• View in gallery

    Mean clinical illness score (A) and rectal temperature (B) for calves that did (vaccinates; squares and short dashed line; n = 9) and did not (controls; circles and solid line; 10) receive 2 doses of a dual-adjuvanted vaccinec that contained killed strains of BHV1, BVDV types 1 (BVDV1) and 2 (BVDV2), bovine respiratory syncytial virus, and parainfluenza type 1 virus and a bacterin against 5 strains of Leptospira (Leptospira canicola, Leptospira grippotyphosa, Leptospira hardjo, Leptospira incterohemorrhagiae, and Leptospira pomona) on study days 0 and 21, as well as calves that were not vaccinated but were experimentally inoculated with a virulent strain of BHV1 (1.04 × 107 TCID50, IV; seeders; triangles and long dashed line; 4) at various times before and after initiation of a BHV1 challenge period (day 56). All calves were Holstein steers and were approximately 4 months old at study initiation (day 0). All calves were commingled in 1 pen for the duration of the study, except for the 3-day period (days 53 to 55) during which the seeders were quarantined and experimentally inoculated with BHV1. The clinical illness scores represent the total score for each group for each day. *Within a day, mean value differs significantly (P ≤ 0.05) between the control and vaccinated groups. See text for further details regarding the clinical illness scoring system.

  • View in gallery

    Mean log2 SN antibody titers against BHV1 (A), BVDV1 (B), and BVDV2 (C) at various times before and after vaccination and BHV1 exposure for the calves of Figure 1. See Figure 1 for remainder of key.

  • View in gallery

    Mean extracellular BHV1-specific (A) and BVDV-specific (B) IFN-γ concentrations at various times before and after vaccination and BHV1 exposure for the controls and vaccinates of Figure 1. See Figure 1 for remainder of key.

  • View in gallery

    Mean percentage of CD4+ (A and B), CD8+ (C and D), and WC-1+ (E and F) T cells that expressed IFN-γ following stimulation with inactivated BVDV (A, C, and E) and inactivated BHV1 (B, D, and F) at various times before and after vaccination and BHV1 exposure for the controls (gray bars) and vaccinates (black bars) of Figure 1. Notice that the scale of the y-axis varies among panels.

  • View in gallery

    Mean percentage of CD4+ (A and B), CD8+ (C and D), and WC-1+ (E and F) T cells that expressed IL-4 following stimulation with inactivated BVDV (A, C, and E) and inactivated BHV1 (B, D, and F) at various times before and after vaccination and BHV1 exposure for the controls (gray bars) and vaccinates (black bars) of Figure 1. See Figure 4 for remainder of key.

  • View in gallery

    Mean percentage of CD4+ (A and B), CD8+ (C and D), and WC-1+ (E and F) T cells that expressed CD44 following stimulation with inactivated BVDV (A, C, and E) and inactivated BHV1 (B, D, and F) at various times before and after vaccination and BHV1 exposure for the controls (gray bars) and vaccinates (black bars) of Figure 1. See Figure 4 for remainder of key.

  • 1. Babiuk LA, van Drunen Littel–van den Hurk S, Tikoo SK. Immunology of bovine herpesvirus 1 infection. Vet Microbiol 1996;53:3142.

  • 2. van Drunen Littel–van den Hurk S. Cell-mediated immune responses induced by BHV-1: rational vaccine design. Expert Rev Vaccines 2007;6:369380.

    • Search Google Scholar
    • Export Citation
  • 3. van Drunen Littel–van den Hurk S, Tikoo SK, van den Hurk JV, et al. Protective immunity in cattle following vaccination with conventional and marker bovine herpesvirus-1 (BHV1) vaccines. Vaccine 1997;15:3644.

    • Search Google Scholar
    • Export Citation
  • 4. Denis M, Kaashoek MJ, van Oirschot JT, et al. Quantitative assessment of the specific CD4+ T lymphocyte proliferative response in bovine herpesvirus 1 immune cattle. Vet Immunol Immunopathol 1994;42:275286.

    • Search Google Scholar
    • Export Citation
  • 5. Davies DH, Carmichael LE. Role of cell-mediated immunity in the recovery of cattle from primary and recurrent infections with infectious bovine rhinotracheitis virus. Infect Immun 1973;8:510518.

    • Search Google Scholar
    • Export Citation
  • 6. Campos M, Ohmann HB, Hutchings D, et al. Role of interferon-gamma in inducing cytotoxicity of peripheral blood mononuclear leukocytes to bovine herpesvirus type 1 (BHV1)–infected cells. Cell Immunol 1989;120:259269.

    • Search Google Scholar
    • Export Citation
  • 7. Tikoo SK, Campos M, Popowych YI, et al. Lymphocyte proliferative responses to recombinant bovine herpes virus type 1 (BHV-1) glycoprotein gD (gIV) in immune cattle: identification of a T cell epitope. Viral Immunol 1995;8:1925.

    • Search Google Scholar
    • Export Citation
  • 8. Guzman E, Price S, Poulsom H, et al. Bovine γδ T cells: cells with multiple functions and important roles in immunity. Vet Immunol Immunopathol 2012;148:161167.

    • Search Google Scholar
    • Export Citation
  • 9. Hoek A, Rutten VP, Kool J, et al. Subpopulations of bovine WC1+ γδ T cells rather than CD4+CD25highFoxp3+ T cells act as immune regulatory cells ex vivo. Vet Res 2009;40:6.

    • Search Google Scholar
    • Export Citation
  • 10. Rouse BT, Babiuk LA. Host defense mechanisms against infectious bovine rhinotracheitis virus; in vitro stimulation of sensitized lymphocytes by virus antigen. Infect Immun 1974;10:681687.

    • Search Google Scholar
    • Export Citation
  • 11. Endsley JJ, Quade MJ, Terharr B, et al. BHV-1–specific CD4+, CD8+, and γδ T cells in calves vaccinated with one dose of a modified live BHV-1 vaccine. Viral Immunol 2002;15:385393.

    • Search Google Scholar
    • Export Citation
  • 12. 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.

    • Search Google Scholar
    • Export Citation
  • 13. Lemaire M, Schynts F, Meyer G, et al. Latency and reactivation of a glycoprotein E negative bovine herpesvirus type 1 vaccine: influence of virus load and effect of specific maternal antibodies. Vaccine 2001;19:47954804.

    • Search Google Scholar
    • Export Citation
  • 14. Chase CCL, Fulton RW, O'Toole D, et al. Bovine herpesvirus 1 modified live virus vaccines for cattle reproduction: balancing protection with undesired effects. Vet Microbiol 2017;206:6977.

    • Search Google Scholar
    • Export Citation
  • 15. Stevens ET, Zimmerman AD, Buterbaugh RE, et al. The induction of a cell-mediated immune response to bovine viral diarrhea virus with an adjuvanted inactivated vaccine. Vet Ther 2009;10:E1E8.

    • Search Google Scholar
    • Export Citation
  • 16. Zimmerman AD, Buterbaugh RE, Herbert JM, et al. Efficacy of bovine herpesvirus-1 inactivated vaccine against abortion and stillbirth in pregnant heifers. J Am Vet Med Assoc 2007;231:13861389.

    • Search Google Scholar
    • Export Citation
  • 17. Newcomer BW, Cofield LG, Walz PH, et al. Prevention of abortion in cattle following vaccination against bovine herpesvirus 1: a meta-analysis. Prev Vet Med 2017;138:18.

    • Search Google Scholar
    • Export Citation
  • 18. Woolums AR, Siger L, Johnson S, et al. Rapid onset of protection following vaccination of calves with multivalent vaccines containing modified-live or modified-live and killed BHV-1 is associated with virus specific interferon gamma production. Vaccine 2003;21:11581164.

    • Search Google Scholar
    • Export Citation
  • 19. Manual of standards for diagnostic tests and vaccines for terrestrial animals. Paris: Office International des Epizooties, 2015;1415.

    • Search Google Scholar
    • Export Citation
  • 20. Bissey LL, Williams AK, Bolin S, et al. Comparison of cytopathic and noncytopathic isolates of bovine viral diarrhea virus by oligonucleotide fingerprinting. J Vet Diagn Invest 1991;3:1621.

    • Search Google Scholar
    • Export Citation
  • 21. Littell RC, Henry PR, Ammerman CB. Statistical analysis of repeated measures data using SAS procedures. J Anim Sci 1998;76:12161231.

  • 22. Quade MJ, Roth JA. Antigen-specific in vitro activation of T-lymphocyte subsets of cattle immunized with a modified live bovine herpesvirus 1 vaccine. Viral Immunol 1999;12:921.

    • Search Google Scholar
    • Export Citation
  • 23. Platt R, Widel PW, Kesl LD, et al. Comparison of humoral and cellular immune response to a pentavalent modified live virus vaccine in three age groups of calves with maternal antibodies, before and after BVDV type 2 challenge. Vaccine 2009;27:45084519.

