Advertisement
Browse Subject Areas
?

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Priming Cross-Protective Bovine Viral Diarrhea Virus-Specific Immunity Using Live-Vectored Mosaic Antigens

  • Shehnaz Lokhandwala,

    Affiliation Department of Veterinary Pathobiology, Texas A&M University, College Station, Texas, United States of America

    ORCID http://orcid.org/0000-0002-0873-8611

  • Xin Fang,

    Affiliation Department of Animal Science, Texas A&M University, College Station, Texas, United States of America

  • Suryakant D. Waghela,

    Affiliation Department of Veterinary Pathobiology, Texas A&M University, College Station, Texas, United States of America

  • Jocelyn Bray,

    Affiliation Department of Veterinary Pathobiology, Texas A&M University, College Station, Texas, United States of America

  • Leo M. Njongmeta,

    Current address: Environmental Health & Safety Office, University of Illinois, Chicago, Illinois, United States of America

    Affiliation Department of Veterinary Pathobiology, Texas A&M University, College Station, Texas, United States of America

  • Andy Herring,

    Affiliation Department of Animal Science, Texas A&M University, College Station, Texas, United States of America

  • Karim W. Abdelsalam,

    Affiliation Department of Veterinary and Biomedical Sciences, South Dakota State University, Brookings, South Dakota, United States of America

  • Christopher Chase,

    Affiliation Department of Veterinary and Biomedical Sciences, South Dakota State University, Brookings, South Dakota, United States of America

  • Waithaka Mwangi

    wmwangi@cvm.tamu.edu

    Affiliation Department of Veterinary Pathobiology, Texas A&M University, College Station, Texas, United States of America

Priming Cross-Protective Bovine Viral Diarrhea Virus-Specific Immunity Using Live-Vectored Mosaic Antigens

  • Shehnaz Lokhandwala, 
  • Xin Fang, 
  • Suryakant D. Waghela, 
  • Jocelyn Bray, 
  • Leo M. Njongmeta, 
  • Andy Herring, 
  • Karim W. Abdelsalam, 
  • Christopher Chase, 
  • Waithaka Mwangi
PLOS
x

Abstract

Bovine viral diarrhea virus (BVDV) plays a key role in bovine respiratory disease complex, which can lead to pneumonia, diarrhea and death of calves. Current vaccines are not very effective due, in part, to immunosuppressive traits and failure to induce broad protection. There are diverse BVDV strains and thus, current vaccines contain representative genotype 1 and 2 viruses (BVDV-1 & 2) to broaden coverage. BVDV modified live virus (MLV) vaccines are superior to killed virus vaccines, but they are susceptible to neutralization and complement-mediated destruction triggered by passively acquired antibodies, thus limiting their efficacy. We generated three novel mosaic polypeptide chimeras, designated NproE2123; NS231; and NS232, which incorporate protective determinants that are highly conserved among BVDV-1a, 1b, and BVDV-2 genotypes. In addition, strain-specific protective antigens from disparate BVDV strains were included to broaden coverage. We confirmed that adenovirus constructs expressing these antigens were strongly recognized by monoclonal antibodies, polyclonal sera, and IFN-γ-secreting T cells generated against diverse BVDV strains. In a proof-of-concept efficacy study, the multi-antigen proto-type vaccine induced higher, but not significantly different, IFN-γ spot forming cells and T-cell proliferation compared to a commercial MLV vaccine. In regards to the humoral response, the prototype vaccine induced higher BVDV-1 specific neutralizing antibody titers, whereas the MLV vaccine induced higher BVDV-2 specific neutralizing antibody titers. Following BVDV type 2a (1373) challenge, calves immunized with the proto-type or the MLV vaccine had lower clinical scores compared to naïve controls. These results support the hypothesis that a broadly protective subunit vaccine can be generated using mosaic polypeptides that incorporate rationally selected and validated protective determinants from diverse BVDV strains. Furthermore, regarding biosafety of using a live vector in cattle, we showed that recombinant human adenovirus-5 was cleared within one week following intradermal inoculation.

Introduction

Bovine viral diarrhea virus (BVDV), an infectious pathogen that is prevalent in cattle herds globally, is a key agent responsible for causing Bovine Respiratory Disease Complex (BRDC) [1]. Infection with BVDV can cause severe diarrhea, respiratory disease, immunosuppression, abortion, congenital malformations, and birth of persistently infected (PI) calves, which play a major role in virus transmission in herds [2]. Immunosuppression caused by acute infection of unprotected calves allows secondary infections to establish and cause pneumonia or enteritis [3]. The secondary infections are responsible for high rates of morbidity and mortality, and it is estimated that the U.S. livestock industry loses >$1billion annually due to BRDC [4, 5].

This virus is classified as a member of the genus Pestivirus within the family Flaviviridae [6]. Two BVDV genotypes (type 1 and 2) are recognized according to serological and genetic relatedness [7]. The BVDV isolates circulating in the world are heterogeneous: BVDV genotype 1 (BVDV-1) is subdivided into a minimum of 12 sub-genotypes (BVDV1a, b, c.…l), whereas BVDV genotype 2 (BVDV-2) is classified into 4 subtypes, 2a-2d [8, 9]. The BVDV can also be divided into cytopathic and non-cytopathic biotypes (cpBVDV and ncpBVDV, respectively), based on their lytic effects on infected cells. The BVDV isolates cause a wide range of disease manifestations, which include sub-clinical and persistent infections, fetal infections, and host immunosuppression [10]. Infected cattle begin to shed the virus into the environment for about ten continuous days starting as early as four days after subclinical infection, whereas PI animals shed the virus for their entire lifetime [11, 12]. The prevalence of PI animals in selected herds in the United States is estimated at 1.7% of the cattle population, and these animals are considered to be the primary source of infection of susceptible animals [13].

BVDV infection in cattle induces high titers of neutralizing antibodies that prevent reinfections especially with the same genotype/sub-genotype [14, 15]. Some studies have demonstrated prevention of clinical signs, but not viral shedding, in cattle upon challenge with BVDV-2 following immunization with BVDV-1 [16, 17]. Failure of vaccination has been attributed to infection with variant genotype(s) as well as development of antigenically distinct viruses in exposed animals [18, 19]. Individual PI cattle may also be a source of genetic variants that amplify following infection of susceptible cattle [20, 21]. However, in the absence of neutralizing antibodies, mutations occur faster and more frequently in BVDV following infection of pregnant animals [22]. Many of the virus genome mutations result in amino acid changes in E2 glycoprotein, a key target of the neutralizing antibodies [21, 23]. The E2 glycoprotein is highly immunogenic and at least nine epitopes have been mapped within three antigenic domains [2428]. One of these antigenic determinants is immunodominant in BVDV-1 and there are three in BVDV-2 that induce neutralizing antibodies in animals [25]. However, it is also reported that viremia can occur despite the presence of neutralizing antibodies in infected animals, and some animals can be protected against BVDV infection in the absence of E2-specific neutralizing antibodies, suggesting a role for neutralizing epitopes from other antigens and/or T cells in protection [29, 30]. Clearance of BVDV infections has also been associated with strain-specific MHC-restricted CD4+ and CD8+ T-cell responses [15, 31]. Cell mediated response to infection is initially provided by E2 and NS2-3 antigen-specific helper CD4+ T-cells [3234].

