Ivpa

Wa

b

a

ARRAA

KMBBIBPI

1

cbmp(sTtwmompit[

0d

Vaccine 28 (2010) 3784–3792

Contents lists available at ScienceDirect

Vaccine

journa l homepage: www.e lsev ier .com/ locate /vacc ine

mmunogenicity of a modified-live virus vaccine against bovine viral diarrheairus types 1 and 2, infectious bovine rhinotracheitis virus, bovinearainfluenza-3 virus, and bovine respiratory syncytial virus whendministered intranasally in young calves

enzhi Xuea,∗, John Ellisb, Debra Matticka, Linda Smitha, Ryan Bradya, Emilio Trigoa

Intervet/Schering-Plough Animal Health, 35500 W. 91st Street, DeSoto, KS 66018, USADepartment of Veterinary Microbiology, Western College of Veterinary Medicine, University, of Saskatchewan, Saskatoon, SK S7N 5B4, Canada

r t i c l e i n f o

rticle history:eceived 4 February 2010eceived in revised form 16 March 2010ccepted 20 March 2010vailable online 8 April 2010

a b s t r a c t

The immunogenicity of an intranasally-administered modified-live virus (MLV) vaccine in 3–8 day oldcalves was evaluated against bovine viral diarrhea virus (BVDV) types 1 and 2, infectious bovine rhino-tracheitis (IBR) virus, parainfluenza-3 (PI-3) virus and bovine respiratory syncytial virus (BRSV). Calveswere intranasally vaccinated with a single dose of a multivalent MLV vaccine and were challenged withone of the respective viruses three to four weeks post-vaccination in five separate studies. There was

eywords:odified-live virus vaccine

ovine respiratory diseasesVDV

BRRSV

significant sparing of diseases in calves intranasally vaccinated with the MLV vaccine, as indicated bysignificantly fewer clinical signs, lower rectal temperatures, reduced viral shedding, greater white bloodcell and platelet counts, and less severe pulmonary lesions than control animals. This was the first MLVcombination vaccine to demonstrate efficacy against BVDV types 1 and 2, IBR, PI-3 and BRSV in calves3–8 days of age.

© 2010 Elsevier Ltd. All rights reserved.

I-3ntranasal vaccination

. Introduction

The bovine respiratory disease (BRD) complex is a signifi-ant problem to the cattle industry, especially in calves enteringeef lots, often resulting in severe economic losses [1–4]. Theajor viral pathogens of the BRD complex include: bovine her-

esvirus type-1, also known as infectious bovine rhinotracheitisIBR) virus, bovine parainfluenza-3 (PI-3) virus, bovine respiratoryyncytial virus (BRSV) and bovine viral diarrhea virus (BVDV) [5,6].hese pathogens, together with Mannheimia haemolytica and Pas-eurella multocida, contribute to the development of shipping fever,ith each pathogen having a differing role in disease develop-ent. Infection with IBR, PI-3 and BRSV results in the destruction

f ciliated respiratory epithelium and each are considered pri-ary respiratory pathogens [5,7]. Although BVDV is not a primary

athogen in the pathogenesis of BRD, it can suppress the hostmmune system and increase the risk of secondary bacterial infec-ions, thus enhancing colonization of the lungs by other pathogens5,8].

∗ Corresponding author. Tel.: +1 913 422 6889; fax: +1 913 422 6074.E-mail address: wenzhi.xue@spcorp.com (W. Xue).

264-410X/$ – see front matter © 2010 Elsevier Ltd. All rights reserved.oi:10.1016/j.vaccine.2010.03.043

Modified-live virus (MLV) vaccines for BVDV, IBR, PI-3 andBRSV are commercially available for the prevention and controlof infections associated with these viruses [9–12]. A majority ofcommercial MLV vaccines targeting these viruses are administeredsubcutaneously, and are capable of inducing protective immuneresponses against BVDV, IBR, PI-3 and BRSV, as well as effectivelycontrolling respiratory diseases associated with these viruses. Inthe current immunization program for most cattle producers, theseparenteral MLV vaccines are used in cattle 3 months of age or older,when maternal immunity has waned to the point that it may haveno affect on immunization.

Attempts to use mucosal vaccination strategies for MLV vaccinesagainst BRD antigens have been reported and were shown to be effi-cacious [13,14]. There are two distinct advantages to vaccinatingyoung calves intranasally, instead of the subcutaneous route. First,intranasal (mucosal) immunization may be more likely to prime ayoung calf’s immune system successfully in the presence of ongo-ing maternal antibodies [13]. Second, antigens delivered mucosally

are more likely to prevent infection rather than just reduce diseaseoccurrence [13,15]. This is especially true for IBR, PI-3 and BRSV,since these viruses infect and replicate in mucosal epithelial cells[5,7]. There are currently no modified-live BVDV, IBR, PI-3 and BRSVcombination vaccines licensed for intranasal use. The purpose of

ne 28

tt2b

2

2

wfTtAaWCwcccl

2

tiaati

2

twsaSiaccodtI

2

p1toirrr

W. Xue et al. / Vacci

hese studies was to test the efficacy of a MLV vaccine, adminis-rated intranasally in 3–8 day old calves, against BVDV types 1 and, IBR, PI-3 and BRSV infections and the respiratory diseases causedy these pathogens.