    • Search Google Scholar
    • Export Citation
  • 24. Townsend J, Duffus WP, Williams DL, et al. Immune production of interferon by cultured peripheral blood mononuclear cells from calves infected with BHV1 and PI3 viruses. Res Vet Sci 1988;45:198205.

    • Search Google Scholar
    • Export Citation
  • 25. Wentink GH, Rutten VP, van Exsel AC, et al. Failure of an in vitro lymphoproliferative assay specific for bovine herpes virus type 1 to detect immunized or latently infected animals. Vet Q 1990;12:175182.

    • Search Google Scholar
    • Export Citation
  • 26. Miller-Edge M, Splitter G. Patterns of bovine T cell-mediated immune responses to bovine herpesvirus 1. Vet Immunol Immunopathol 1986;13:301319.

    • Search Google Scholar
    • Export Citation
  • 27. Rutten VP, Wentink GH, de Jong WA, et al. Determination of BHV1-specific immune reactivity in naturally infected and vaccinated animals by lymphocyte proliferation assays. Vet Immunol Immunopathol 1990;25:259267.

    • Search Google Scholar
    • Export Citation
  • 28. van Drunen Littel–van den Hurk S, Myers D, Doig PA, et al. Identification of a mutant herpesvirus-1 (BHV-1) in post-arrival outbreaks of IBR in feedlot calves and protection with conventional vaccination. Can J Vet Res 2001;65:8188.

    • Search Google Scholar
    • Export Citation
  • 29. Tough DF, Borrow P, Sprent J. Induction of bystander T cell proliferation by viruses and type I interferon in vivo. Science 1996;272:19471950.

    • Search Google Scholar
    • Export Citation
  • 30. Li Causi E, Parikh SC, Chudley L, et al. Vaccination expands antigen-specific CD4+ memory T cells and mobilizes bystander central memory T cells. PLoS One 2015;10:e0136717.

    • Search Google Scholar
    • Export Citation
  • 31. Elgueta R, de Vries VC, Noelle RJ. The immortality of humoral immunity. Immunol Rev 2010;236:139150.

  • 32. Walz PH, Givens MD, Rodning SP, et al. Evaluation of reproductive protection against bovine viral diarrhea virus and bovine herpesvirus-1 afforded by annual revaccination with modified-live viral or combination modified-live/killed viral vaccines after primary vaccination with modified-live viral vaccine. Vaccine 2017;35:10461054.

    • Search Google Scholar
    • Export Citation
  • 33. Bovine rhinotracheitis vaccine, killed virus. 9 USC §113.216.

  • 34. Roth JA, Carter DP. Comparison of bovine herpesvirus 1 vaccines for rapid induction of immunity. Vet Ther 2000;1:220228.

  • 35. Straub OC, Mawhinney IC. Vaccination to protect calves against infectious bovine rhinotracheitis. Vet Rec 1988;122:407411.

Advertisement

Cell-mediated and humoral immune responses to bovine herpesvirus type 1 and bovine viral diarrhea virus in calves following administration of a killed-virus vaccine and bovine herpesvirus type 1 challenge

View More View Less
  • 1 Boehringer Ingelheim Vetmedica Inc, 2621 North Belt Hwy, St Joseph, MO 64506.
  • | 2 RTI LLC, 801 32nd Ave, Brookings, SD 57006.
  • | 3 RTI LLC, 801 32nd Ave, Brookings, SD 57006.
  • | 4 RTI LLC, 801 32nd Ave, Brookings, SD 57006.
  • | 5 Department of Veterinary and Biomedical Sciences, College of Agriculture and Biological Sciences, South Dakota State University, Brookings, SD 57007.
  • | 6 RTI LLC, 801 32nd Ave, Brookings, SD 57006.
  • | 7 RTI LLC, 801 32nd Ave, Brookings, SD 57006.
  • | 8 RTI LLC, 801 32nd Ave, Brookings, SD 57006.
  • | 9 Department of Veterinary and Biomedical Sciences, College of Agriculture and Biological Sciences, South Dakota State University, Brookings, SD 57007.

Abstract

OBJECTIVE To evaluate cell-mediated and humoral immune responses of calves receiving 2 doses of a dual-adjuvanted vaccine containing inactivated bovine herpesvirus type 1 (BHV1) and bovine viral diarrhea virus types 1 (BVDV1) and 2 (BVDV2) before and after exposure to BHV1.

ANIMALS 24 Holstein steers negative for anti-BHV1 antibodies and proliferative cell-mediated immune responses against BHV1 and BVDV.

PROCEDURES Calves were randomly assigned to 3 groups. The vaccinated group (n = 10) received 2 doses of vaccine on days 0 and 21. Control (n = 10) and seeder (4) groups remained unvaccinated. Calves were commingled during the study except for the 3-day period (days 53 to 55) when seeders were inoculated with BHV1 (1.04 × 107 TCID50, IV) to serve as a source of virus for challenge (days 56 through 84). Rectal temperature and clinical illness scores were monitored, and blood and nasal specimens were obtained for determination of clinicopathologic and immunologic variables.

RESULTS After BHV1 challenge, mean rectal temperature and clinical illness scores were lower for vaccinates than controls. In vaccinates, antibody titers against BHV1 and BVDV2, but not BVDV1, increased after challenge as did extracellular and intracellular interferon-γ expression, indicating a T helper 1 memory response. Additional results of cell marker expression were variable, with no significant increase or decrease associated with treatment.

CONCLUSIONS AND CLINICAL RELEVANCE Calves administered 2 doses of a killed-virus vaccine developed cell-mediated and humoral immune responses to BHV1 and BVDV, which were protective against disease when those calves were subsequently exposed to BHV1.

Abstract

OBJECTIVE To evaluate cell-mediated and humoral immune responses of calves receiving 2 doses of a dual-adjuvanted vaccine containing inactivated bovine herpesvirus type 1 (BHV1) and bovine viral diarrhea virus types 1 (BVDV1) and 2 (BVDV2) before and after exposure to BHV1.

ANIMALS 24 Holstein steers negative for anti-BHV1 antibodies and proliferative cell-mediated immune responses against BHV1 and BVDV.

PROCEDURES Calves were randomly assigned to 3 groups. The vaccinated group (n = 10) received 2 doses of vaccine on days 0 and 21. Control (n = 10) and seeder (4) groups remained unvaccinated. Calves were commingled during the study except for the 3-day period (days 53 to 55) when seeders were inoculated with BHV1 (1.04 × 107 TCID50, IV) to serve as a source of virus for challenge (days 56 through 84). Rectal temperature and clinical illness scores were monitored, and blood and nasal specimens were obtained for determination of clinicopathologic and immunologic variables.

RESULTS After BHV1 challenge, mean rectal temperature and clinical illness scores were lower for vaccinates than controls. In vaccinates, antibody titers against BHV1 and BVDV2, but not BVDV1, increased after challenge as did extracellular and intracellular interferon-γ expression, indicating a T helper 1 memory response. Additional results of cell marker expression were variable, with no significant increase or decrease associated with treatment.

CONCLUSIONS AND CLINICAL RELEVANCE Calves administered 2 doses of a killed-virus vaccine developed cell-mediated and humoral immune responses to BHV1 and BVDV, which were protective against disease when those calves were subsequently exposed to BHV1.

Bovine herpesvirus type 1 is an important pathogen of cattle that causes clinical disease of the respiratory (eg, infectious bovine rhinotracheitis and conjunctivitis) or reproductive (eg, abortion, infectious pustular vulvovaginitis, and balanoposthitis syndrome) tracts as well as encephalitis.1–3 Cattle with BHV1 infections of the upper respiratory tract frequently develop anorexia, fever, tachypnea, cough, and nasal discharge and lesions, which can lead to pathological alterations of the upper respiratory tract and secondary respiratory tract infections caused by other viral or bacterial pathogens associated with the bovine respiratory disease complex (ie, shipping fever).1–3 Bovine herpesvirus type 1 alters the immune system directly by interacting with immune cells and indirectly by altering the release of various cytokines.1,4 Economic losses associated with primary BHV1 infections are largely the result of respiratory and reproductive tract diseases. However, BHV1 can become latent in infected animals and recrudesce subsequent to stress or synthetic corticosteroid treatment, even in the presence of BHV1-neutralizing antibodies. Cattle with recrudescent BHV1 infections can develop clinical disease and serve as a source of infection for naïve animals.2,5

The immune system response to BHV1 infection and vaccination is not completely understood. Interferon γ is a key marker in the development of an acquired immune response because it is produced by only memory (both helper [CD4+] and cytotoxic [CD8+]) and γδ T cells. The production of IFN-γ by T lymphocytes is important for lysis of BHV1-infected cells, but IFN-γ by itself is not cytotoxic to nonimmune PBMCs.6 Proliferation of BHV1-specific PBMCs is inhibited when CD4+ T cells are depleted but is not affected when CD8+ or γδ T cells are depleted.4 Consequently, CD4+ T cells are commonly considered the primary cells involved in a CMI response because CD4+ Th1 cells produce the IFN-γ necessary for activation of macrophages and CD8+ T cells that destroy virus-infected cells.1,7 CD4+ Th2 cells produce IL-4, the cytokine that induces an antibody response.1,7 γδ T cells have a number of functions within the immune system, and their exact role in humoral immunity and CMI remains ambiguous at best. Bovine γδ T cells have regulatory T-cell characteristics (ie, downregulate the immune response following pathogen clearance), serve as costimulatory cells that release IFN-γ or present antigens, and act as a bridge between cell-mediated and humoral immune functions.2,8,9