Despite availability of vaccines, BVDV prevalence has not markedly reduced due, in part, to failure of the vaccines to confer broad protection [35, 36]. Currently, both killed and modified live virus (MLV) vaccines are commercially available [37]. The killed vaccine elicits primarily a humoral response with minimal cell mediated response, whereas MLV vaccines are better at inducing CD4+ and CD8+ T-cells responses in addition to antibody responses [38]. Since the presence of BVDV-specific maternal antibodies interferes with efficacy of BVDV vaccines, especially MLV, immunization is usually delayed until most of the maternal BVDV antibodies have waned [19, 39]. However, BVDV-specific antibodies in each animal decline at different rates and thus, antibody titers in some calves fall below protective levels much earlier than expected, and in the presence of PI calves in the herd, there is a high risk of infection [40]. MLV vaccines are currently the most efficacious, but genotype-specific vaccines are not effective at conferring cross-protection and thus, protection against BVDV-1 and 2 requires a vaccine formulation that contains a representative of genotype type 1 and 2 viruses. The MLV vaccines are not considered to be safe since the attenuated virus can revert to wild type virus, cause in-utero infections and mucosal disease, carry the risk of vaccine contamination with adventitious viruses, and are immunosuppressive [41, 42]. Furthermore, MLV strains may cause ovarian lesions leading to infertility in cows [43]. Both killed and MLV vaccine virus are traditionally grown in MDBK cells and recent findings show that calves fed colostrum from some dams vaccinated with killed BVDV vaccine formulated with adjuvant have a high incidence of a syndrome characterized by spontaneous bleeding, severe anemia with heavy bone marrow damage. There is evidence to show that the damage is due to maternal alloantibodies induced by the vaccines against bovine cell antigens, including MHC-I molecules, and the syndrome has been named bovine neonatal pancytopenia [4446].

Given the limitations of the current vaccines, there is a need to develop improved vaccines for safe, robust, and broad protection against diverse BVDV genotypes. Empirical selection and validation of protective immune targets that are conserved among diverse BVDV strains can be used to generate novel mosaic antigens for development of a contemporary vaccine. Similar strategies have been used to develop broadly protective vaccines to overcome a wide Influenza and HIV-1 genetic diversity [4749]. The BVDV envelop (E2) and non-structural (NS2-3) antigens are immunodominant, and neutralizing antibody as well as T-cell responses directed against these antigens can confer protection [50, 51]. Importantly, evaluation of BVDV-specific immune responses following resolution of acute infection has revealed that the E2, NS2-3, and N-terminal protease fragment (Npro) antigens contain CD4+ T-cell epitopes [31]. In addition, MHC DR-restricted T-cell epitopes have been identified from conserved regions of E2 and NS2-3 [31, 34, 5153].

In this study, we generated a prototype vaccine composed of recombinant adenoviruses expressing three novel mosaic polypeptide chimeras, designated NproE2123; NS231; and NS232. These antigens incorporated neutralizing epitopes, defined and predicted IFN-γ-inducing CD4+ T-cell as well as cytotoxic T lymphocyte determinants that are highly conserved among BVDV-1a, b, and BVDV-2 genotypes [24, 25, 31, 51, 52]. In addition, strain-specific protective antigens from disparate BVDV strains whose genome sequences are available were included to broaden coverage. We compared the immunogenicity and protective efficacy of this prototype adenovirus-vectored vaccine to a commercial MLV vaccine in calves.

Adenovirus-vectored subunit vaccines are undergoing clinical trials in readiness for deployment [54, 55]; there is concern that persistence of the construct in host tissues may increase chances of generating replication-competent progenies if recombination with closely related viruses occurs. Thus, we set out to determine replication-incompetent recombinant human adenovirus-5 persistence at the skin injection site, the draining lymph node, and the spleen of calves following intradermal inoculation.

Materials and Methods

Design of genes encoding novel mosaic BVDV antigens

Published reports on protective BVDV antibody and T-cell epitopes, sequenced genomes, and bioinformatics tools were used to design novel mosaic polypeptides, which incorporated consensus and strain-specific key antigenic determinants from BVDV-1 and 2 strains [24, 25, 31, 47, 48, 52, 56, 57]. Analysis of sequenced genomes showed that the Npro antigen is highly conserved, but the E2 and the NS2-3 antigens have conserved and variable domains. Amino acid sequences of the E2 proteins from currently defined BVDV-1 or BVDV-2 genotypes were aligned and three novel mosaic E2 polypeptides, designated E21,2,3 (E21-3), each containing consensus E2 determinants plus defined strain-specific neutralization epitopes were selected, and wherever there was no consensus at a specific amino acid position for the BVDV-1 genotypes, amino acid from the BVDV-1b sequence was selected since this is the most prevalent sub-type in North America. The E21-3 polypeptide sequence was fused in-frame to the C-termini of the Npro polypeptide and the resultant chimeric polypeptide, designated NproE21-3, was used to generate a codon-optimized synthetic gene, designated npro-e21a-e21b-e22 (nproe21-3,), that also included flag tag sequence at the 3’ end. Two additional mosaic polypeptides that incorporated consensus amino acids from diverse NS2-3 proteins, designated NS2-31 (from BVDV-1 genotypes) and NS2-32 (from BVDV-2 genotypes) were similarly designed and used to generate two synthetic gene sequences, designated ns2-31, and ns2-32, respectively, that also included the flag tag sequence at the 3’ end. Synthetic genes were codon-optimized, custom-made, cloned into pUC57 vector, and sequence-verified by GenScript Inc., NJ, USA.

Generation of recombinant adenovirus plasmid expression constructs

The three synthetic genes (nproe21-3, ns2-31, and ns2-32) were subcloned into pDonR vector using the Gateway Technology (Life Technologies, NY, USA) to generate shuttle constructs. Positive clones were identified by PCR screening of plasmid DNA in bacteria colonies using vector-specific forward primer and gene-specific reverse primer. Authentic entry constructs, designated pDonRNproE21-3, pDonRNS2-31, and pDonRNS2-32, respectively were selected by DNA sequencing. The selected constructs were used to transfer each gene into pAd adenovirus plasmid backbone by homologous recombination (Gateway Technology, Life Technologies, NY, USA) and recombinant constructs were identified by PCR screening as above. Authentic recombinant plasmid constructs, designated pAdNproE21-3, pAdNS2-31, and pAdNS2-32, respectively were selected after DNA sequencing.

Protein expression by plasmid constructs and generation of recombinant adenoviruses

Protein expression was evaluated by immunocytometric analysis of human embryonic kidney (HEK)-293A cells grown in 12-well tissue culture plates and transfected with 1 μg of the selected clones of the pAd DNA constructs, and then probed with anti-FLAG mAb at 48 hr. post-transfection as previously described [58]. Five clones of each pAd construct were selected based on efficiency of protein expression as judged by the immunocytometric analysis, and 2 μg DNA of each construct was digested with Pac-I restriction enzyme. The digested DNA was transfected into HEK-293A cells grown in 6-well plates to generate recombinant adenoviruses that were designated AdNproE21-3, AdNS2-31, and AdNS2-32, respectively. In addition, adenovirus expressing luciferase (AdLuc) was generated to serve as a negative control. Protein expression by the AdNproE21-3, AdNS2-31, and AdNS2-32 adenoviruses was tested and validated by immunocytometric analysis of infected HEK-293A cells as above, whereas fluorescence was used to confirm luciferase expression.