. Materials and methods

.1. Calves

Colostrum-deprived (CD) calves (JR Livestock Inc, Lake Mills, IA)ere used in BVDV, IBR and PI-3 studies. The calves were removed

rom dams at birth before taking colostrum and fed milk placer.he CD calves were housed in individual hutches before vaccina-ion and in separated pens after vaccination at 3–8 days of age.ll calves were virus neutralizing (VN) antibody-free to BVDV, IBRnd PI-3 at vaccination. The BRSV study was conducted at theestern College of Veterinary Medicine (Saskatoon, Saskatchewan,

anada). Neonatal Holstein calves, obtained from local dairies,ere removed from dams at birth, and the calves were fed bovine

olostrum that contained minimal antibody to BRSV, so that thealves had antibodies to BVDV, IBR and PI-3, but not to BRSV. Thesealves were kept outdoors in individual pens until the time of chal-enge.

.2. Vaccination

A combination viral vaccine1, containing modified-live BVDVypes 1 and 2, IBR, PI-3 and BRSV, as well as, avirulent M. haemolyt-ca and P. multocida, was given intranasally as a single, 1 mL doset 3–8 days of age. All antigens in the vaccine serial were batchedt their minimum protective dose (MPD) level. In all studies, con-rol animals received 1 mL of sterile water diluent, administeredntranasally as well.

.3. Challenge viruses

BVDV NY-1, a non-cytopathic (ncp) type 1b strain, was used ashe BVDV type 1 challenge virus. BVDV 1373, an ncp type 2 strain,as used as challenge virus in the BVDV type 2 study. The Cooper

train of IBR was used to challenge calves in the IBR study. The BVDVnd IBR challenge viruses were obtained from National Veterinaryervices Laboratories (NVSL, Ames, IA). The challenge virus usedn the PI-3 study, Reisinger SF-4, was originally provided by NVSLnd had been previously passed in animals. The lung lavage fluidollected from these animals was used as PI-3 challenge virus. Thehallenge inoculum for the BRSV study consisted of a lung washbtained from a newborn calf infected with BRSV, as previouslyescribed [16]. The PI-3 and BRSV lung lavage fluids were confirmedo be free of bacterial contamination, Mycoplasma spp., BVDV andBR by standard 9CFR tests.

.4. Clinical assessment

Calves were observed for clinical signs on days −1 and 0,rior to challenge, and days 1 through 14 after challenge (daysthrough 8 for BRSV only). Clinical assessments were made at

he same time each morning by investigators who were unaware

f the treatment codes in each study. Clinical signs monitoredncluded depression, nasal and ocular discharges, coughing andectal temperatures. Clinical signs of diarrhea and mucosal hemor-hage for BVDV, nasal lesions and plaques for IBR, and respiratoryate for BRSV were also recorded in their respective studies.

1 Vista Once SQ, Intervet Inc. Millsboro, Delaware, USA.

(2010) 3784–3792 3785

For the BRSV study, calves were euthanized by a barbiturateoverdose of Euthanyl Forte (MTC Pharmaceuticals, Cambridge,ON, Canada) 8 days after challenge. Moreover, in all studies,if calves were observed at any time with severe signs of res-piratory distress, including pronounced open-mouthed, laboredbreathing (>100 breathes/min), severe signs of depression and/orrecumbence with total reluctance to rise, or with PaO2 val-ues <45 mm Hg (range 40–45 mm Hg), they were euthanizedimmediately according to USDA/CFIA Animal Care Committeeguidelines.

2.5. Sample collection

Blood samples were collected at the day of vaccination, chal-lenge and 14 days post-challenge, and SN antibody titers weredetermined for each virus. Deep nasal swab specimens wereobtained from both nares at 1-day prior to challenge through 10days post-challenge for BVDV, IBR and PI-3 studies. After collection,swabs were placed in 3 mL of cold transport medium consisting ofDulbecco’s Modified Eagle Medium (HyClone, Logan, UT) supple-mented with 10% horse serum (Sigma, St. Louis, MO), 1% gelatin(Fisher Scientific, Fair Lawn, NJ), 1× Antibiotic Antimycotic Solu-tion (Sigma, St. Louis, MO) and 30 �g/mL Gentamicin (Sigma, St.Louis, MO). Collection of nasal swabs in the BRSV study started1-day prior to challenge and ended 8 days after challenge. Swabspecimens for BRSV isolation were placed in 1 mL of transportmedium as described previously [13]. All swab specimens werestored at −70◦ C or below until they were cultured for quantita-tive virus isolation. For the BVDV studies, white blood cell (WBC)and platelet counts were conducted from 2 days pre-challengethrough 8 days post-challenge by a clinical pathology laboratory(Physicians Reference Laboratory, Overland Park, KS). In the BRSVstudy, arterial blood samples were collected from the caudal tho-racic aorta, and oxygen tension (mm Hg) measurements wereperformed by use of a gas analyzer (Ciba-Corning, Medfield, MA)[13].

2.6. Virus neutralizing antibody analysis

The virus neutralizing (VN) antibody titers to BVDV, IBR and PI-3 were measured by use of a standard microplate VN procedure.Briefly, two-fold dilutions of each serum sample were made on 96-well tissue culture plates, and approximately 100–200 TCID50 (50%tissue culture infectious dose) of each respective virus was addedto each serum dilution. After 3 days of incubation on bovine kidneycell monolayers at 37 ◦C, with 5% CO2, the plates were observedfor cytopathic effect (CPE). The neutralizing antibody titer of eachsample was determined using standard laboratory methods. Bovinerespiratory syncytial virus-specific IgG ELISAs were performed aspreviously described to determine BRSV antibody endpoint titers[13].