Vaccination of cattle against BHV1 is effective for disease prevention. After BHV1 establishes an infection, the virus replicates and spreads by intracellular and extracellular routes, thereby avoiding anti-BHV1 antibodies.1,2,10 Because BHV1 can spread intracellularly, an effective immune response must include the development of CMI as well as anti-BHV1 antibodies (ie, humoral immunity). Multiple studies5,6,10,11 have been conducted to evaluate the ability of various BHV1 vaccines to induce CMI. Traditionally, MLV vaccines are considered superior to KV vaccines for the production of both humoral immunity and CMI; however, the potential for attenuated vaccine strains of BHV1 to cause latent infections in vaccinated animals has caused some to question the safety of MLV vaccines.2,12–14 Killed-virus vaccines are generally considered safer than MLV vaccines because they do not contain live BHV1 and therefore cannot establish latent infections that can subsequently recrudesce.2,15,16 The ability of a KV vaccine to induce a CMI response sufficient to protect against disease caused by BHV1 has been scrutinized.2 Results of a recent meta-analysis17 indicate that the level of protection against disease caused by BHV1 conferred by KV vaccines was similar to that of MLV vaccines.

The purpose of the study reported here was to evaluate the CMI response in calves that received a dual-adjuvanted vaccine containing killed strains of BHV1, BVDV types 1 (BVDV1) and 2 (BVDV2), bovine respiratory syncytial virus, and parainfluenza type 3 virus and a bacterin against 5 strains of Leptospira (Leptospira canicola, Leptospira grippotyphosa, Leptospira hardjo, Leptospira incterohemorrhagiae, and Leptospira pomona) and were subsequently challenged with BHV1. The humoral immune response and select clinical variables of those calves were also evaluated.

Materials and Methods

Animals

The study protocol was reviewed and approved by the fully constituted Institutional Animal Care and Use Committee of RTI LLC. Twenty-four approximately 4-month-old Holstein steers were purchased for the study. Prior to purchase, all calves were confirmed to be free of persistent infection with BVDV by means of immunohistochemical analysis of ear notch specimens. Prior to study initiation, all calves were also confirmed to be seronegative for antibodies against BHV1 and negative for proliferative responses against BHV1 and BVDV antigens as described.18

At arrival to the research facility (day -30), all calves were commingled in one 279-m2 pen. The pen had a concrete base and was bedded with straw. It was equipped with an automatic waterer, bale feeders, and approximately 24 m of bunk space as well as a chute and working area. Calves were fed a commercially available dairy calf grower feed that contained monensin (40 g/ton) for prevention and control of coccidiosis caused by Eimeria bovis and Eimeria zuernii at a rate of approximately 2.3 to 3.6 kg/calf/d. Calves had ad libitum access to water and long-stemmed grass hay throughout the study period.

Calves were allowed a 30-day acclimation period prior to study initiation (initial vaccination; day 0). Calves were monitored daily, and any that were observed to be injured or moribund were evaluated by a veterinarian and treated accordingly. Because some of the calves had signs of coccidiosis, all calves received a 5-day treatment course of amprolium on days 13 through 17. Briefly, the automatic waterer was turned off and water was provided in a tank. A 9.6% amprolium solutiona was added to the water in accordance with the label instructions (ie, 473 mL of amprolium solution/378.5 L of water). The pen was also cleaned and bedded with fresh straw.

Study design

Upon arrival at the research facility (day -30), the 24 calves were randomly assigned to 1 of 3 treatment groups by use of software.b Calves in the vaccinated group (vaccinates; n = 10) received 2 doses of a KV respiratory vaccine. Calves in the control group (controls; n = 10) were not vaccinated. Calves in the seeder group (seeders; n = 4) were not vaccinated and were experimentally inoculated with BVH1 so that they could serve as a source of virus exposure for vaccinates and controls during the challenge period. The day vaccinates received the first dose of vaccine was designated day 0, and the challenge period was designated as days 56 through 84. Rectal temperature was recorded for and clinical illness scores were assigned to calves at predetermined times throughout the study period as measures of clinical status. Nasal swab specimens and blood samples were obtained from calves at predetermined times for determination of viral shedding, SN antibody titers against various viruses, and serum concentrations and PBMC expressions of various cell markers and cytokines. All individuals who performed the clinical assessments of calves, as well as those who performed the laboratory assays, remained unaware of (ie, were blinded to) the treatment group assignment of each calf throughout the duration of the study.

Vaccination

All calves in the vaccine group received 5 mL of a dual-adjuvanted vaccinec that contained killed strains of BHV1 (McKercher strain), BVDV1 (Singer strain), BVDV2 (5912 strain), bovine respiratory syncytial virus, and parainfluenza type 3 virus and a bacterin against 5 strains of Leptospira (L canicola, L grippotyphosa, L hardjo, L incterohemorrhagiae, and L pomona) on days 0 and 21. The vaccine was administered SC in the left side of the neck on day 0 and in the right side of the neck on day 21. A new sterile 6-mL syringe and 18-gauge 3/4-inch needle were used to administer the vaccine to each calf. Calves in the control and seeder groups were not vaccinated and were administered sham injections on days 0 and 21. All 24 calves were visually observed for evidence of local or systemic adverse vaccine reactions on a daily basis for at least a week after each vaccination event.

Experimental inoculation of seeders with BHV1

On day 53, the 4 calves in the seeder group were removed from the group pen to a quarantine area, where they were experimentally inoculated with 2 mL (1.04 × 107 TCID50) of virulent BHV1 (Cooper straind), IV. The seeders remained in the quarantine area until day 56 when they were returned to the group pen to serve as a challenge source of BHV1 for controls and vaccinates.

BHV1 challenge and clinical assessment

Seeder calves were returned to the group pen on day 56 and commingled with the controls and vaccinates for the remainder of the observation period. All calves were observed on a daily basis. On days 56 and 59 through 70 (days 0 and 3 through 14 after BHV1 challenge initiation), each calf was assigned a clinical illness score and had its rectal temperature measured with a self-calibrating digital thermometer. The clinical illness score represented the summation of the subjective semiquantitative assessments of respiration (0 = normal, 1 = short and rapid breaths, 2 = mild dyspnea, or 3 = severe dyspnea), nasal discharge (0 = none or slight, 1 = mucoid or blood-tinged, 2 = mucopurulent, or 3 = purulent), nasal lesions (0 = none, 1= erythema, 2 = 1 to 5 plaques, or 3 = < 5 plaques), ocular discharge (0 = none or slight, 1 = moderate, 2 = conjunctivitis, or 3 = purulent), attitude (0 = normal, 1 = mildly depressed, 2 = moderately depressed, or 3 = severely depressed), and cough (0 = absent or 1 = present).

Sample collection

Blood samples were collected by jugular venipuncture from all calves on days -7, 56, 70, and 83. Additional blood samples were collected from vaccinates and controls on days 0, 21, 28, 63, and 66 and from seeders on day 53. From each calf during each acquisition, blood was collected into a blood collection tube without any additives (12.5 mL) and a tube with EDTA (9 mL) as an anticoagulant.

Nasal swab specimens were collected from seeder calves on day 53 (prior to experimental inoculation with BHV1) and from all calves on days 56, 59 through 66, 70, and 84 (days 0, 3 through 10, 14, and 28 after BHV1 challenge initiation). Briefly, during each acquisition, a single swabe was gently inserted into each nare (1 swab/nare), rotated, and then placed in a tube containing nasal transport medium.

SN assays

Standard virus neutralization microtiter assays were used to determine SN antibody titers against BHV1, BVDV1, and BVDV2 for all calves on days 56, 70, and 83 (days 0, 14, and 18 after BHV1 challenge initiation); from vaccinates and controls on days -7, 0 (initial vaccination), 21 (booster vaccination), and 28; and from seeders on day 53 (experimental inoculation with BHV1). Within 24 hours after collection, blood samples collected into blood collection tubes without any additives were centrifuged at 1,500 × g and 4°C for 15 minutes. Serum was then harvested from each sample and stored frozen at −20°C until analysis.