One clone of each recombinant adenovirus was selected for amplification based on protein expression. The bulk viruses were tested for protein expression as above and following titer determination, replication competence of the recombinant adenoviruses was determined by immunocytometric analysis of HEK-293A (which supports adenovirus replication) and MDBK cells (susceptible to adenovirus infection, but do not support replication of replication-incompetent adenovirus) infected overnight with one MOI of each virus construct and then probed with an in-house generated rabbit anti-adenovirus polyclonal IgG (1:500 dilution) followed by an alkaline-phosphatase-conjugated anti-Rabbit IgG (1:1000) (Jackson ImmunoResearch, Cat #711-055-152) secondary antibody and Fast Red TR–Naphthol AS-MX as the substrate (Sigma, F4523) to evaluate infectivity.

Validation of the mosaic antigens using BVDV-specific antibodies and T-cells

Authenticity of the mosaic NproE21-3, NS2-31, and NS2-32 antigens was confirmed by immunocytometric analysis using E2-specific neutralizing monoclonal antibodies (mAbs) and polyclonal antibodies (pAbs) generated against diverse BVDV strains. Briefly, HEK-293A cells grown in 12-well plates were infected for 48 hr. with AdNproE21-3, AdNS2-31, AdNS2-32, or AdLuc and probed with anti-BVDV E2 mAbs 348 and 26A (VMRD, Inc., Pullman, WA), goat anti-BVDV polyclonal sera (VMRD), and bovine anti-BVDV hyperimmune sera from steers immunized and challenged with multiple BVDV-1 and 2 strains [59]. Antigen authenticity was further confirmed by ELISA and Western Blot analysis using the above mentioned antibodies.

The authenticity of the T-cell epitopes in the mosaic antigens was validated by proliferation assays using peripheral blood mononuclear cells (PBMCs) isolated from the BVDV-immunized steers [59]. Recombinant NproE21-3, NS2-31 and NS2-32 antigens were expressed by using recombinant baculoviruses in High Five cells (Thermo Fisher Scientific) generated using the Bac-to-Bac HBM TOPO Secreted Expression System (Thermo Fisher Scientific) as per manufacturer’s instructions and validated as above. These antigens were then affinity purified using Anti-FLAG M2 Affinity Gel (Sigma) and used at 5μg/ml to conduct 3H-Thymidine incorporation assays to quantify antigen-specific T cell responses as previously described [58]. Heat killed BVDV-1b (CA0401186a) and BVDV-2 (A125) at 5μg/ml served as positive control antigens, whereas medium alone was the negative control. The outcome of the cell proliferation was presented as counts per minute (cpm).

Immunization and challenge of calves

Three groups (A, B, and C), of age-matched BVDV sero-negative and virus-free weaned Holstein calves (n = 5) were identified as previously described [60] and used in this study as shown in Table 1. Each calf in group A was inoculated subcutaneously (SQ) with a cocktail, designated AdBVDV, containing the AdNproE21-3, AdNS2-31, and AdNS2-32 recombinant adenoviruses (5 x 1010 TCID50/construct) formulated in adjuvant E (BenchMark-Vaxliant). Each calf in group B was similarly inoculated, but with a commercial MLV BVDV-1 and 2 vaccine (Bovi-Shield Gold, Zoetis Inc., Kalamazoo, MI), whereas each calf in group C was inoculated with the recombinant AdLuc formulated in adjuvant E. Seventy-nine days post-priming, the AdBVDV vaccinees and the negative controls received inoculation of the respective priming immunogen and dose as above. One hundred and forty nine days post-boosting, all the calves were challenged by intranasal administration of 2 x 106 TCID50 of BVDV-1373 using a human nasal atomizer. (http://www.teleflexarcatalog.com/anesthesia-respiratory/airway/categories/552).

Cellular and humoral immune responses

Two weeks post-priming and bi-weekly thereafter, PBMCs were isolated to evaluate and quantify proliferation of BVDV-specific T-cell responses as previously described [58]. The PBMCs (2.5 x 105 cells/well) were cultured for 72 hr. at 37°C in triplicate wells of round-bottom 96-well plates in a total volume of 100 μl of complete RPMI-1640 (cRPMI) medium containing 2.5 μg/ml defined BVDV CD4+ T-cell epitope peptides [32]. The positive control was cRPMI medium containing 1.3 μg/ml concanavalin A (ConA), whereas medium alone served as a negative control. Cells were labeled with 0.25 μCi of 3H-thymidine for 6 hr., harvested using a semi-automatic cell harvester (Tomtec Life Sciences, Hamden, CT), and the incorporated 3H-thymidine was counted with a Micro-Beta liquid scintillation counter (Perkin Elmer, Waltham, MA). The incorporation of 3H-thymidine by the proliferating PBMCs was presented as mean counts per minute (cpm) of triplicate wells (±1 SD).

The PBMCs were also used to quantify BVDV-specific IFN-γ-secreting cells by EliSpot assay as previously described [58]. The PBMCs (2.5 x 105 cells/well) were seeded into triplicate wells of MultiScreen-HA plates (EMD Millipore, Billerica, MA) in a final volume of 100 μl cRPMI medium containing 2.5 μg/ml BVDV CD4+ T-cell epitope peptides. The positive control was 1.3 μg/ml ConA, whereas medium alone served as a negative control. The plates were incubated for 36 hr. at 37°C, washed, developed, and dried overnight as previously described [58]. Following quantification of the spots using an EliSpot reader (AID, Diagnostika GmbH, Germany), the mean number of spots in the negative control wells was subtracted from the mean number of spots in the cognate test wells to determine the mean number of BVDV-specific IFN-γ-secreting PBMCs and the results were presented as the mean number of spot-forming cells/106 PBMCs.

Sera from blood collected two weeks post-boost and one week pre-challenge were tested to determine BVDV-1 and BVDV-2 neutralizing antibody titers using BVDV-1 (Singer, NADL, BJ, TGAC, CA0401186a) and BVDV-2 (890, 1373, A125) strains as previously described [61, 62]. Briefly, serum was heat inactivated at 56°C for 30 min, and 25 μl of each serum was serially diluted (2-fold) in cell culture media without FBS in 96-well microtiter plates. Stock BVDV virus containing 100 TCID50/25μl was added to each test well. In each test, a positive control serum was also included. This serum/virus mixture was incubated for 1 hr., at 37°C, MDBK cells added, and the plates were incubated at 37°C in a humidified atmosphere of 5% CO2 for 72 hr. The cells were observed daily for CPE for cytopathic strains, whereas the non-cytopathic strains were detected by Immuno-peroxidase assay [63]. The results were presented as serum neutralization titers (SNT) [64].