2.7. Quantitative virus isolation

For BVDV, IBR and PI-3 studies, virus isolation from nasal swabswas conducted using bovine kidney cell monolayers in 96-well tis-sue culture plates. Briefly, following centrifugation of samples, thesupernatants were used to infect cell monolayers for virus quantita-tion. After 3 days of incubation at 37 ◦C, with 5% CO2, the TCID50/mL

was calculated for IBR and PI-3 viruses by CPE observation of cellmonolayers. The titers of BVDV type 1 and type 2 were calculated byimmunofluorescence of cell cultures stained by BVDV type-specificmonoclonal antibodies. BRSV shedding was quantitatively deter-mined by a plaque assay method using bovine embryonic lung (BEL)fibroblasts as previously described [17].

3 ine 28

2

wmtiag

2

d2

c1ac1eEv

Coceisil

vatafAI

(rcdwpwa

gcwaiAc

2

tccl

786 W. Xue et al. / Vacc

.8. Postmortem analysis

At the end of the BRSV study, the respiratory tract of each calfas collected and analyzed to determine the percentage of pneu-onic tissue as previously described [18]. Tracings were made of

he dorsal surface, and the percentage area of pneumonic lung,ncluding characteristic atelectatic, as well as, emphysematousreas for each tracing was determined by a software imaging pro-ram (Universal Imaging Corp., West Chester, PA).

.9. Experimental design

Five separate immunogenicity studies were implemented in 3–8ay old CD calves for each viral antigen: BVDV type 1, BVDV type, IBR, PI-3 and BRSV.

For the BVDV type 1 study, 15 calves were intranasally vac-inated with a single dose of the MLV vaccine as described, and6 calves received sterile water diluent intranasally. Three weeksfter vaccination, the calves were commingled and intranasallyhallenged with an aerosolized virulent type 1 BVDV strain NY-. The challenge was performed by spraying 2 mL of virus intoach nostril, using a Devilbiss Atomizer (Devilbiss, Somerset, PA).ach animal received approximately 5 × 106 TCID50 of challengeirus.

A total of 29 calves were enrolled in the BVDV type 2 study.alves in the vaccinate group (n = 14) received a single 1 mL dosef the MLV vaccine administrated intranasally, and calves in theontrol group (n = 15) received a 1 mL dose of sterile water dilu-nt. On day 21 post-vaccination, the calves were commingled andntranasally challenged with an aerosolized virulent type 2 BVDVtrain 1373. The challenge was performed by spraying 2 mL of virusnto each nostril, using a Devilbiss Atomizer. Each calf was chal-enged with approximately 1.6 × 105 TCID50 of challenge virus.

In the IBR study, 16 calves received a single 1 mL dose of the MLVaccine administrated intranasally, and 14 control calves received1 mL dose of sterile water diluent. Four weeks post-vaccination,

he calves were commingled and intranasally challenged with anerosolized virulent IBR virus Cooper strain. The challenge was per-ormed by spraying 2 mL of virus into each nostril, using a Devilbisstomizer. Each animal received approximately 1.3 × 105 TCID50 of

BR challenge virus.For the PI-3 study, 31 calves were randomized into vaccinate

n = 15) and control groups (n = 16). Calves in the vaccinate groupeceived a single 1 mL intranasal dose of the MLV vaccine, andalves in control group were given a 1 mL dose of sterile wateriluent intranasally. Twenty-eight days after vaccination, the calvesere mixed and intranasally challenged with an aerosolized in vivoassed PI-3 lung wash using a Devilbiss Atomizer. The challengeas performed by spraying 2 mL of virus into each nostril. Each

nimal received approximately 1.6 × 107 TCID50 of PI-3 virus.Finally, for the BRSV study, nine calves received a 1 mL sin-

le dose of the MLV vaccine intranasally, whereas seven controlalves received 1 mL of sterile water diluent intranasally. The calvesere commingled and challenged 21 days after vaccination via

erosol delivery using Ultra-Neb 99 ultrasonic nebulizers (Dev-lbiss, Somerset, PA) in the procedure described previously [13].pproximatelly 106.2 PFUs of virulent BRSV were used to challengealves.

.10. Data analysis

In each study, the two treatment groups (vaccinate and con-rol) were analyzed and compared with respect to the primarylinical signs including rectal temperature, nasal and ocular dis-harges, depression, leukopenia, oral or nasal mucosal lesions, lungesions, pO2 values and virus shedding, using PROC NPAR1WAY of

(2010) 3784–3792

SAS® Version 9.1.3 to perform the Fisher Exact test. The preventedfraction and associated confidence interval were estimated.

3. Results

3.1. The intranasal administration of the MLV vaccine in 3–8 dayold calves produced a measurable antibody response

All colostrum-deprived calves were free of VN antibodies to eachof the five viruses at the time of vaccination (Table 1). At challenge,nearly all vaccinated calves exhibited minimal VN antibody titers toBVDV types 1 and 2, IBR and PI-3. ELISA antibody titers to BRSV werethe same at challenge as they were at the time of vaccination. Uponconclusion of each challenge observation period, VN titers to eachvirus were significantly (p < 0.05) higher in the vaccinated calveswhen compared to control calves. These results demonstrated thatcalves were primed by intranasal vaccination with the MLV vaccine,even though there was an absence of significant antibody titers atthe time of challenge. Similar antibody results were also obtainedin the BRSV ELISA.

