Search Google Scholar
    • Export Citation
  • 8.

    Ellis J, Gow S & West K, et al. Response of calves to challenge exposure with virulent bovine respiratory syncytial virus following intranasal administration of vaccines formulated for parenteral administration. J Am Vet Med Assoc 2007;230:233243.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 9.

    West K, Petrie L & Haines DM, et al. The effect of formalin-inactivated vaccine on respiratory disease associated with bovine respiratory syncytial virus infection in calves. Vaccine 1999;17:809820.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 10.

    Will JA, Bisgard GE. Cardiac catheterization of unanesthetized large domestic animals. J Appl Physiol 1972;33:400401.

  • 11.

    West K, Ellis J. Functional analysis of antibody responses of feedlot cattle to bovine respiratory syncytial virus following vaccination with mixed vaccines. Can J Vet Res 1997;61:2833.

    • Search Google Scholar
    • Export Citation
  • 12.

    West K, Bogdan J & Hamel A, et al. A comparison of diagnostic methods for the detection of bovine respiratory syncytial virus in experimental clinical specimens. Can J Vet Res 1998;62:245250.

    • Search Google Scholar
    • Export Citation
  • 13.

    Besser TE, McGuire TC & Gay CC, et al. Transfer of functional immunoglobulin G (IgG) antibody into the gastrointestinal tract accounts for IgG clearance in calves. J Virol 1988;62:22342237.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 14.

    Tizard IR. Regulation of acquired immunity. In: Veterinary immunology: an introduction. Philadelphia: WB Saunders Co, 2009;216.

  • 15.

    Fulton RW, Briggs RE & Payton ME, et al. Maternally derived humoral immunity to bovine viral diarrhea virus (BVDV) 1a, BVDV1b, BVDV2, bovine herpesvirus-1, parainfluenza-3 virus bovine respiratory syncytial virus, Mannheimia haemolytica and Pasteurella multocida in beef calves, antibody decline by half-life studies and effect on response to vaccination. Vaccine 2004;22:643649.

    • Search Google Scholar
    • Export Citation
  • 16.

    O'Neill RG, Woolliams JA & Glass EJ, et al. Quantitative evaluation of genetic and environmental parameters determining antibody response induced by vaccination against bovine respiratory syncytial virus. Vaccine 2006;24:40074016.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 17.

    O'Neill RG, Fitzpatrick JL & Glass EJ, et al. Optimisation of the response to respiratory virus vaccines in cattle. Vet Rec 2007;161:269270.

  • 18.

    Kimman TG, Westenbrink F, Straver PJ. Priming for local and systemic antibody memory responses to bovine respiratory syncytial virus: effect of amount of virus, virus replication, route of administration and maternal antibodies. Vet Immunol Immunopathol 1989;22:145160.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 19.

    Ellis JA, Hassard LE & Cortese VS, et al. The effect of perinatal vaccination on humoral and cellular immune responses in cows and young calves. J Am Vet Med Assoc 1996;208:393400.

    • Search Google Scholar
    • Export Citation
  • 20.

    Mawhinney IC, Burrows MR. Protection against bovine respiratory syncytial virus challenge following a single dose of vaccine in young calves with maternal antibody. Vet Rec 2005;156:139143.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 21.

    Harmeyer SS, Murray J & Imrie C, et al. Efficacy of a live bovine respiratory syncytial virus vaccine in seropositive calves. Vet Rec 2006;159:456457.

  • 22.

    Vangeel I, Antonis AF & Fluess M, et al. Efficacy of a modified live intranasal bovine respiratory syncytial virus vaccine in 3-week-old calves experimentally challenged with BRSV. Vet J 2007;174:627635.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 23.

    Ellis J, West K & Cortese V, et al. Effect of maternal antibodies on induction and persistence of vaccine-induced immune responses against bovine viral diarrhea virus type II in young calves. J Am Vet Med Assoc 2001;219:351356.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 24.

    Ellis JA, Waldner C & Rhodes C, et al. Longevity of protective immunity to experimental bovine herpesvirus-1 infection following inoculation with a combination modified-live virus vaccine in beef calves. J Am Vet Med Assoc 2005;227:123128.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 25.

    Viuff B, Tjørnehøj K & Larsen L, et al. Replication and clearance of respiratory syncytial virus: apoptosis is an important pathway of virus clearance after experimental infection with bovine respiratory syncytial virus. Am J Pathol 2002;161:21952207.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 26.

    Lemaire M, Meyer G & Baranowski E, et al. Production of bovine herpesvirus type 1-seronegative latent carriers by administration of a live-attenuated vaccine in passively immunized calves. J Clin Microbiol 2000;38:42334238.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 27.

    Kleiboeker SB, Lee SM & Jones CA, et al. Evaluation of shedding of bovine herpesvirus 1, bovine viral diarrhea virus 1, and bovine viral diarrhea virus 2 after vaccination of calves with a multivalent modified-live virus vaccine. J Am Vet Med Assoc 2003;222:13991403.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 28.

    Tizard I. Immunity at body surfaces. In: Veterinary immunology: an introduction. Philadelphia: WB Saunders Co, 2009;253254.

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