Clinical parameters: Viremia/WBC-platelet counts

Calves were observed daily pre-immunization, post-immunization and post-challenge for coughing, nasal discharge and diarrhea. Rectal temperature post-challenge was taken daily [65]. To determine virus titers post-challenge, blood was collected in vacutainer tubes (containing Sodium-EDTA) by jugular venipuncture, freeze-thawed to lyse cells, centrifuged and supernatants were used to determine BVDV titers as previously described [66]. Whole blood was used to determine CBC using Cell-Dyn 3700 analyzer (Abbott Diagnostics, Lake Forest, IL 60045, USA) with veterinary package as bovines for automated counts (WBC, RBC, Hgb, MCV, PLT). Thin blood smears were stained with Giemsa for differential white blood cell counts [67]. Platelet count verification, WBC count verification, RBC and WBC morphology was evaluated microscopically.

Persistence of recombinant adenovirus in cattle

Presence of recombinant replication-incompetent adenovirus in cattle was tracked for three weeks post-inoculation by rescue of virus from tissue biopsies taken from the intradermal inoculation site. Briefly, recombinant adenovirus (5 x 109 ifu) was inoculated (I.D) into nine marked sites on the neck of four steers. One skin biopsy was taken from each site using a 4mm Biopsy Punch (American Screening, Shreveport, LA) on days 1–7, 14, and 21. In addition, skin biopsies were concurrently collected from the flank region of each steer to serve as negative controls. The steers were euthanized three weeks post-inoculation and samples of draining lymph node and spleen were collected. The fresh tissue samples collected were snap frozen in liquid nitrogen, ground and then resuspended in 1 ml DMEM (Invitrogen). Following centrifugation, supernatants were filtered through 0.22 μm pore membrane, and 0.5 ml was added to one well of HEK-293A cells (which supports adenovirus replication) grown in 12-well plates. Supernatant from HEK-293A cells infected overnight with the recombinant adenovirus, and subjected to the same treatment as above was used as a positive control. Three days post-infection, presence of adenovirus in the HEK-293A cells was evaluated by immunocytometric analysis using the rabbit anti-adenovirus polyclonal IgGs as above. Medium from the above HEK-293A cells was used to infect fresh cells, and seven days later the above process was repeated to confirm presence or absence of adenovirus.

Statistical analysis

Analysis of Variance (ANOVA) followed by Tukey’s multiple comparison test was used to analyze the significance of the differences in BVDV-specific immune responses and disease indices between the treatments (groups A and B) and the negative control (group C) using GraphPad Prism 6 (Version 6.07, GraphPad Software, Inc. La Jolla, USA). Statistical significance was considered when P < 0.05.

Ethics statement

The study was conducted in accordance with the Public Health Service Policy on Humane Care and Use of Laboratory Animals as specified in the Health Research and Extension Act of 1985 (Public Law 99–158) or in accordance with the U.S Department of Agriculture policies as required by the Animal Welfare Act of 1966 (7.USC.2131 et seq) as amended in 1970, 1976, and 1985. The research protocol: AUP21010-65 was reviewed and approved by the Texas A&M University Institutional Animal Care and Use Committee to ensure compliance with PHS standards. All animal care facilities are inspected twice per year. The facilities and procedures for maintenance and care of animals are accredited by the American Association for Accreditation of Laboratory Animal Care. Efforts were made to minimize suffering, and at the completion of the study, the calves were euthanized with an overdose of sodium pentobarbital. This method is approved by the Panel on Euthanasia of the American Veterinary Medical Association.

Results and Discussion

Expression constructs encoding novel mosaic BVDV antigens

Three synthetic genes (designated nproe21-3, ns2-31, and ns2-32) encoding novel BVDV mosaic antigens were designed as depicted in Fig 1A. The nproe21-3 chimeric gene encodes the N-terminal protease fragment (Npro), a consensus BVDV-1a envelope glycoprotein E2 mosaic gene (e21), a consensus BVDV-1b envelope glycoprotein E2 mosaic gene (e22), and a consensus BVDV-2 envelope glycoprotein E2 mosaic gene (e23) fused in-frame to flag-tag. The ns2-31 DNA fragment encodes a consensus BVDV-1 Nonstructural protein 2–3 fused in-frame to flag-tag, whereas the ns2-32 DNA fragment encodes a consensus BVDV-2 Nonstructural protein 2–3 fused in-frame to flag-tag (Fig 1A).

thumbnail
Fig 1. BVDV antigen expression constructs.

A) Schematic diagram of expression cassettes encoding BVDV mosaic antigens: i) Composition of the nproe21-3, chimeric gene: npro encodes Npro antigen; e21 encodes a mosaic BVDV-1a E2 envelope glycoprotein; e22 encodes a mosaic BVDV-1b E2 envelope glycoprotein; whereas e23 encodes a mosaic BVDV-2 E2 envelope glycoprotein. ii) ns2-31 encodes a mosaic BVDV-1 nonstructural protein 2–3. iii) ns2-33 encodes a mosaic BVDV-2 nonstructural protein 2–3. A gene (flag) encoding the FLAG tag was fused in-frame at the 3’ end of each chimeric gene for tracking protein expression and transcription was under the direction of the CMV promoter (CMVp). The genes were cloned into adenovirus backbone plasmid vector and the resultant constructs were designated pAdNproE21-3, pAdNS2-31, and pAdNS2-32, respectively. B) Protein expression by recombinant plasmid constructs: The plasmid DNA constructs encoding the three genes described in (A) above were transfected into HEK-293A cell monolayers and protein expression was evaluated by immunocytometric analysis using anti-FLAG M2-AP Conjugate as follows: HEK-293A cells monolayers were transfected with the following constructs: i) pAdNproE21-3; ii) pAdNS2-31; iii) pAdNS2-32; and iv) pAd vector (negative control). C) Protein expression by recombinant adenovirus constructs: HEK-293A cells monolayers were infected with the following recombinant adenovirus: i) AdNproE21-3; ii) AdNS2-31; iii) AdNS2-32; and iv) Ad-Luciferase. Protein expression was evaluated by immunocytometric analysis as above.

https://doi.org/10.1371/journal.pone.0170425.g001

Expression of the mosaic BVDV antigens

Immunocytometric analysis of HEK-293A cells transfected with the pAdNproE21-3, pAdNS2-31, or pAdNS2-32 constructs probed with anti-FLAG mAb confirmed that each construct expressed the encoded antigen (Fig 1B). Similarly, immunocytometric analysis of HEK-293A cells infected with the AdNproE21-3, AdNS2-31, or AdNS2-32 recombinant adenoviruses probed with anti-FLAG mAb confirmed protein expression (Fig 1C). Analysis of replication competency confirmed that the AdNproE21-3, AdNS2-31, and AdNS2-32 recombinant adenoviruses were replication-incompetent.

Novel mosaic BVDV antigens are recognized by multiple BVDV-specific antibodies

Authenticity of the mosaic antigens (NproE21-3, NS2-31, and NS2-32) expressed by the recombinant adenoviruses was confirmed by immunocytometric analysis of infected HEK-293A cells probed with BVDV neutralizing monoclonal antibodies and polyclonal sera raised against diverse BVDV strains (Fig 2). Anti-BVDV polyclonal sera from immunized goat and cattle reacted with all three recombinant antigens (NproE21-3, NS2-31, and NS2-32), whereas monoclonal antibodies 26A and 348, specific for the glycoprotein E2, reacted with NproE21-3 antigen only (Fig 2A). The outcome confirmed that neutralization epitopes in the mosaic NproE21-3 antigen were correctly expressed, and that the NS2-31 and NS2-32 mosaic antigens were specifically recognized by anti-BVDV polyclonal sera. Thus, these antigens were expected to induce authentic BVDV-specific immune responses in cattle. This expected outcome was consistent with previous demonstration that multicomponent mosaic antigens generated using this strategy elicit broadly protective pathogen-specific immune responses [6870].

thumbnail
Fig 2. Validation of B-cell epitopes in the mosaic BVDV antigens.