Titers of serum-neutralizing antibodies against each virus were determined in duplicate for each serum sample. The reference strains used in the assays were the Cooper straind for BHV1, cytopathic Singer strainf for BVDV1, and cytopathic A125 strainf for BVDV2. Briefly, serum samples were thawed and heat inactivated at 56°C for 30 minutes. For each serum sample for each SN assay, serial 2-fold dilutions from 1:2 to 1:256 were added to wells of a 96-well microtiter plate. To each well, 50 μL of MEM containing < 500 TCID50 of the designated virus reference strain was added. The plate was incubated at 37°C for 1 hour. Then, the contents of each well were transferred to the corresponding well of a 96-well microtiter tissue culture plateg that contained a monolayer of bovine turbinate cells.h The plates were incubated at 37°C in an atmosphere with 5% CO2 for 3 (BHV1) or 5 (BVDV1 and BVDV2) days, after which each well was evaluated for the presence of virus-induced cytopathic effects.19 For each serum sample, the reciprocal of the highest dilution that prevented cytopathic effect was recorded as the SN antibody titer for the virus in question. Geometric mean antibody titers were calculated by use of log2 titers.

BHV1 isolation

Virus isolation was used to determine whether BHV1 was present in nasal swab specimens. The swabs were removed from the transport medium tubes, then the tubes were vortexed and centrifuged at 600 × g and 4°C for 5 minutes. Each nasal swab fluid sample underwent BHV1 isolation in triplicate. For each replicate, 200 μL of the nasal swab fluid sample was added to a well of a microtiter tissue culture platef that contained a monolayer of BVDV-free bovine turbinate cells.h The plate was incubated at 37°C in an atmosphere with 5% CO2 for 2 to 3 days. Then, the wells were stained with an anti-BHV1 monoclonal antibodyi and read with a fluorescence microscope.j A sample was considered positive if BHV1-specific staining was detected in any of the 3 replicate wells.

Cellular immunologic assays

The BHV1-specific and BVDV-specific IFN-γ concentrations and expressions of various cell surface markers and cytokines were determined for vaccinates and controls on days -30, 0, 21, 28, 56, 63, and 66. Blood samples collected into tubes containing EDTA were processed by means of Ficollk gradient centrifugation as described15 to harvest PBMCs.

Extracellular IFN-α ELISA—Isolated PBMCs from each blood sample were incubated in 8 wells of a 96-well microtiter tissue culture plate.g Two of those 8 wells contained MEM (nonstimulated [negative] control), 2 contained Pokeweed mitogenl (1 μg/well; stimulated [positive] control), 2 contained inactivated BVDV (TGAC strain20; MOI, 0.01), and 2 contained inactivated BHV1 (Cooper straind; MOI, 0.01). The plate was incubated at 37°C in an atmosphere with 5% CO2 for 5 days. The cell culture supernatant was removed from each well, and the IFN-γ concentration in the supernatant was quantified by use of a bovine IFN-γ ELISAm in accordance with the manufacturer's instructions. The light absorbance of each sample was measured at a wavelength of 450 nm with a microplate reader.n A standard curve for each plate was produced by use of online software,o and the IFN-γ concentration in each sample was determined by comparison with the standard curve.

Flow cytometric cellular assays—Expression of cell surface markers (CD44 [memory cells] and WC-1 [γδ T cells]) and intracellular cytokines (IFN-γ and IL-4) by PBMCs was determined by flow cytometry. For each sample, aliquots of PBMC suspensions were added to 12 wells of each of two 96-well microtiter tissue culture plates.g Within each plate, 3 of the 12 wells contained MEM (nonstimulated [negative] control), 3 contained concanavalin Al (stimulated [positive] control; 1 μg/well), 3 contained inactivated BVDV (TGAC strain20; MOI, 0.01), and 3 contained inactivated BHV1 (Cooper straind; MOI, 0.01). Plates were incubated at 37°C in an atmosphere with 5% CO2 for 96 hours. Then, 1 mL of brefeldin Ap (1 μg/mL), a golgi transport inhibitor, was added to each well to prevent the release of intracellular IFN-γ and IL-4. Extracellular tagging of membrane surface molecules was performed on all stimulated PBMC samples by the addition of antibodies against CD4,q CD8,r CD44,s or WC-1t in designated wells. The PBMC samples in all wells were permeabilized with saponinp for determination of intracellular expression of IFN-γ or IL-4. Then, the PBMCs on one plate were treated with a monoclonal antibody against IFN-γ,u whereas those of the other plate were treated with a monoclonal antibody against IL-4.v The stained plates underwent flow cytometry,w and the resulting cell marker data were analyzed with softwarex specific for flow cytometric analysis.

Statistical analyses

Comparisons among treatment groups over time were analyzed by ANOVA for repeated measures with commercially available softwarey as described.21 Briefly, the statistical model consisted of treatment (vaccinated, control, or seeder), sample acquisition time (time), and the interaction between treatment and time. The effect of treatment was analyzed with animal within treatment as the error term, and the effect of time and any interactions were analyzed with the residual as the error term. When a significant effect (P ≤ 0.05) or tendency (P ≤ 0.10) of treatment was detected, pairwise comparisons from the analysis were used to determine level of significance. Values of P ≤ 0.05 were considered significant for all analyses.

Results

Clinical observations

No vaccine-related adverse effects were observed in any of the calves following either vaccination event. One vaccinated animal became debilitated and was euthanized prior to receiving the booster vaccination on day 21. Necropsy results indicated that the calf had severe unresolved coccidiosis. None of the other 23 study calves required additional treatments or medical interventions during the observation period.

All 4 seeders developed clinical signs consistent with BHV1 infection following experimental inoculation with the virus. During the 17 days after BHV1 inoculation, no coughing was noted and the general attitude remained normal for all 4 seeders. During the first 14 days of the challenge period (days 56 through 70), 1 seeder was tachypneic for 1 day, all 4 seeders had an abnormal nasal discharge for at least 1 day, 3 seeders had nasal lesions on days 56 through 66, and all 4 seeders had an abnormal ocular discharge on days 56 through 65 and 70. The mean clinical illness score for the seeders peaked on day 62 (6 days after initiation of the BHV1 challenge period) and decreased to below that at initiation of the challenge period by day 67 (11 days after the BHV1 challenge initiation; Figure 1).

Figure 1—
Figure 1—

Mean clinical illness score (A) and rectal temperature (B) for calves that did (vaccinates; squares and short dashed line; n = 9) and did not (controls; circles and solid line; 10) receive 2 doses of a dual-adjuvanted vaccinec that contained killed strains of BHV1, BVDV types 1 (BVDV1) and 2 (BVDV2), bovine respiratory syncytial virus, and parainfluenza type 1 virus and a bacterin against 5 strains of Leptospira (Leptospira canicola, Leptospira grippotyphosa, Leptospira hardjo, Leptospira incterohemorrhagiae, and Leptospira pomona) on study days 0 and 21, as well as calves that were not vaccinated but were experimentally inoculated with a virulent strain of BHV1 (1.04 × 107 TCID50, IV; seeders; triangles and long dashed line; 4) at various times before and after initiation of a BHV1 challenge period (day 56). All calves were Holstein steers and were approximately 4 months old at study initiation (day 0). All calves were commingled in 1 pen for the duration of the study, except for the 3-day period (days 53 to 55) during which the seeders were quarantined and experimentally inoculated with BHV1. The clinical illness scores represent the total score for each group for each day. *Within a day, mean value differs significantly (P ≤ 0.05) between the control and vaccinated groups. See text for further details regarding the clinical illness scoring system.

Citation: American Journal of Veterinary Research 79, 11; 10.2460/ajvr.79.11.1166

No coughing was noted for any calf in either the control or vaccinated group during the BHV1 challenge period. Among the 10 controls, 1 and 2 calves had an abnormally depressed attitude on days 65 and 66 (9 and 10 days after initiation of the BHV1 challenge period), respectively. Moreover, the mean clinical attitude score for controls was significantly higher (ie, calves were more depressed) than that for vaccinates on both days 65 (P = 0.042) and 66 (P < 0.001). Three controls and 1 vaccinate were assigned an abnormal respiration score on 1 day during the BHV1 challenge period, but the mean respiration score did not differ significantly between the vaccinated and control groups at any time during that period.

Seven of 10 controls and 5 of 9 vaccinates developed an abnormal nasal discharge on at least 1 day during the first 14 days of the BHV1 challenge period. Nasal discharge scores increased from baseline in both groups significantly (P = 0.004) through the challenge period, with the highest levels seen for both groups on days 64 through 70. No significant differences were noted between treatment groups. The nasal discharge score also increased significantly (P = 0.026) from baseline after challenge. The mean nasal discharge score for controls was significantly (P = 0.002) greater than that for vaccinates on day 66. Six of 10 controls and 2 of 9 vaccinates developed nasal lesions.

Eight of 10 controls and 8 of 9 vaccinates had an abnormal ocular discharge on at least 1 day during the first 14 days of the BHV1 challenge period. The high proportion of calves that developed ocular discharge in both the control and vaccinated groups appeared somewhat contradictory when viewed in conjunction with the results for the other clinical variables assessed. Prior to initiation of the study, several of the study calves were determined to have and were treated for conjunctivitis caused by Moraxella bovoculi. Unresolved M bovoculi infection might have interfered with our ability to assess ocular signs associated with the BHV1 challenge.