3.2. Efficacy against BVDV type 1 challenge

After challenge with BVDV type 1 strain NY-1, all control calves(16/16) developed clinical diseases associated with BVDV infection,including nasal discharge, coughing, diarrhea and depression afterday 5 post-challenge (Fig. 1A). In contrast, only one-fifth (3/15) ofvaccinated calves developed clinical signs, which were minimal innature. The prevented fraction was 80% for the vaccinate groupand the statistical analysis indicated that the total average dailyclinical score in the control group was 16.8, which is significantly(p < 0.01) higher than that for the vaccinate group (daily averageclinical score of 1.2). The rectal temperatures of calves were mea-sured daily from days −1 to 14 post-challenge. All control calveshad temperatures higher than 40 ◦C during the observation period,while only 3 of 15 vaccinates developed fever (Fig. 1B). Moreover,the vaccinated group had significantly (p < 0.05) lower tempera-tures at days 7–9 post-challenge than the control group, when thehighest rectal temperatures were recorded.

BVDV infection usually results in leukopenia. Results of WBCcounts demonstrated a decrease beginning on day 2 post-challenge.The WBC decrease in control calves (>45%) was more pronouncedthan in MLV-vaccinated calves (<20%) (Fig. 1C), and WBC countsin vaccinate group were significant higher (p < 0.05) than controlgroup at days 3 and 4 post-challenge. In addition, the platelet countdropped beginning at day 2 post-challenge for all calves; however,the control calves displayed a greater decline in platelet countswhen compared to the vaccinated calves (Fig. 1D). Specifically,the vaccinate group showed significantly (p < 0.05) higher plateletcounts at days 6, 7 and 8 post-challenge than the control calves.

Nasal swab samples were collected from challenged calves fromday −1 to day 10 post-challenge for detection of virus shedding. Allcalves were negative for virus isolation prior to challenge, the dayof challenge and for the first 2 days following challenge (Fig. 1E).Statistical analysis of virus isolation, in terms of positive or nega-tive, and the total number of days positive, indicate that there weresignificantly (p < 0.01) less vaccinates (4/15) that shed virus thancontrols (16/16). Additionally, the MLV-vaccinated calves werepositive for significantly (p < 0.01) fewer days (mean = 0.4 days)compared to the controls (mean = 4.38 days). The titers of isolatedBVDV1 from controls were significantly (p < 0.05) higher than vac-cinates at days 5–10 post-challenge as well.

3.3. Efficacy against BVDV type 2 challenge

BVDV type 2 challenge strain 1373 is a virulent strain andcauses severe clinical disease. All control calves (15/15) developed

W. Xue et al. / Vaccine 28 (2010) 3784–3792 3787

Table 1Induction of a virus neutralizing antibody or ELISA antibody response to each vaccine virus in 3–8 day old calves, intranasally vaccinated with a MLV vaccine.

Antigens target Treatment groups Antibody titers* at different days

Vaccination Challenge Termination

BVDV1 Vaccinate 0 0 24.8Control 0 0 6.7

BVDV2 Vaccinate 0 1.4 68.7Control 0 0 2.3

IBR Vaccinate 0 1.6 7.0Control 0 0 3.5

PI-3 Vaccinate 0 1.2 35.3Control 0 0 13.4

BRSV Vaccinate 4.0 4.3 9.3Control 4.0 3.9 3.0

* The virus neutralizing antibody titers listed are the geometric mean titers (GMT) to BVDV types 1 and 2, IBR, and PI-3 viruses; and ELISA unit titers to BRSV.

Fig. 1. Intranasal vaccination protected young calves from clinical diseases caused by type 1 BVDV challenge. The vaccinated calves were protected from development ofsevere clinical signs (A); elevated rectal temperatures (B); leukopenia (C); decrease of platelet counts (D); and virus shedding (E). Statistical values are *p < 0.05 and **p < 0.01.

3788 W. Xue et al. / Vaccine 28 (2010) 3784–3792

F by tys ase oft l valu

snIc(dMic4trhw

bt

ig. 2. Intranasal vaccination protected young calves from clinical diseases causedevere clinical diseases (A); elevated rectal temperatures (B); leukopenia (C); decreype 2 BVDV challenge was significantly decreased by the vaccination (F). Statistica

evere clinical disease associated with BVDV2 infection, includingasal discharge, coughing, diarrhea and depression post-challenge.

n comparison, fewer vaccinated calves (6/14) developed clini-al signs from the challenge, which were moderate in natureFig. 2A). Statistical analysis of depression, diarrhea and nasalischarge demonstrated prevented fractions of 70–100% for theLV-vaccinates, indicating the immunogenicity of the vaccine

n reducing clinical disease caused by BVDV2 challenge. Post-hallenge, all control calves developed temperatures higher than0 ◦C for 4–10 days. However, only 3 of 14 vaccinated calves showedemperatures higher than 40 ◦C for only 1–3 days (Fig. 2B). As aesult, the proportion of calves with fever (≥40 ◦C) was significantly

igher (p < 0.01) in the control group versus the vaccinate group,ith a prevented fraction of 79% for the MLV-vaccinates.