Authenticity of the adenovirus-expressed novel BVDV mosaic antigens was confirmed by immunocytometric analysis using E2-specific neutralizing monoclonal antibodies 26A and 348 (both neutralize BVDV-1 & 2); bovine anti-BVDV hyper-immune serum (generated by immunizing steers multiple times with BVDV-1 & 2 vaccines followed by boosting with killed diverse BVDV-1 & 2 strains and then challenged with wild type BVDV-1 & 2 strains (The sera have high BVDV-1 & 2 neutralizing titers [59]); and goat anti-BVDV polyclonal serum generated against multiple wild-type BVDV-1 & 2 strains. A) HEK-293A cells expressing NproE21-3; B) HEK-293A cells expressing NS2-31; C) HEK-293A cells expressing NS2-32; and D) HEK-293A cells expressing luciferase.

https://doi.org/10.1371/journal.pone.0170425.g002

Novel mosaic BVDV antigens are recognized by BVDV-specific T lymphocytes

The NproE21-3, NS2-31, and NS2-32 antigens stimulated robust proliferation of PBMCs from BVDV-immunized steers (Fig 3). The recall responses stimulated by the mosaic antigens were significantly (P<0.01) higher than the responses elicited by whole killed BVDV-1b or BVDV-2, suggesting that the mosaic antigens are likely to prime and amplify robust antigen-specific immune responses in vivo (Fig 3). These outcomes showed that the mosaic antigens were properly processed to generate peptides that were presented by MHC molecules to cognate BVDV-specific memory T-cells. Previous studies have shown that mosaic antigens are processed by host APCs to generate relevant peptides for MHC presentation to elicit protective T-cell responses [71, 72].

thumbnail
Fig 3. Validation of mosaic antigens using BVDV-specific T-cells.

Authenticity of T-cell epitopes in the mosaic BVDV antigens was validated by proliferation assay using PBMCs from a BVDV-1 & 2 hyper-immune steer [59]. The data shown is minus background counts from negative control (media alone) treatment. The asterisks denote a statistically significant difference (P<0.01) between the proliferation induced by the NproE21-3, NS2-31 and the NS2-32 antigens and both whole killed viruses BVDV-1b and BVDV-2. This outcome is representative of assays conducted using PBMCs from other BVDV immune steers.

https://doi.org/10.1371/journal.pone.0170425.g003

Mosaic antigens elicited stronger BVDV-specific T-cell immune responses

Immunogenicity and protective efficacy of the AdNproE21-3, AdNS2-31, and AdNS2-32 recombinant adenovirus cocktail, designated AdBVDV, was evaluated in steers using a homologous prime-boost immunization regimen (Table 1 and Fig 4). One week after the AdBVDV vaccinees were boosted, the cocktail elicited higher, but not significantly different, BVDV-specific IFN-γ-secreting PBMCs as well as BVDV-specific PBMC proliferation compared to the vaccinees that received the commercial MLV BVDV vaccine (Fig 5A & 5C). The mean responses mounted by the AdBVDV vaccinees, but not the MLV vaccinees, were significantly higher (P<0.05) than the negative controls. Before challenge (five months after the AdBVDV vaccinees were boosted) the AdBVDV-induced mean IFN-γ+ response had increased and was significant (P<0.05) compared to the negative controls, whereas the mean IFN-γ+ response in the MLV vaccinees had already declined (Fig 5B). This decrease in the mean IFN-γ response in the MLV BVDV vaccine treatment group one week before challenge, might have had an impact on clearance of the challenge virus.

thumbnail
Fig 4. Immunization timeline.

On day -228 pre-challenge, cattle in the treatment group were vaccinated with a cocktail of the recombinant adenoviruses expressing mosaic BVDV antigens (AdBVDV), whereas positive control cattle received a commercial MLV BVDV vaccine. Negative control cattle were inoculated with the recombinant AdLuc. On day -149 pre-challenge, the cattle were boosted with the respective priming inoculum and dose (Table 1). On day 0, all the cattle were challenged by intranasal delivery of a BVDV-1373 using an atomizer. Blood samples were collected on selected days (0, 3, 6, 10, 12, 13 and 15), whereas clinical observations and rectal temperatures were monitored and recorded daily from days 1–15 post-challenge.

https://doi.org/10.1371/journal.pone.0170425.g004

thumbnail
Fig 5. Mosaic BVDV vaccine elicited strong T-cell responses.

Immunization of cattle with adenovirus-vectored mosaic BVDV vaccine primed and expanded BVDV-specific T-cells. A commercial MLV BVDV vaccine served as a positive control, whereas AdLuciferase served as a negative control. EliSpot assays were used to evaluate BVDV-specific IFN-γ-secreting PBMC responses against defined BVDV T-cell epitopes and data is shown for A) One-week post-boost; and B) Day 0 of challenge. Outcome is presented as IFN-γ+ SFC/106 PBMC. Cell proliferation assays were used to evaluate BVDV-specific PBMC responses and data is shown for C) One-week post-boost; and D) one-week pre-challenge. Proliferation of the PBMCs in response to defined BVDV T-cell epitopes is presented as the means ± standard deviations of 3H-thymidine incorporation by the cells from triplicate wells. In both assays, medium alone served as the negative control and the data shown is minus media background counts. The group mean is represented by a bar. Asterisks denote statistically significant differences, *P<0.05.

https://doi.org/10.1371/journal.pone.0170425.g005

The mean BVDV-specific PBMC proliferation increased in both the vaccinated groups, but only the AdBVDV- and not the MLV-induced response was significantly different (P<0.05) from the AdLuc control group (Fig 5D). The increase in mean IFN-γ response and PBMC proliferation in the AdBVDV treatment group at five months post-boost, were not significantly different from the responses recorded at one week post-boost (Fig 5).