For both the control and vaccinated groups, the mean clinical illness score increased significantly (P < 0.001) from baseline after challenge (Figure 1). The mean clinical illness score for the control group was consistently greater than that for the vaccinated group between days 61 and 68 (5 through 12 days after initiation of challenge period); however, it did not differ significantly between those 2 groups on any of those days.

The mean rectal temperatures for all 3 groups during the first 14 days of the challenge period were plotted (Figure 1). Among the 10 controls, 7 had an abnormally high rectal temperature (range, 39.4° to 40.7°C) on at least 1 day during the BHV1 challenge period, and 4 of those 7 had an abnormally high rectal temperature on multiple days. Among the 9 vaccinates, only 4 had an abnormally high rectal temperature (range, 39.6° to 39.8°C) on 1 day during the challenge period. Three of the 4 seeders had an abnormally high rectal temperature (range, 39.6° to 40.4°C) on at least 1 day during the challenge period. The mean rectal temperature for the control group was significantly greater than that for the vaccinated group on days 64 (8 days after initiation of the BHV1 challenge period; P = 0.046), 65 (P = 0.003), and 66 (P = 0.029).

SN antibody titers

The mean log2 SN antibody titers against BHV1, BVDV1, and BVDV2 for all 3 groups of calves at various times throughout the observation period were plotted (Figure 2). All 24 calves were seronegative for antibodies against BHV1 on days -7 and 0 (immediately before vaccinates received the first dose of vaccine). On day 21 (day that vaccinates received the booster dose of vaccine), all controls and seeders remained seronegative for antibodies against BHV1, whereas 6 of 9 vaccinates were seropositive for antibodies against BHV1 (mean log2 anti-BHV1 antibody titer, 1.4; range, 1 to 4). On day 28, all vaccinates were seropositive (mean log2 anti-BHV1 antibody titer, 4.5; range, 3 to 6) for antibodies against BHV1. On day 56 (initiation of BHV1 challenge period), all 10 controls remained seronegative for antibodies against BHV1, whereas the 9 vaccinates had a mean log2 anti-BHV1 antibody titer of 3.8 (range, 2 to 5). During the BHV1 challenge period (days 56 through 84), the mean SN anti-BHV1 antibody titer increased dramatically for all 3 groups. On day 70 (14 days after initiation of the challenge period), 5 of 10 controls had seroconverted and had a mean log2 anti-BHV1 antibody titer of 1.9 (range, 1 to 6). By day 83 (27 days after initiation of the challenge period), all 10 controls had seroconverted and had a mean log2 anti-BHV1 antibody titer of 5.2 (range, 3 to 6). The fact that all 10 controls seroconverted to BHV1 during the challenge period indicated that the virus was actively circulating among the study calves.

Figure 2—
Figure 2—

Mean log2 SN antibody titers against BHV1 (A), BVDV1 (B), and BVDV2 (C) at various times before and after vaccination and BHV1 exposure for the calves of Figure 1. See Figure 1 for remainder of key.

Citation: American Journal of Veterinary Research 79, 11; 10.2460/ajvr.79.11.1166

Prior to study initiation, all calves were either seronegative or had low SN antibody titers against BVDV 1 (Figure 2). On day 0, 1 control and 1 vaccinate had anti-BVDV1 antibody titers of 1 and 2, respectively, which were believed to be the result of residual maternally derived antibodies. All 10 controls were seronegative for antibodies against BVDV1 on day 21 and at all subsequent sample acquisition times for the duration of the observation period. The mean log2 anti-BVDV1 antibody titer for the vaccinates was 5.7 (range, 3 to 9) on day 28 (7 days after administration of the booster dose of vaccine), increased to 6.3 on day 56 (initiation of the BHV1 challenge period), remained fairly stable throughout the challenge period, and was 6.2 (range, 4 to 8) on day 83.

On day 0, 2 of the 10 controls and 4 of 9 vaccinates had an SN anti-BVDV2 antibody titer of 1, whereas 2 controls and 2 vaccinates had an SN anti-BVDV2 antibody titer of 2 (Figure 2), most likely owing to residual maternally derived antibodies. All 10 controls were seronegative for anti-BVDV2 antibodies on day 21 and remained seronegative for the remainder of the observation period. For the 9 vaccinates that survived to receive the booster dose of vaccine on day 21, the mean log2 anti-BVDV2 antibody titer was 0.33 (range, < 1 to 5) on day 28, 4.1 (range, 1 to 6) on day 56, 4.3 (range, 3 to 7) on day 70, and 4.4 (range, 1 to 6) on day 83. One vaccinate remained seronegative for anti-BVDV2 antibodies until day 83, at which time it had an anti-BVDV2 antibody titer of 2.

The four seeders were seronegative for antibodies against BHV1, BVDV2, and BVDV2 until after experimental inoculation with BHV1. Following BHV1 inoculation, all 4 seeders seroconverted to BHV1. The mean SN anti-BHV1 antibody titer was 6.8 (range, 4 to 8) on day 70 and 7.2 (range, 4 to 8) on day 83. All 4 seeder calves remained seronegative for antibodies against BVDV1 throughout the observation period. One seeder had an anti-BVDV2 antibody titer of 1 on day 56 but was seronegative again on days 70 and 83.

BHV1 shedding

All 4 seeder calves had nasal swab specimens that tested positive for BHV1 following experimental inoculation with the virus on day 53. In fact, BHV1 was isolated from nasal swab specimens obtained from at least 3 of the 4 seeders on days 56 and 59 through 64 (ie, first week of the BHV1 challenge period) and from 2 and 1 seeders on days 65 and 66, respectively (Table 1). The virus was not isolated from any of the vaccinates or controls until days 61 and 62 (5 and 6 days after initiation of the BHV1 challenge period), respectively. All 10 controls had at least 1 BHV1-positive nasal swab specimen (ie, was shedding the virus) between days 62 and 70, with 6 controls actively shedding the virus for < 4 days. In general, onset of BHV1 shedding was delayed and the duration of BHV1 shedding was shorter in vaccinates relative to controls. The virus was not isolated at any time from 1 of the 9 vaccinates.

Table 1—

Number of calves from which BHV1 was isolated.

Study dayVaccinatesControlsSeeders
56003
59003
60003
61104
62154
63363
64563
65672
66681
70670

All calves were Holstein steers that were approximately 4 months old at study initiation. Vaccinates (n = 9) received 5 mL of a dual-adjuvanted vaccinea that contained killed strains of BHV1, BVDV types 1 (BVDV1) and 2 (BVDV2), bovine respiratory syncytial virus, and parainfluenza type 3 virus and a bacterin against 5 strains of Leptospira (Leptospira canicola, Leptospira grippotyphosa, Leptospira hardjo, Leptospira incterohemorrhagiae, and Leptospira pomona), SC, on study days 0 and 21. Controls (n = 10) and seeders (4) were not vaccinated. All calves were commingled in 1 pen throughout the study, except for a 3-day period (days 53 through 55) during which the seeders were quarantined and experimentally inoculated with 2 mL (1.04 × 107 TCID50) of a virulent strain of BHV1, IV. Seeders were reintroduced into the pen with the rest of the calves on day 56, and nasal swab specimens for BHV1 isolation were obtained from all calves at predetermined times for 2 weeks (days 56 through 70; BHV1 challenge period).

Extracellular BHV1- and BVDV-specific IFN-γ concentrations

For both the control and vaccinated groups, extracellular IFN-γ concentrations measured in the supernatants from PBMCs incubated with MEM (unstimulated [negative] controls) remained low throughout the observation period, and the extracellular IFN-γ concentrations measured in the supernatants from PBMCs incubated with pokeweed mitogen (stimulated [positive] controls) were greater than those for negative controls at all time points. On day 28, the mean extracellular IFN-γ concentration of the positive control samples for the vaccinated group was greater than that for positive control samples for the control group. For both the control and vaccinated groups, the mean extracellular IFN-γ concentration for positive control samples on days 63 and 66 (7 and 10 days after initiation of the BHV1 challenge period) was greater than that on day 56 (data not shown). The mean extracellular BHV1-specific IFN-γ concentration over time for the control and vaccinated groups was plotted (Figure 3). For both groups, the mean extracellular BHV1-specific IFN-γ concentration remained fairly stable prior to initiation of the BHV1 challenge period (day 56), then it increased for both groups. That increase was marked for the vaccinated group (the mean extracellular BHV1-specific IFN-γ concentration went from 981 pg/mL on day 28 to 7,164 pg/mL on day 66) but was less dramatic for the control group (the mean extracellular BHV1-specific IFN-γ concentration went from 339 pg/mL on day 28 to 2,823 pg/mL on day 66). On day 63 (7 days after initiation of the BHV1 challenge period), the mean extracellular BHV1-specific IFN-γ concentration for the vaccinated group (6,617 pg/mL) was significantly (P = 0.032) greater than that for the control group (1,766 pg/mL). For both the vaccinated and control groups, the mean extracellular BHV1-specific IFN-γ concentration on days 56, 63, and 66 was significantly greater than the corresponding mean concentrations on days 0 and 28.