Results of WBC counts showed that the counts of control calvesegan decreasing on day 2 post-challenge, with a greater reduc-ion from days 3 through 8, when compared to MLV-vaccinates

pe 2 BVDV challenge. The vaccinated calves were protected from development ofplatelet counts (D); and virus shedding (E). Fatality of calves caused by the virulentes are *p < 0.05 and **p < 0.01.

(Fig. 2C). The vaccinate group had a slight overall decrease inWBC counts (about 20% drop) from days 3 to 6, then recoveredto normal levels. However, the control group did not fully recoverits WBC count to normal levels during the challenge observationperiod. The difference between control and MLV-vaccinated calvesin WBC counts was significant (p < 0.05), with control calves devel-oping leukopenia from the challenge. Platelet counts started todecrease on day 2 post-challenge for all calves (Fig. 2D). However,the decrease was significantly different (p < 0.05) between the con-trol and vaccinate calves. The mean platelet count of control calvesdecreased from 820 K/mm3 at challenge day to <256 K/mm3 at day8 post-challenge. In contrast, vaccinated calves had platelet counts

decrease from 700 K/mm3 at challenge day to 531 K/mm3 at day 8post-challenge.

Nasal swab samples were collected from challenged calves fromday −1 to day 10 post-challenge for virus shedding detection. Allcontrol calves shed virus at multiple days during the challenge

ne 28

pcSttsp7

ocpds

3

didic(svd

a4ad

Fc

W. Xue et al. / Vacci

eriod, with high titers of shed virus compared to MLV-vaccinatedalves, which shed virus for a much shorter period after challenge.tatistical analysis of the virus isolation results demonstrated thathere were significantly (p < 0.01) less vaccinates that shed virushan controls. Furthermore, the BVDV titers from controls, wereignificantly (p < 0.05) higher than vaccinates at days 5 through 9ost-challenge (Fig. 2E), with peak viral shedding occurring at daysand 8 post-challenge.

Due to the BVDV2 challenge, 9 of 15 control animals died,r were euthanized for humane reasons, on days 10–14 post-hallenge (Fig. 2F). In the vaccinate group, one calf died on day 11ost-challenge, but necropsy results could not confirm the cause ofeath to be associated to the BVDV2 challenge. Statistical analysishowed a prevented fraction of 88% for the MLV vaccination group.

.4. Efficacy against IBR challenge

After challenge with IBR Cooper strain, 14 of 14 control calveseveloped clinical signs associated with IBR infection, includ-

ng nasal lesions, nasal discharge, ocular discharge, coughing andepression. Total clinical scores were significantly (p < 0.05) higher

n the control calves than the vaccinated calves (Fig. 3A). Specifi-ally, control calves (14/14) showed significantly more nasal lesionsp < 0.01) than vaccinated calves (4/16). Eleven of 14 control calveshowed nasal discharge on various days, but only 3 of 16 MLV-accinated calves displayed nasal discharge. This difference in nasalischarge between control and vaccinates was significant (p < 0.01).

All control calves showed elevated rectal temperatures >40 ◦C,nd 13 of 14 controls had rectal temperatures of 40–41.2 ◦C for–11 days after challenge, with temperatures peaking at days 5nd 6 post-challenge (Fig. 3B). In contrast, 4 of 16 vaccinated calveseveloped elevated temperatures of >40 ◦C, each for only 1 day. The

ig. 3. Intranasal vaccination protected young calves from development of severe clinicahallenge. Statistical values are *p < 0.05 and **p < 0.01.

(2010) 3784–3792 3789

proportion of calves with fever (>40 ◦C) was significantly (p < 0.01)different between vaccinate and control groups.

Virus isolated from nasal swabs at day 1 post-challenge wasprobably residual challenge virus, while the virus isolated at day 2post-challenge and thereafter was likely replicating virus. Althoughall animals, both controls and vaccinates, shed viruses at variousdays after challenge, results demonstrated that control calves shedhigher amounts of the IBR virus and for a longer time (Fig. 3C).The maximum post-challenge titers of isolated viruses were signif-icantly higher (p < 0.01) from control calves (8.2 log10 TCID50/mL)than from the vaccinated calves (6.4 log10 TCID50/mL).

3.5. Efficacy against PI-3 virus challenge

PI-3 typically does not cause severe disease in cattle. After chal-lenge, the majority of the control animals (14/16) developed clinicalsigns of respiratory disease, including dyspnea, coughing and nasaldischarge (Fig. 4A). In contrast, clinical signs were observed in only2 of 15 vaccinated calves during the observation period. The per-centage of calves developing clinical disease associated with PI-3challenge was significantly lower (p < 0.01) in the vaccinate groupversus the control group, with an estimated prevented fraction of85% for the MLV vaccination group.

In addition, the control calves developed higher rectal temper-atures after PI-3 challenge, compared to MLV-vaccinated calves.Elevated rectal temperatures were observed in control animals

from day 2 post-challenge, returning to levels comparable to thevaccinated group after day 6 post-challenge (Fig. 4B). Statisticalanalysis demonstrated that the maximum elevated temperaturewas significantly higher (p < 0.01) in the control group than in thevaccinated group.

l signs (A); elevated rectal temperatures (B); and virus shedding (C) caused by IBR

3790 W. Xue et al. / Vaccine 28 (2010) 3784–3792

F gns (AS

ecsmtvctc

3

ofipdcsit(g

pa(g

aT

ig. 4. Intranasal vaccination protected young calves from development of clinical sitatistical values are *p < 0.05 and **p < 0.01.