Mosaic antigens elicited cross-protective BVDV-specific antibody responses

Following boosting of the AdBVDV vaccinees, the levels of BVDV neutralizing serum antibodies against five BVDV-1 strains and three BVDV-2 strains were evaluated at one-week post-boost and one-week pre-challenge (Figs 6 and 7). The adenovirus cocktail induced higher mean neutralizing antibody titers post-boost against all BVDV-1 strains compared to the responses stimulated by the commercial MLV BVDV vaccine and the AdLuc controls. The difference between the mean titers however, was significant only for the non-cytopathic BVDV-1b BJ (P<0.05) and BVDV CA0401186a strains (AdBVDV vs MLV, P<0.05; AdBVDV vs AdLuc, P<0.01) (Fig 6A). Furthermore, the mean AdBVDV titers increased up to five months post-boost (one-week pre-challenge) against 4 of 5 BVDV-1 strains whereas, the mean MLV titers either remained the same or declined. These mean AdBVDV titers remained significantly higher (P<0.05) than the MLV vaccinees and the AdLuc controls for the BJ strain, and only the AdLuc controls for the cytopathic BVDV-1a NADL strain. Interestingly, for all three BVDV-2 strains, the mean titers of the MLV vaccinees were higher (in contrast to BVDV-1) than the AdBVDV vaccinees post-boost. These mean MLV titers were significantly higher (P<0.05) than the AdBVDV vaccinees only for strain A125 and significantly higher than the AdLuc controls for all three strains (P<0.05 for strain 890; P<0.01 for strains 1373 and A125). The mean BVDV-2-specific titers in both the AdBVDV vaccinees as well as the MLV vaccinees increased before challenge. Thus overall, the AdBVDV vaccine cocktail was able to induce high titers against all 8 BVDV strains tested in 3 out of 5 calves, whereas the MLV vaccine was able to induce high titers against only BVDV-2 strains. It is also noteworthy that the 3 AdBVDV vaccinees had substantially higher neutralizing titers (1:1024–1:2048) when compared to the MLV vaccinees (1:32–1:256) against the NADL strain which is a component of the commercial MLV vaccine they received.

thumbnail
Fig 6. Mosaic BVDV vaccine induced BVDV-1 specific neutralizing antibodies.

Serum neutralization assays were used to evaluate BVDV-1-specific neutralization titers at A) One-week post-boost; and B) one-week pre-challenge against five BVDV type 1 strains. Mean group titers are represented by the bars. Statistically significant differences between the groups are denoted by asterisks. *P<0.05; **P<0.01.

https://doi.org/10.1371/journal.pone.0170425.g006

thumbnail
Fig 7. Mosaic BVDV vaccine induced BVDV-2 specific neutralizing antibodies.

BVDV-2-specific neutralization titers against three BVDV type 2 strains were evaluated at A) two weeks post-boost; and B) one week pre-challenge. Mean group titers are represented by the bars. Statistically significant differences between the groups are denoted by asterisks *P<0.05; **P<0.01.

https://doi.org/10.1371/journal.pone.0170425.g007

Clinical observations, hematology and viremia

Following the BVDV challenge, there were no obvious differences in clinical score among all the animals, however, characteristic biphasic pyrexia was observed for the negative controls but not for the AdBVDV or the MLV vaccinees (Fig 8A). On day 5 post-challenge, the transient rise in mean rectal temperatures of the negative controls was significantly higher (P<0.001) than the MLV vaccinees but not the AdBVDV vaccinees. The mean rectal temperatures for the negative controls rose again on day 9, peaked at day 10 and normalized by day 11 post-challenge. The mean temperatures of the controls were significantly higher than AdBVDV vaccinees on days 9 (P<0.05) and 10 (P<0.001) post challenge, and the MLV vaccinees on days 8 (P<0.01), 9 (P<0.001) and 10 (P<0.001) post-challenge (Fig 8A). The negative control animals also exhibited transient leucopenia from days 6 to 9 post-challenge with a 32–40% reduction against baseline (day 0) white blood cell (WBC) counts. This reduction of WBCs in the negative controls was significant compared to the AdBVDV vaccinees on days 6 (P<0.05) and 9 (P<0.01) post-challenge, and the MLV vaccinees on days 6 (P<0.01), 7 (P<0.01) and 9 (P<0.001) post-challenge (Fig 8B). There was no significant difference in platelet counts among the treatment groups post-challenge. On days 7 and 10 post-challenge, no virus was detected in all the AdBVDV vaccinees (Table 2). However, BVD virus was detected from the blood of one of the steers that received the commercial MLV BVDV vaccine on day 7 but not on day 10 post-challenge, and from the blood of all the negative controls up to day 15 post-challenge (Table 2).

thumbnail
Fig 8. Clinical manifestations post-challenge.

A) Mean rectal temperature fluctuation; and B) Mean change ratios of white blood cell counts in the vaccinated and negative control groups post-challenge. Asterisks denote statistically significant differences as compared to the negative controls. *P<0.05; **P<0.01 and ***P<0.001.

https://doi.org/10.1371/journal.pone.0170425.g008

thumbnail
Table 2. Virus isolation from calves on day 7 and day 10 post-challenge.

https://doi.org/10.1371/journal.pone.0170425.t002

Recombinant adenovirus inoculated intradermally is short lived

Persistence of recombinant replication-incompetent adenovirus at the intradermal inoculation site, the draining lymph node, and the spleen was monitored by HEK-293A cell-dependent virus rescue followed by immunocytometric analysis using adenovirus-specific polyclonal antibody. One-day post-inoculation, adenovirus was readily recovered from the skin biopsies collected from the inoculation sites, but not from the control sites (Fig 9B and 9C). Virus recovery decreased drastically by day two post-inoculation and very few viral particles were recovered at day three (Fig 9E and 9H). No virus was recoverable from all skin biopsies collected on days 4–7 post-inoculation (Fig 9K). Skin biopsies collected on days 14 and 21, and draining lymph node and spleen samples collected on day 21 were all negative (Fig 9N and 9O). The medium from the HEK-293A cells used to test the samples collected on days 4–7, 14, and 21, was negative after a second round of screening. These outcomes are consistent with previous findings in rodents [73]. Given that ABSL2 biocontainment is required for in vivo studies using the replication-incompetent adenovirus, data from this pilot study suggest that it is safe to downgrade biocontainment after seven days post-inoculation. However, the fate of the vector genome in cattle and environmental risk assessment will need to be determined.

thumbnail
Fig 9. Persistence of replication-incompetent adenovirus in cattle.

Viable recombinant adenovirus inoculated intradermally is only recoverable within three days. Presence of adenovirus rescued from tissue samples of four steers at defined time points was tracked by immunocytometric analysis of HEK-293A cells. Representative data from one steer is shown: A, D, G, J, and M are positive controls at 24 hr., 48 hr., 72 hr., day 7, and day 21, respectively. B, E, H, and K, are skin biopsies taken from the inoculation sites on the neck of the steers at 24 hr., 48 hr., 72 hr., and day 7, respectively, whereas C, F, I, and L, are cognate control skin biopsies taken concurrently from the flank. N and O are draining lymph node and spleen samples, respectively, collected three weeks post-inoculation.

https://doi.org/10.1371/journal.pone.0170425.g009

Conclusions

The purpose of this study was to develop an efficacious prototype BVDV vaccine which a) overcomes the several disadvantages associated with the MLV vaccine mentioned previously and b) provides broad protection against multiple BVDV genotypes. To this end, we designed mosaic polypeptide consensus sequences of highly immunogenic BVDV antigens such as Npro, E2 glycoprotein and the Nonstructural protein 2–3 based on multiple genotypes. We selected live replication deficient adenovirus as a vector for delivery of these antigens to prime strong humoral as well as cell mediated immune responses. Polyclonal anti-BVDV sera and monoclonal anti-E2 antibodies strongly recognized these mosaic antigens by immunocytometric analysis. Furthermore, PBMCs from BVDV immune steers proliferated strongly upon stimulation by these mosaic antigens. The above outcomes confirmed the authenticity of both B-cell and T-cell epitopes in all the mosaic antigens.