Figure 3—
Figure 3—

Mean extracellular BHV1-specific (A) and BVDV-specific (B) IFN-γ concentrations at various times before and after vaccination and BHV1 exposure for the controls and vaccinates of Figure 1. See Figure 1 for remainder of key.

Citation: American Journal of Veterinary Research 79, 11; 10.2460/ajvr.79.11.1166

The mean extracellular BVDV-specific IFN-γ concentration for the control group mirrored that of the vaccinated group until day 56, after which it continued to increase for the vaccinated group but decreased for the control group (Figure 3). Both group and time significantly affected extracellular BVDV-specific IFN-γ concentration. On day 63 (7 days after initiation of the BHV1 challenge), the mean extracellular BVDV-specific IFN-γ concentration for the vaccinated group (4,584 pg/mL) was significantly (P = 0.043) greater than that for the control group (1,465 pg/mL). As observed with BHV1, for both the vaccinated and control groups, the mean extracellular BVDV-specific IFN-γ concentration on days 56, 63, and 66 was significantly greater than the corresponding mean concentrations on days 0 and 28.

Expression of cell surface markers and intracellular cytokines

The mean percentages of CD4+, CD8+, and WC-1+ (γδ) T cells expressing intracellular IFN-γ (Figure 4), intracellular IL-4 (Figure 5), and CD44 on the cell membrane (Figure 6) following stimulation with BVDV and BHV1 over time for controls and vaccinates were plotted. Despite minor fluctuations in the expression of cell surface CD44 and intracellular IFN-γ and IL-4, results of statistical analyses indicated that expression of those variables was not significantly affected by group, time, or the interaction between group and time. In general, expression pattern fluctuations in CD4+ and CD8+ cells, although minor in magnitude, were more noticeable than those in WC-1+ cells. Also, the percentages of CD4+, CD8+, and WC-1+ cells that expressed CD44 tended to be much higher, compared with the respective percentages of those cells that expressed IFN-γ and IL-4. Expression patterns of IFN-γ, IL-4, and CD44 varied among cell types (CD4+, CD8+, and WC-1+) and on the basis of type of stimulation (BVDV or BHV1) and time relative to vaccination and BHV1 challenge.

Figure 4—
Figure 4—

Mean percentage of CD4+ (A and B), CD8+ (C and D), and WC-1+ (E and F) T cells that expressed IFN-γ following stimulation with inactivated BVDV (A, C, and E) and inactivated BHV1 (B, D, and F) at various times before and after vaccination and BHV1 exposure for the controls (gray bars) and vaccinates (black bars) of Figure 1. Notice that the scale of the y-axis varies among panels.

Citation: American Journal of Veterinary Research 79, 11; 10.2460/ajvr.79.11.1166

Figure 5—
Figure 5—

Mean percentage of CD4+ (A and B), CD8+ (C and D), and WC-1+ (E and F) T cells that expressed IL-4 following stimulation with inactivated BVDV (A, C, and E) and inactivated BHV1 (B, D, and F) at various times before and after vaccination and BHV1 exposure for the controls (gray bars) and vaccinates (black bars) of Figure 1. See Figure 4 for remainder of key.

Citation: American Journal of Veterinary Research 79, 11; 10.2460/ajvr.79.11.1166

Figure 6—
Figure 6—

Mean percentage of CD4+ (A and B), CD8+ (C and D), and WC-1+ (E and F) T cells that expressed CD44 following stimulation with inactivated BVDV (A, C, and E) and inactivated BHV1 (B, D, and F) at various times before and after vaccination and BHV1 exposure for the controls (gray bars) and vaccinates (black bars) of Figure 1. See Figure 4 for remainder of key.

Citation: American Journal of Veterinary Research 79, 11; 10.2460/ajvr.79.11.1166

Discussion

In the present study, calves that received 2 SC doses of a multivalent KV vaccine containing inactivated strains of BHV1, BVDV1, BVDV2, bovine respiratory syncytial virus, and parainfluenza virus type 3 and a bacterin against 5 strains of Leptospira developed humoral and CMI responses to both the BHV1 and BVDV components of the vaccine. Cell-mediated immunity is particularly important for protection against disease caused by BHV-1. Comparison of extracellular IFN-γ assay and flow cytometric results over time between vaccinates and unvaccinated controls suggested that the vaccine induced a Th1 (CMI) response. In fact, the vaccinates had a clear increase in cell-associated BHV1-specific IFN-γ production by CD4+ and CD8+ T cells following vaccination but not after BHV1 challenge. Data regarding IFN-γ production by γδ (WC-1+) T cells were inconsistent, likely owing to variation in the baseline (initial) number of γδ T cells between controls and vaccinates, which masked vaccine-induced effects. Mean intracellular expression of IL-4 did not differ significantly between vaccinates and controls at any time during the observation period. Collectively, those results suggested that the Th1 response but not the Th2 response differed between vaccinates and controls.

The CD44 expression data indicated that controls generally had fewer activated or memory cells than vaccinates throughout the observation period, although the magnitude of that difference was not significant. Also, the changes in CD44+ expression for the control group mirrored those of the vaccinated group throughout the observation period. Thus, the importance of that finding in terms of immune system function remains to be defined.

During the BHV1 challenge period, the mean proportion of IFN-γ–positive CD8+ cells remained fairly stable for the vaccinated group, whereas it increased for the control group. This was not unexpected because the BHV1 challenge period was the first time that controls were exposed to the virus. It has been theorized that CMI, in conjunction with humoral and other nonspecific immune mechanisms, is the primary mechanism by which cattle recover from an initial BHV1 infection.1 Results of another study22 indicate that the proportion of CD25-positive CD8+ cells decreases in unvaccinated calves with active BHV1 infections. However, in yet another study11 PBMCs obtained from calves vaccinated with an MLV vaccine had a significant increase in CD25 (interleukin-2 receptor α chain) expression by CD4+, CD8+, and γδ T cells following in vitro stimulation with live BHV1. Calves with maternal antibodies that were vaccinated with a pentavalent MLV vaccine and then challenged with active BVDV2 had a significant increase in mean expression of CD25, IFN-γ, and IL-4 by CD4+, CD8+, and γδ T cells following stimulation with BVDV1 and BVDV2, but as in the present study, few significant differences in the expression of those cytokines were observed following stimulation with active BHV1.23 Cattle vaccinated with an MLV vaccine against BHV1 have an increase in expression of BHV1-specific CD25 by CD4+ T cells at 2, 3, 4, 6, and 7 weeks after vaccination and by γδ T cells at all times after vaccination22; however, BHV1-specific CD25 expression by CD8+ T cells did not change in those cattle following vaccination. Cattle vaccinated with an adjuvanted KV vaccine against BVDV had an increase in the percentage of CD8+, γδ, and CD4+ T cells that expressed BVDV-specific IFN-γ throughout the observation period; at 7, 14, and 31 days after vaccination; and at 42 days after vaccination, respectively, compared with unvaccinated control cattle.15 In the present study, calves vaccinated with an adjuvanted KV vaccine had a greater percentage of CD4+ and CD8+ cells that expressed BVDV-specific IFN-γ on days 0 (administration of first dose of vaccine), 21 (administration of second dose of vaccine), and 28 than did unvaccinated controls.

In the present study, the percentage of CD4+, CD8+, and γδ T cells expressing BHV1-specific IFN-γ for vaccinates increased from baseline (day -7) following vaccination prior to the BHV1 challenge period. That finding suggested that the vaccine induced a CMI response against BHV1 by 28 days after administration of the first dose (and 7 days after administration of the booster dose). During the BHV1 challenge period, the extracellular BHV1-specific IFN-γ concentration for vaccinates increased 7-fold from that on day 28, which was indicative of a robust CMI response. Interferon-γ is produced by only memory (both helper and cytotoxic) and γδ T cells; therefore, it is an important indicator of an acquired immune response. Production of IFN-γ is vital for CMI owing to its direct antiviral activity and its prominence as an activator of the cytotoxic response.7 Several reports4,13,24–28 describe the effects of BHV1 stimulation on IFN-γ production by immune and nonimmune PBMCs. Spontaneous in vitro production of BHV1-specific IFN-γ was observed in unstimulated cultures of PBMCs obtained from calves following experimental inoculation with 2 doses of live BHV1 4 to 7 weeks apart.24 Peripheral blood mononuclear cells obtained from colostrum-deprived neonatal calves that received either a low or high dose of an MLV vaccine containing a glycoprotein E–negative strain of BHV1, intranasally, were stimulated to produce BHV1-specific IFN-γ on at least 1 occasion following vaccination when not exposed to dexamethasone, but could not be stimulated to produce IFN-γ when exposed to dexamethasone.13 In vaccinated cattle, IFN-γ production varies and BHV1-specific proliferative responses are not necessarily correlated with protection from disease when challenged with the virus.4,25 Differences in the proliferative immune response among cattle could result from interindividual variation in immune status or the ability of PBMCs to discriminate among serologically cross-reactive BHV1 isolates.26 However, in 1 study,27 lymphocytes obtained from cattle vaccinated against 3 strains of BHV1 had similar proliferative responses, as determined by 3H thymidine uptake, when stimulated with an alternate strain of live BHV1 virus in vitro.27 For cattle of another study,28 BHV1-specific proliferative responses increased following vaccination, but when vaccinated and unvaccinated control cattle underwent a BHV1 challenge, the proliferative immune response was stronger for a mutant strain of the virus than for a known strain in both groups. Genetic differences among cattle can result in the induction of different types of immune responses. The immune status of cattle, and hence its variability, can be affected by the type of vaccine (MLV or KV) used, type of adjuvants in the vaccine, and number of vaccinations administered. All those factors have the potential to affect an animal's ability to recognize and mount an immune response to both conserved and variable epitopes of the vaccinal virus strain.