Nasal swab samples for virus isolation were collected fromach calf prior to challenge (days −1 and 0) and days 1–10 post-hallenge. The results demonstrated that the control calves shedignificantly (p < 0.01) more virus than the vaccinated calves. Theedian PI-3 titer isolated from the vaccinated group was less than

hat in the control group. Statistical analysis of the duration ofirus shedding showed that the vaccinated group had a signifi-antly (p < 0.01) shorter duration of virus shedding (2 days) thanhe control group (6 days). Fig. 4C presents the virus titers shed byalves from both vaccinate and control groups.

.6. Efficacy against BRSV challenge

After challenge with virulent BRSV, the control animals devel-ped clinical signs of severe respiratory disease, including highever, high respiratory rates, dyspnea, depression, anorexia, cough-ng and nasal discharge beginning at approximately day 3ost-challenge and continuing through to the end of the study onay 8 post-challenge. Vaccinated calves developed milder clini-al disease and showed significantly lower (p < 0.05) total clinicalcores than the controls following challenge. Respiratory ratesncreased in both groups of challenged calves (Fig. 5A). Althoughhe difference in maximum respiratory rates was not significantp = 0.14) between vaccinated and control groups, the vaccinatedroup displayed a lower respiratory rate at days 5–8 post-challenge.

Arterial blood oxygen concentration (PO2) is a key indicator ofulmonary function. Arterial blood was collected from all calvest day 6 post-challenge, and the (PO2) in the vaccinated calves

80.7 mm Hg) was significantly (p < 0.05) higher than in the controlroup (65.2 mm Hg) (Fig. 5B).

Nasal swab samples for virus isolation were collected from eachnimal prior to challenge and on days 2–8 days post-challenge.he samples were titrated on BEL cells and titers of shed virus

); elevated rectal temperatures (B); and virus shedding (C) caused by PI-3 challenge.

were expressed as plaque-forming units per mL (PFU/mL). Titrationresults demonstrated that the control animals shed significantly(p < 0.05) more BRSV than the vaccinated animals, and the medianBRSV titer for the vaccinated group was less than the control group.Furthermore, virus shedding was significantly (p < 0.05) higher incontrol calves on days 4–7 post-challenge, compared to the vacci-nated calves (Fig. 5C).

Lungs were collected six days after challenge from the two euth-anized calves (1 each from vaccinated and control groups), andeight days after challenge from all of the remaining calves, andpneumonic lesions were measured in the lungs. Although the lunglesion scores of the vaccinated group (28.1%) was not significantlydifferent (p = 0.07) than the control group, it was still measurablylower compared to the score of control calves (40%) (Fig. 5D).

4. Discussion

Currently available parenteral combination MLV vaccinesagainst BVDV types 1 and 2, IBR, PI-3 and BRSV can clearly induceprotective responses against these viral pathogens [9,11,12]; how-ever, no comparable combination vaccine is commercially availablefor intranasal administration. The development of a multivalentMLV vaccine for mucosal immunization has potential advantagescompared to traditional parenteral vaccination. First, given thevariability in passive transfer of maternal immunoglobulin, it is dif-ficult to predict when parenteral vaccination of young calves willconfer immune protection from these viruses. Mucosal immuniza-tion may be more likely to immunologically prime young calves in

the face of maternal antibodies. In that case, whenever the mater-nal antibodies decay to nonprotective concentrations, intranasallyvaccinated calves would have active immunity, and avoid the “win-dow of susceptibility” to virus infection [15]. A second advantage isthat mucosal vaccination is more likely to prevent viral infection,

W. Xue et al. / Vaccine 28 (2010) 3784–3792 3791

F ses ca( od oxa

rl

ttmeicpnmtiiataBdwttct

sTvi

ig. 5. Intranasal vaccination protected young calves from clinical respiratory diseaA), titers of shed virus (C), and percentages of lung lesions (D). After challenge, blore *p < 0.05 and **p < 0.01.

ather than simply reducing disease, by preferentially stimulatingocal (IgA) responses [15,19].

Many commercial MLV vaccines are available for control ofhe BRD complex, but nearly all of the vaccines are adminis-ered by either subcutaneous or intramuscular routes. Combination

odified-live intranasal IBR and PI-3 vaccines have documentedfficacy [20–22] and are the only vaccines currently licensed forntranasal administration in cattle [23,24]. In fact, intranasal vac-ination using IBR and PI-3 vaccines has been accepted as a goodractice for many years within the cattle industry [25,26]. Immu-ization via mucosal administration with BRSV vaccines, eitheronovalent or multivalent, has been reported [13,15]. Results of

he studies reported in this paper indicate that the vaccine used formmunization was completely efficacious in intranasally immuniz-ng against all viruses contained in the product. To our knowledge,t the time of preparation of this manuscript, the vaccine used inhis paper is the only one currently approved by the regulatoryuthorities to be used intranasally to against BVDV, IBR, PI3 andRSV pathogens. This vaccine induced immune responses in 3–8ay old calves that significantly reduced disease after challengeith IBR, PI-3 and BRSV, as well as BVDV types 1 and 2. Moreover,

o our knowledge, results of this study document for the first timehat intranasal vaccination can be successfully employed to signifi-antly reduce disease resulting from challenge with virulent BVDVypes 1 and 2 in young calves.