Calves immunized with a cocktail of recombinant adenoviruses expressing these antigens had stronger IFN-γ+ and proliferation responses to defined BVDV CD4+ T-cell epitopes as compared to calves vaccinated with the commercial BVDV MLV vaccine. In addition, the AdBVDV vaccinees had higher serum neutralizing titers against BVDV-1 than the MLV vaccinees. In case of BVDV-2, the MLV vaccinees had higher mean titers one week post-boost, but the AdBVDV mean titers increased over time and before challenge were equivalent or higher than the MLV vaccinees for 2 of 3 strains tested. Importantly, both BVDV-1 and BVDV-2 neutralizing antibody titers along with the cellular IFN-γ+ and proliferation immune responses considerably increased for up to five months post-boost (one week before challenge) in most AdBVDV vaccinees, whereas only the BVDV-2 specific titers and the mean proliferation responses amplified in the MLV vaccinees. Upon challenge with a BVDV-2a strain, both vaccinated groups showed no clinical signs of infection. The negative controls however, had a mild fever on day 5 post-challenge followed by a more severe pyrexia on day 10 post-challenge. Moreover, the negative controls also had significantly lower WBC counts than both vaccinated groups. Rapid clearance of virus is an attractive trait in a BVDV vaccine. All the AdBVDV vaccinees had cleared the virus as early as 7 days post-challenge, whereas one MLV vaccinee was still viremic on day 7 but not on day 10 post-challenge. All negative controls remained viremic up to day 15 post-challenge. With regards to the safety concern and ABSL2 biocontainment when using human Ad5 as a delivery vector, we showed that the replication-incompetent Ad5 virus is cleared from the inoculation site within four days post-injection and is not recovered from either the draining lymph node or the spleen after 21 days post-inoculation.

Overall, data from this study showed that the AdBVDV prototype vaccine is more immunogenic and offers better cross-protection than the commercial MLV vaccine in terms of cell mediated and neutralizing antibody responses. As far as protective efficacy is concerned, the AdBVDV vaccine performed at par if not better than the MLV vaccine upon challenge by BVDV-2a strain. Notably, this study is the first to report heterologous protection using subunit BVDV vaccines. Future studies with larger animal sample sizes, different vaccine doses and challenge with diverse BVDV strains need to be conducted to further optimize the AdBVDV prototype vaccine.

The protective potential of the BVDV E2 antigen has been successfully demonstrated in the past using various delivery platforms like live-vectors, DNA immunizations or as a recombinant protein produced in different expression systems [7477]. Current efforts are now focused on enhancing this potential using modern adjuvants and antigen carriers such as PRR activators, APC targeting molecules and silica nanoparticles [7881]. This study highlights the cross-protective potential of the novel mosaic polypeptides and is the first to report heterologous protection using subunit BVDV vaccines. Thus, future studies using these mosaic polypeptide sequences in conjunction with modern immune-response enhancing strategies may lead to a very effective and cross-protective BVDV vaccine.

Acknowledgments

Dr. Glenn A. Holub (Department of Animal Science, Texas A&M University, College Station: for help with sourcing steers), and Dr. Clay Ashley, Dr. Zoe Browning, and Ms. Destiny Taylor (VetMed Park staff) for help with animal handling.

Author Contributions

  1. Conceptualization: SDW WM.
  2. Formal analysis: SL XF WM.
  3. Funding acquisition: SDW WM.
  4. Investigation: SL XF SDW JB LMN KWA CC WM.
  5. Methodology: SDW WM.
  6. Project administration: WM.
  7. Supervision: WM.
  8. Validation: WM.
  9. Visualization: SL XF.
  10. Writing – original draft: SL XF SDW WM.
  11. Writing – review & editing: SL SDW AH CC WM.