During the BHV1-challenge period of the present study, vaccinates had a vigorous BVDV-specific response (as evidenced by the increase in the extracellular BVDV-specific IFN-γ concentration) in conjunction with the strong BHV1-specific response, which was likely a bystander effect. A bystander, or nearest-neighbor, effect occurs in response to some viral infections and is characterized by a marked transient increase in total T-cell numbers and activation of specific T cells by heterologous antigens.29 The presence of a bystander effect suggests that proliferative responses of T cells are not entirely antigen specific. In mice, the bystander effect is driven by cytokines, with IFN-α and IFN-β (IFN-I) having primary roles as inducers of bystander cells.29 In humans that received a tetanus toxoid vaccine, the presence of a bystander effect was identified on the basis of an increase in the proliferative responses of T helper memory cells to the tetanus toxoid antigens as well as 2 unrelated and non-cross-reactive common recall antigens (purified proteins derived from tuberculin and Candida albicans).30,31 In those studies,30,31 an increase in IFN-γ production by tetanus toxoid-specific T cells was also observed following stimulation with purified proteins derived from tuberculin and C albicans. In mice, a recall response to tetanus toxoid also induces proliferation of CD4+ T cells previously activated by a specific unrelated antigen.29 Elucidation of the significance of a bystander effect has been ongoing for several years. Interferon-I is believed to be involved in sustaining immunological memory as well as inducing a bystander response.29 It is possible that the 2 are interconnected and that the bystander effect may help sustain long-term memory responses, although additional research is necessary to understand the underlying mechanisms.

The serologic response following vaccination for the vaccinates of the present study was similar to that reported in cattle of other studies.11,12,16,23,26 Antibodies are necessary for neutralization of extracellular virus.2 Prior to initiation of the present study, all calves were seronegative for anti-BHV1 antibodies and some had very low antibody titers against BVDV1 or BVDV2, which were likely residual maternally derived antibodies. Following vaccination, all vaccinates developed measurable SN antibody titers against BHV1, BVDV1, and BVDV2, whereas the controls remained seronegative for antibodies against all 3 viruses. After initiation of the BHV1 challenge period, all the controls seroconverted to BHV1 but remained seronegative for antibodies against BVDV1 and BVDV2 for the duration of the study. The mean anti-BHV1 antibody titer for vaccinates continued to increase during the BHV1 challenge period as did the anti-BVDV2 antibody titers, albeit to a lesser extent. That finding may indicate that the bystander effect evidenced by an increase in IFN-γ production by BVDV-stimulated PBMCs spilled over to a Th2-mediated response against BVDV2; however, a similar response was not detected against BVDV1. A bystander humoral effect has been described in humans who developed antibodies against Clostridium tetani as well as unrelated antigens after receiving a tetanus toxoid.31 In another study,32 an increase in SN antibody titers against BVDV1 and BVDV2 following experimental infection with BHV1 was observed in vaccinated cattle and unvaccinated cattle exposed to BVDV. To our knowledge, the present study was the first to describe a bystander effect in regard to an anti-BVDV antibody response in vaccinated calves following a BHV1 challenge. The fact that a possible serologic bystander effect was observed for BVDV2, but not BVDV1, was interesting and warrants further investigation.

Results of the present study indicated that the KV vaccine used to vaccinate the calves of the vaccinated group was effective in eliciting both a CMI and humoral immune response and protecting against disease when those calves were exposed to BHV1. Virus isolation results indicated that the seeder calves successfully spread BHV1 to both vaccinates and controls. During the first 2 weeks of the BHV1 challenge period, vaccinates had a lower mean rectal temperature and lower clinical illness score than did the controls. Body temperature (and by proxy, rectal temperature) is considered a primary indicator for vaccine efficacy when BHV1 challenge models are used.33 The vaccinates of the present study received 2 doses of a KV vaccine, and the mitigation of disease severity following subsequent exposure of those calves to BHV1 was comparable to that induced by MLV and KV vaccines in cattle of other studies.3,11,13,28,32,34,35 Calves vaccinated with a KV vaccine at 8 and 12 weeks or at 4, 8, and 12 weeks old developed fewer clinicals signs of disease and weighed significantly more at the end of the observation period than unvaccinated calves.35 Results of a recent meta-analysis17 suggest that vaccination of cattle against BHV1 with a commercially available MLV or KV vaccine decreases the risk of abortion following BHV1 exposure by approximately 60%.

Results of the present study indicated that calves that received 2 SC doses of a multivalent KV vaccine containing inactivated strains of BHV1, BVDV1, BVDV2, bovine respiratory syncytial virus, and parainfluenza virus type 3 and a bacterin against 5 strains of Leptospira developed humoral and CMI responses to both the BHV1 and BVDV components of the vaccine, which conferred protection against disease when those calves were subsequently exposed to BHV1. During the BHV1 challenge period the concentrations of both BHV1-specific and BVDV-specific IFN-γ (indicators of CMI) increased for both vaccinates and controls, but the magnitude of those increases for vaccinates was several fold greater than that for controls. Also during the BHV1 challenge period, the controls developed SN antibody titers against BHV1 as expected but remained seronegative for antibodies against BVDV1 and BVDV2, whereas the vaccinates had an increase in SN antibody titers against both BHV1 and BVDV2, but not BVDV1. Collectively, those results suggested that the vaccine induced effective CMI and humoral responses against BHV1 when vaccinates were exposed to that virus as well as a bystander CMI response against BVDV1 and BVDV2 and a bystander humoral response against BVDV2 but not BVDV1. The latter finding was interesting, and further research into the mechanisms involved in the development of bystander effects is warranted.

Acknowledgments

The study was performed at the RTI LLC clinical and laboratory facilities located in South Dakota.

The authors thank Dr. Alan Young of South Dakota State University for proliferation assay and flow cytometry analysis and Dr. George Perry of South Dakota State University for statistical analysis.

ABBREVIATIONS

BHV1

Bovine herpesvirus type 1

BVDV

Bovine viral diarrhea virus

CMI

Cell-mediated immunity

IFN

Interferon

IL-4

Interleukin-4

KV

Killed virus

MEM

Minimum essential medium

MLV

Modified-live virus

MOI

Multiplicity of infection

PBMC

Peripheral blood mononuclear cell

SN

Serum neutralization

Th1

T helper 1

Th2

T helper 2

WC-1

γδ T-cell glycoprotein marker

Footnotes

a.

Corid, 9.6% oral solution, Merial Ltd, Duluth, Ga.

b.

RAND function, Microsoft Excel, Microsoft Corp, Edina, Minn.

c.

Triangle 10, Boehringer Ingelheim Vetmedica Inc, Duluth, Ga.

d.

Cooper strain (lot No. 00-22), Center for Veterinary Biologics, Ames, Iowa.

e.

FLOQSwab, Copan Flock Technologies Srl, Brescia, Italy.

f.

National Veterinary Services Laboratories, Ames, Iowa.

g.

Grenier Bio-One, Frickenhausen, Germany.

h.

American Type Culture Collection, Manassas, Va.

i.

Anti-BHV1 monoclonal antibody (lot No. P140-1222-001), South Dakota State University, Brookings, SD.

j.

CK40° fluorescence microscope, Olympus America Inc, Melville, NY.

k.

Ficoll-Paque, GE Healthcare, Amersham, England.

l.

Sigma-Aldrich Corp, St Louis, Mo.

m.

AbD Serotec, Kidlington, England.

n.

Tecan Sunrise, Männedorf, Switzerland.

o.

MyAssays.com. Accessed Oct 1, 2018.

p.

BD Cytofix/Cytoperm Fixation/Permeabilization Kit, BD Biosciences, San Jose, Calif.

q.

GC50A, Washington State University, Pullman, Wash.

r.

ST8, Washington State University, Pullman, Wash.

s.

BAG40A, Washington State University, Pullman, Wash.

t.

BAG25A, Washington State University, Pullman, Wash.

u.

MCA1783PE, AbD Serotec, Kidlington, England.

v.

MCA1820PE, AbD Serotec, Kidlington, England.

w.

Accuri C6 Flow Cytometer, BD Biosciences, San Jose, Calif.

x.

Cell Sampler Software, BD Biosciences, San Jose, Calif.

y.