A feature of BVDV infection resulting from both natural expo-ure, and, potentially, vaccination, is immunosuppression [27–29].herefore, a concern was that intranasal inoculation using a MLVaccine containing both BVDV types 1 and 2 might create a risk ofmmunosuppression. In this study, vaccinated calves did not show

used by BRSV challenge. Vaccinated calves had significantly lower respiratory ratesygen tension values in control calves significantly decreased (B). Statistical values

any clinical illness post-vaccination, and, importantly, as well, hadno decrease in WBC counts. These findings indicate that immuno-suppression is not likely to be a safety concern with this MLVvaccine.

The MLV vaccine studied here was a multivalent vaccine, includ-ing avirulent cultures of P. multocida and M. haemolytica bacterialantigens. These two antigens also induced strong immunity inyoung calves and protected the calves from challenges with virulentP. multocida and M. haemolytica strains (unpublished data).

In previous studies with BRSV, mucosal immunization pro-vided clinical protection associated with an anamnestic mucosalIgA response, without a notable systemic IgG response at chal-lenge [13,18]. Similar results were demonstrated herein, in thatno significant VN antibodies were detected in the serum to any ofthe viruses in vaccinated calves at the time of challenge; however,serological VN antibody titers to BVDV types 1 and 2, IBR and PI-3 viruses all increased anemnestically following challenge. Sincemucosal IgA and cell-mediated immune responses, including IFN-�, were not monitored in the current studies, no conclusions canbe reached regarding which immune effector responses were asso-ciated with the observed disease-sparing. It is possible that morethan one vaccine-stimulated immunologic mechanism can conferclinical protection from different virulent virus challenges. Never-theless, the vaccine used in this study has previously been shownto induce antigen-specific T-cell subset activation to each of the

five viruses in parenterally vaccinated calves [30]. Together, thesefindings clearly demonstrate that intranasal administration of thisvaccine to young calves primes immune responses, including mem-ory responses that reduce disease resulting from challenge witheach of the viruses. Practically, cattle producers like to physically

3 ine 28

hitiba(

cushaPabetp

A

cUraT

R

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

792 W. Xue et al. / Vacc

andle the calves as fewer times as possible to avoid the stressest may bring to the animal. Therefore, it is much easier for themo vaccinate young calves intranasally and wait until the wean-ng of calves for a second vaccination, either by subcutaneous ory intranasal route. The MLV vaccine in this study demonstrated6 months of duration of immunity to BVDV and IBR antigens

unpublished data) when administered intranasally.In conclusion, the demonstration that combination viral vac-

ines, containing IBR, PI-3, BRSV and BVDV types 1 and 2, can besed intranasally against each of these viral pathogens has sub-tantial clinical importance. For many years, this type of vaccineas been used parenterally in cattle 3 months of age or older, as anid in the prevention of respiratory diseases associated with IBR,I-3 and BRSV, and BVDV types 1 and 2. Due to the expected vari-bility in passive immunity in cattle populations, many calves wille susceptible to infection prior to this time. These results extendfficacy to calves as young as 3 days old, thereby providing a toolo aid in closing the window of susceptibility between the neonataleriod and weaning.

cknowledgements

All host animal immunogenicity studies in this manuscript wereonducted at the Intervet Inc animal research facility and at theniversity of Saskatchewan, following ethical guidelines from the

espective regulatory agencies, USDA and CFIA. The authors thankll technicians involved in the animal care and sample collections.he authors also thank David Whiteman for statistical data analysis.

eferences

[1] Kelly AP, Janzen ED. A review of morbidity and mortality rates and diseaseoccurrence in North America feedlot cattle. Can Vet J 1986;27:496–500.

[2] Barrett D. Bovine respiratory disease–a clinician’s perspective. Cattle Pract1998;6:251–5.

[3] Caldow G, Nettleton P. Pneumonia: identifying the cause agent. Cattle Pract2000;8:131–4.

[4] Scott P. Epidemiology and treatment of bovine respiratory in beef cattle. CattlePract 1997;5:283–8.

[5] Cusack PMV, McMeniman N, Lean IJ. The medicine and epidemiology of bovinerespiratory disease in feedlots. Aust Vet J 2003;81:480–7.

[6] Martin SW, Bohac JG. The association between serological titers in infectiousbovine rhinotracheitis virus, bovine viral diarrhea virus, parainfluenza-3 virus,respiratory syncytial virus and treatment for respiratory disease in Ontariofeedlot calves. Can J Vet Res 1986;50:351–8.

[7] Baker JC, Frey ML. Bovine respiratory syncytial virus. Vet Clin North Am LargeAnim Pract 1985;1:259–75.

[8] Richer L, Marois P, Lamontagne L. Association of bovine viral diarrhoea viruswith multiple viral infections in bovine respiratory disease outbreaks. Can Vet

J 1988;29:713–7.

[9] Fulton RW, Confer AW, Burge LJ, Perino LJ, d’Offay JM, Payton ME, et al. Antibodyresponses by cattle after vaccination with commercial viral vaccines containingbovine herpesvirus-1, bovine viral diarrhea virus, parainfluenza-3 virus, andbovine respiratory syncytial virus immunogens and subsequent revaccinationat day 140. Vaccine 1995;13:725–33.

[

[

(2010) 3784–3792

10] Schunicht OC, Booker CW, Jim GK, Guichon PT, Wildman BK, Hill BW. Com-parison of a multivalent viral vaccine program versus univalent viral vaccineprogram on animal health, feedlot performance, and carcass characteristics offeedlot calves. Can Vet J 2003;44:43–50.

11] Peters AR, Thevasagayam SJ, Wiseman A, Salt JS. Duration of immunity of aquadrivalent vaccine against respiratory disease caused by BHV-1, PI3V, BVDV,and BRSV in experimentally infected calves. Prev Vet Med 2004;66:63–77.

12] Donkersgoed JV, van der Hurk JV, McCartney D, Harland RJ. Comparativeserological responses in calves to eight commercial vaccines against infec-tious bovine rhinotracheitis, parainfluenza-3, bovine respiratory syncytial, andbovine viral diarrhea viruses. Can Vet J 1991;32:727–33.

13] Ellis J, Gow S, West K, Waldner C, Rhodes C, Mutwiri G, et al. Responsesof calves to challenge exposure with virulent bovine respiratory syncytialvirus following intranasal administration of vaccines formulated for parenteraladministration. J Am Vet Med Assoc 2007;230:233–43.

14] Gogey S, Fays KD, Versali MF, Gautier S, Thiry E. Glycol chitosan improves theefficacy of intranasally administrated replication defective human adenovirustype 5 expressing glycoprotein D of bovine herpesvirus 1. Vaccine 2004;22:1946–53.

15] Kimman TG, Westenbrink F, Staver PJ. Priming for local and systemic antibodymemory responses to bovine respiratory syncytial virus: effect of amount ofvirus, virus replication, route of administration and maternal antibodies. VetImmunol Immunopathol 1989;22:145–60.

16] West K, Petrie L, Haines DM, Konoby C, Clark EG, Martin K, et al. The effectof formalin-inactivated vaccine on respiratory disease associated with bovinerespiratory syncytial virus infection in calves. Vaccine 1999;17:809–20.

17] West K, Bogdan J, Hamel A, Nayar G, Morley PS, Haines DM, et al. A comparisonof diagnostic methods for the detection of bovine respiratory syncytial virus inexperimental clinical specimens. Can J Vet Res 1998;62:245–50.

18] West K, Petrie L, Konoby C, Haines DM, Cortese V, Ellis JA. The efficacy ofmodified-live bovine respiratory syncytial virus vaccines in experimentallyinfected calves. Vaccine 1999;18:907–19.

19] Tizard I. Immunity at body surfaces. In: Veterinary Immunology, An Introduc-tion. Philadelphia: WB Saunders Co; 2004, 67–74.

20] Todd JD. Intranasal vaccination of cattle against IBR and PI3: field and labora-tory observations in dairy, beef, and neonatal calf populations. Dev BiologicalStandard 1976;33:391–5.

21] Burroughs AL, Morrill JL, Bostwick JL. Immune response of dairy cattleto parainfluenza-3 virus in intranasal infectious bovine rhinotracheitis-parainfluenza-3 virus vaccine. Can J Comp Med 1982;46:264–6.

22] Martin W, Willson P, Curtis R, Allen B, Acres S. A field trial, of preshipmentvaccination, with intranasal infectious bovine rhinotracheitis-parainfluenza-3vaccines. J Comp Med 1983;47:245–9.

23] Bowland SL, Shewen PE. Bovine respiratory disease: commercial vaccines cur-rently available in Canada. Can Vet J 2000;41:33–48.

24] Cortese V. Bovine vaccines and her vaccination programs. In: Smith B, editor.Large Animal Internal Medicine. Philadelphia: Mosby; 2002. p. 1403–11.

25] Kahrs RF. Infectious bovine rhinotracheitis and infectious pustular vulvovagini-tis. In: Kahrs RF, editor. Viral Diseases Of Cattle. Ames: Iowa State UniversityPress; 2001. p. 159–70.

26] Kahrs RF. Parainfluenza type-3. In: Kahrs RF, editor. Viral Diseases Of Cattle.Ames: Iowa State University Press; 2001. p. 185–93.

27] Bolin SR, McClurkin AW, Coria MF. Effects of bovine viral diarrhea virus on thepercentages and absolute numbers of circulating B and T lymphocytes in cattle.Am J Vet Res 1985;46:884–6.

28] Ellis JA, Davis WC, Belden EL, Pratt DL. Flow cytofluorimetric analysis of lym-phocyte subset alterations in cattle infected with bovine viral diarrhea virus.Vet Pathol 1988;25:231–6.

29] Walz PH, bell TG, Wells JL, Grooms DL. Relationship between degree of viremiaand disease manifestation in calves with experimentally induced bovine viraldiarrhea virus infection. Am J Vet Res 2001;62:1095–103.

30] Platt R, Burdett W, Roth JA. Induction of antigen-specific T-cell subset activationto bovine respiratory disease viruses by a modified-live virus vaccine. Am J VetRes 2006;67:1179–84.

本文献由“学霸图书馆-文献云下载”收集自网络，仅供学习交流使用。

学霸图书馆（www.xuebalib.com）是一个“整合众多图书馆数据库资源，

提供一站式文献检索和下载服务”的24 小时在线不限IP

图书馆。

图书馆致力于便利、促进学习与科研，提供最强文献下载服务。

图书馆导航：

图书馆首页 文献云下载 图书馆入口 外文数据库大全 疑难文献辅助工具