SAS, version 9.4, SAS Institute Inc, Cary, NC.

References

  • 1. Babiuk LA, van Drunen Littel–van den Hurk S, Tikoo SK. Immunology of bovine herpesvirus 1 infection. Vet Microbiol 1996;53:3142.

  • 2. van Drunen Littel–van den Hurk S. Cell-mediated immune responses induced by BHV-1: rational vaccine design. Expert Rev Vaccines 2007;6:369380.

    • Search Google Scholar
    • Export Citation
  • 3. van Drunen Littel–van den Hurk S, Tikoo SK, van den Hurk JV, et al. Protective immunity in cattle following vaccination with conventional and marker bovine herpesvirus-1 (BHV1) vaccines. Vaccine 1997;15:3644.

    • Search Google Scholar
    • Export Citation
  • 4. Denis M, Kaashoek MJ, van Oirschot JT, et al. Quantitative assessment of the specific CD4+ T lymphocyte proliferative response in bovine herpesvirus 1 immune cattle. Vet Immunol Immunopathol 1994;42:275286.

    • Search Google Scholar
    • Export Citation
  • 5. Davies DH, Carmichael LE. Role of cell-mediated immunity in the recovery of cattle from primary and recurrent infections with infectious bovine rhinotracheitis virus. Infect Immun 1973;8:510518.

    • Search Google Scholar
    • Export Citation
  • 6. Campos M, Ohmann HB, Hutchings D, et al. Role of interferon-gamma in inducing cytotoxicity of peripheral blood mononuclear leukocytes to bovine herpesvirus type 1 (BHV1)–infected cells. Cell Immunol 1989;120:259269.

    • Search Google Scholar
    • Export Citation
  • 7. Tikoo SK, Campos M, Popowych YI, et al. Lymphocyte proliferative responses to recombinant bovine herpes virus type 1 (BHV-1) glycoprotein gD (gIV) in immune cattle: identification of a T cell epitope. Viral Immunol 1995;8:1925.

    • Search Google Scholar
    • Export Citation
  • 8. Guzman E, Price S, Poulsom H, et al. Bovine γδ T cells: cells with multiple functions and important roles in immunity. Vet Immunol Immunopathol 2012;148:161167.

    • Search Google Scholar
    • Export Citation
  • 9. Hoek A, Rutten VP, Kool J, et al. Subpopulations of bovine WC1+ γδ T cells rather than CD4+CD25highFoxp3+ T cells act as immune regulatory cells ex vivo. Vet Res 2009;40:6.

    • Search Google Scholar
    • Export Citation
  • 10. Rouse BT, Babiuk LA. Host defense mechanisms against infectious bovine rhinotracheitis virus; in vitro stimulation of sensitized lymphocytes by virus antigen. Infect Immun 1974;10:681687.

    • Search Google Scholar
    • Export Citation
  • 11. Endsley JJ, Quade MJ, Terharr B, et al. BHV-1–specific CD4+, CD8+, and γδ T cells in calves vaccinated with one dose of a modified live BHV-1 vaccine. Viral Immunol 2002;15:385393.

    • Search Google Scholar
    • Export Citation
  • 12. 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.

    • Search Google Scholar
    • Export Citation
  • 13. Lemaire M, Schynts F, Meyer G, et al. Latency and reactivation of a glycoprotein E negative bovine herpesvirus type 1 vaccine: influence of virus load and effect of specific maternal antibodies. Vaccine 2001;19:47954804.

    • Search Google Scholar
    • Export Citation
  • 14. Chase CCL, Fulton RW, O'Toole D, et al. Bovine herpesvirus 1 modified live virus vaccines for cattle reproduction: balancing protection with undesired effects. Vet Microbiol 2017;206:6977.

    • Search Google Scholar
    • Export Citation
  • 15. Stevens ET, Zimmerman AD, Buterbaugh RE, et al. The induction of a cell-mediated immune response to bovine viral diarrhea virus with an adjuvanted inactivated vaccine. Vet Ther 2009;10:E1E8.

    • Search Google Scholar
    • Export Citation
  • 16. Zimmerman AD, Buterbaugh RE, Herbert JM, et al. Efficacy of bovine herpesvirus-1 inactivated vaccine against abortion and stillbirth in pregnant heifers. J Am Vet Med Assoc 2007;231:13861389.

    • Search Google Scholar
    • Export Citation
  • 17. Newcomer BW, Cofield LG, Walz PH, et al. Prevention of abortion in cattle following vaccination against bovine herpesvirus 1: a meta-analysis. Prev Vet Med 2017;138:18.

    • Search Google Scholar
    • Export Citation
  • 18. Woolums AR, Siger L, Johnson S, et al. Rapid onset of protection following vaccination of calves with multivalent vaccines containing modified-live or modified-live and killed BHV-1 is associated with virus specific interferon gamma production. Vaccine 2003;21:11581164.

    • Search Google Scholar
    • Export Citation
  • 19. Manual of standards for diagnostic tests and vaccines for terrestrial animals. Paris: Office International des Epizooties, 2015;1415.

    • Search Google Scholar
    • Export Citation
  • 20. Bissey LL, Williams AK, Bolin S, et al. Comparison of cytopathic and noncytopathic isolates of bovine viral diarrhea virus by oligonucleotide fingerprinting. J Vet Diagn Invest 1991;3:1621.

    • Search Google Scholar
    • Export Citation
  • 21. Littell RC, Henry PR, Ammerman CB. Statistical analysis of repeated measures data using SAS procedures. J Anim Sci 1998;76:12161231.

  • 22. Quade MJ, Roth JA. Antigen-specific in vitro activation of T-lymphocyte subsets of cattle immunized with a modified live bovine herpesvirus 1 vaccine. Viral Immunol 1999;12:921.

    • Search Google Scholar
    • Export Citation
  • 23. Platt R, Widel PW, Kesl LD, et al. Comparison of humoral and cellular immune response to a pentavalent modified live virus vaccine in three age groups of calves with maternal antibodies, before and after BVDV type 2 challenge. Vaccine 2009;27:45084519.

    • Search Google Scholar
    • Export Citation
  • 24. Townsend J, Duffus WP, Williams DL, et al. Immune production of interferon by cultured peripheral blood mononuclear cells from calves infected with BHV1 and PI3 viruses. Res Vet Sci 1988;45:198205.

    • Search Google Scholar
    • Export Citation
  • 25. Wentink GH, Rutten VP, van Exsel AC, et al. Failure of an in vitro lymphoproliferative assay specific for bovine herpes virus type 1 to detect immunized or latently infected animals. Vet Q 1990;12:175182.

    • Search Google Scholar
    • Export Citation
  • 26. Miller-Edge M, Splitter G. Patterns of bovine T cell-mediated immune responses to bovine herpesvirus 1. Vet Immunol Immunopathol 1986;13:301319.

    • Search Google Scholar
    • Export Citation
  • 27. Rutten VP, Wentink GH, de Jong WA, et al. Determination of BHV1-specific immune reactivity in naturally infected and vaccinated animals by lymphocyte proliferation assays. Vet Immunol Immunopathol 1990;25:259267.

    • Search Google Scholar
    • Export Citation
  • 28. van Drunen Littel–van den Hurk S, Myers D, Doig PA, et al. Identification of a mutant herpesvirus-1 (BHV-1) in post-arrival outbreaks of IBR in feedlot calves and protection with conventional vaccination. Can J Vet Res 2001;65:8188.

    • Search Google Scholar
    • Export Citation
  • 29. Tough DF, Borrow P, Sprent J. Induction of bystander T cell proliferation by viruses and type I interferon in vivo. Science 1996;272:19471950.

    • Search Google Scholar
    • Export Citation
  • 30. Li Causi E, Parikh SC, Chudley L, et al. Vaccination expands antigen-specific CD4+ memory T cells and mobilizes bystander central memory T cells. PLoS One 2015;10:e0136717.

    • Search Google Scholar
    • Export Citation
  • 31. Elgueta R, de Vries VC, Noelle RJ. The immortality of humoral immunity. Immunol Rev 2010;236:139150.

  • 32. Walz PH, Givens MD, Rodning SP, et al. Evaluation of reproductive protection against bovine viral diarrhea virus and bovine herpesvirus-1 afforded by annual revaccination with modified-live viral or combination modified-live/killed viral vaccines after primary vaccination with modified-live viral vaccine. Vaccine 2017;35:10461054.

    • Search Google Scholar
    • Export Citation
  • 33. Bovine rhinotracheitis vaccine, killed virus. 9 USC §113.216.

  • 34. Roth JA, Carter DP. Comparison of bovine herpesvirus 1 vaccines for rapid induction of immunity. Vet Ther 2000;1:220228.

  • 35. Straub OC, Mawhinney IC. Vaccination to protect calves against infectious bovine rhinotracheitis. Vet Rec 1988;122:407411.

Contributor Notes

Dr. Van Anne's present address is Van Anne Veterinary Service, Gering, NE 69341.

Address correspondence to Dr. Chase (christopher.chase@sdstate.edu).