References

  1. 1. Fulton RW, Purdy CW, Confer AW, Saliki JT, Loan RW, Briggs RE, et al. Bovine viral diarrhea viral infections in feeder calves with respiratory disease: interactions with Pasteurella spp., parainfluenza-3 virus, and bovine respiratory syncytial virus. Can J Vet Res. 2000;64(3):151–9. pmid:10935880
  2. 2. Campbell JR. Effect of bovine viral diarrhea virus in the feedlot. Vet Clin North Am Food Anim Pract. 2004;20(1):39–50. pmid:15062473
  3. 3. Chase CC, Elmowalid G, Yousif AA. The immune response to bovine viral diarrhea virus: a constantly changing picture. Vet Clin North Am Food Anim Pract. 2004;20(1):95–114. pmid:15062477
  4. 4. Smith RA. North American cattle marketing and bovine respiratory disease (BRD). Animal Health Research Reviews. 2009;10(02):105–8.
  5. 5. Jones C, Chowdhury S. Bovine Herpesvirus Type 1 (BHV-1) is an Important Cofactor in the Bovine Respiratory Disease Complex. Veterinary Clinics of North America: Food Animal Practice. 2010;26(2):303–21. pmid:20619186
  6. 6. Olafson P, Mac CA, Fox FH. An apparently new transmissible disease of cattle. Cornell Vet. 1946;36:205–13. Epub 1946/07/01. pmid:20995890
  7. 7. Ridpath JF, Bolin SR, Dubovi EJ. Segregation of bovine viral diarrhea virus into genotypes. Virology. 1994;205(1):66–74. Epub 1994/11/15. pmid:7975238
  8. 8. Vilcek S, Durkovic B, Kolesarova M, Greiser-Wilke I, Paton D. Genetic diversity of international bovine viral diarrhoea virus (BVDV) isolates: identification of a new BVDV-1 genetic group. Veterinary research. 2004;35(5):609–15. Epub 2004/09/17. pmid:15369663
  9. 9. Giangaspero M, Harasawa R, Weber L, Belloli A. Genoepidemiological evaluation of Bovine viral diarrhea virus 2 species based on secondary structures in the 5 ' untranslated region. J Vet Med Sci. 2008;70(6):571–80. pmid:18628597
  10. 10. Baker JC. The clinical manifestations of bovine viral diarrhea infection. Vet Clin North Am Food Anim Pract. 1995;11(3):425–45. Epub 1995/11/01. pmid:8581856
  11. 11. Brownlie J, Clarke MC, Howard CJ, Pocock DH. Pathogenesis and epidemiology of bovine virus diarrhoea virus infection of cattle. Ann Rech Vet. 1987;18(2):157–66. Epub 1987/01/01. pmid:3619343
  12. 12. Brock KV, Grooms DL, Ridpath J, Bolin SR. Changes in levels of viremia in cattle persistently infected with bovine viral diarrhea virus. J Vet Diagn Invest. 1998;10(1):22–6. Epub 1998/04/04. pmid:9526856
  13. 13. Bolin SR, McClurkin AW, Coria MF. Frequency of persistent bovine viral diarrhea virus infection in selected cattle herds. Am J Vet Res. 1985;46(11):2385–7. Epub 1985/11/01. pmid:3000232
  14. 14. Howard CJ, Clarke MC, Brownlie J. Protection against respiratory infection with bovine virus diarrhoea virus by passively acquired antibody. Vet Microbiol. 1989;19(3):195–203. pmid:2541531
  15. 15. Potgieter LN. Immunology of bovine viral diarrhea virus. Vet Clin North Am Food Anim Pract. 1995;11(3):501–20. pmid:8581860
  16. 16. Dean HJ, Leyh R. Cross-protective efficacy of a bovine viral diarrhea virus (BVDV) type 1 vaccine against BVDV type 2 challenge. Vaccine. 1999;17(9–10):1117–24. pmid:10195622
  17. 17. Fairbanks KK, Rinehart CL, Ohnesorge WC, Loughin MM, Chase CC. Evaluation of fetal protection against experimental infection with type 1 and type 2 bovine viral diarrhea virus after vaccination of the dam with a bivalent modified-live virus vaccine. J Am Vet Med Assoc. 2004;225(12):1898–904. pmid:15643841
  18. 18. Bolin SR, Ridpath JF. Assessment of protection from systemic infection or disease afforded by low to intermediate titers of passively acquired neutralizing antibody against bovine viral diarrhea virus in calves. Am J Vet Res. 1995;56(6):755–9. pmid:7653884
  19. 19. Kelling CL. Evolution of bovine viral diarrhea virus vaccines. Vet Clin North Am Food Anim Pract. 2004;20(1):115–29. Epub 2004/04/06. pmid:15062478
  20. 20. Stokstad M, Brownlie J, Collins ME. Analysis of variation of bovine viral diarrhoea virus E2 sequence following transplacental infection of cattle. Veterinary Microbiology. 2004;102(3–4):141–5. pmid:15327790
  21. 21. Neill JD, Newcomer BW, Marley SD, Ridpath JF, Givens MD. Genetic change in the open reading frame of bovine viral diarrhea virus is introduced more rapidly during the establishment of a single persistent infection than from multiple acute infections. Virus Research. 2011;158(1–2):140–5. pmid:21470568
  22. 22. Neill JD, Newcomer BW, Marley SD, Ridpath JF, Givens MD. Greater numbers of nucleotide substitutions are introduced into the genomic RNA of bovine viral diarrhea virus during acute infections of pregnant cattle than of non-pregnant cattle. Virol J. 2012;9(1):1–8.
  23. 23. Hertig C, Stalder H, Peterhans E. Genetic-Heterogeneity within the Coding Regions of E2 and Ns3 in Strains of Bovine Viral Diarrhea Virus. Gene. 1995;153(2):191–5. pmid:7875587
  24. 24. Deregt D, Bolin S. R., van den Hurk J., Ridpath J. F., and Gilbert S. A.. Mapping of a type 1-specific and a type-common epitope on the E2 (gp53) protein of bovine viral diarrhea virus with neutralization escape mutants. Virus Research 1998.;53(1):81–90. pmid:9617771
  25. 25. Deregt D, van Rijn PA, Wiens TY, van den Hurk J. Monoclonal antibodies to the E2 protein of a new genotype (type 2) of bovine viral diarrhea virus define three antigenic domains involved in neutralization. Virus Res. 1998;57(2):171–81. pmid:9870585
  26. 26. Donis RO, Corapi W, Dubovi EJ. Neutralizing monoclonal antibodies to bovine viral diarrhoea virus bind to the 56K to 58K glycoprotein. J Gen Virol. 1988;69 (Pt 1):77–86.
  27. 27. Liang R, van den Hurk JV, Babiuk LA, van Drunen Littel-van den Hurk S. Priming with DNA encoding E2 and boosting with E2 protein formulated with CpG oligodeoxynucleotides induces strong immune responses and protection from Bovine viral diarrhea virus in cattle. J Gen Virol. 2006;87(Pt 10):2971–82. pmid:16963756
  28. 28. Paton DJ, Lowings JP, Barrett AD. Epitope mapping of the gp53 envelope protein of bovine viral diarrhea virus. Virology. 1992;190(2):763–72. pmid:1381537
  29. 29. Nobiron I, Thompson I, Brownlie J, Collins ME. DNA vaccination against bovine viral diarrhoea virus induces humoral and cellular responses in cattle with evidence for protection against viral challenge. Vaccine. 2003;21(17–18):2082–92. Epub 2003/04/23. pmid:12706698
  30. 30. Endsley JJ, Roth JA, Ridpath J, Neill J. Maternal antibody blocks humoral but not T cell responses to BVDV. Biologicals. 2003;31(2):123–5. pmid:12770543
  31. 31. Collen T, Morrison WI. CD4(+) T-cell responses to bovine viral diarrhoea virus in cattle. Virus Res. 2000;67(1):67–80. pmid:10773320
  32. 32. Collen T, Carr V, Parsons K, Charleston B, Morrison WI. Analysis of the repertoire of cattle CD4(+) T cells reactive with bovine viral diarrhoea virus. Vet Immunol Immunopathol. 2002;87(3–4):235–8. Epub 2002/06/20. pmid:12072240
  33. 33. Howard CJ, Clarke MC, Sopp P, Brownlie J. Immunity to bovine virus diarrhoea virus in calves: the role of different T-cell subpopulations analysed by specific depletion in vivo with monoclonal antibodies. Vet Immunol Immunopathol. 1992;32(3–4):303–14. Epub 1992/05/01. pmid:1321529
  34. 34. Beer M, Wolf G., Pichler J., Wolfmeyer A., and Kaaden O. R.. Cytotoxic T-lymphocyte responses in cattle infected with BVDV. Vet Microbiol 1997.;58:9–22. pmid:9451457
  35. 35. Bolin SR, Grooms DL. Origination and consequences of bovine viral diarrhea virus diversity. Vet Clin North Am Food Anim Pract. 2004;20(1):51–68. Epub 2004/04/06. pmid:15062474
  36. 36. O'Rourke K. BVDV: 40 years of effort and the disease still has a firm hold. Journal of the American Veterinary Medical Association. 2002;220(12):1770–3. pmid:12092944
  37. 37. Theurer ME, Larson RL, White BJ. Systematic review and meta-analysis of the effectiveness of commercially available vaccines against bovine herpesvirus, bovine viral diarrhea virus, bovine respiratory syncytial virus, and parainfluenza type 3 virus for mitigation of bovine respiratory disease complex in cattle. J Am Vet Med Assoc. 2015;246(1):126–42. pmid:25517335
  38. 38. Reber AJ, Tanner M, Okinaga T, Woolums AR, Williams S, Ensley DT, et al. Evaluation of multiple immune parameters after vaccination with modified live or killed bovine viral diarrhea virus vaccines. Comparative immunology, microbiology and infectious diseases. 2006;29(1):61–77. Epub 2006/03/18. pmid:16543046
  39. 39. Ellis J, West K, Cortese V, Konoby C, Weigel D. Effect of maternal antibodies on induction and persistence of vaccine-induced immune responses against bovine viral diarrhea virus type II in young calves. J Am Vet Med Assoc. 2001;219(3):351–6. Epub 2001/08/11. pmid:11497051
  40. 40. Fulton RW, Briggs RE, Payton ME, Confer AW, Saliki JT, Ridpath JF, et al. Maternally derived humoral immunity to bovine viral diarrhea virus (BVDV) 1a, BVDV1b, BVDV2, bovine herpesvirus-1, parainfluenza-3 virus bovine respiratory syncytial virus, Mannheimia haemolytica and Pasteurella multocida in beef calves, antibody decline by half-life studies and effect on response to vaccination. Vaccine. 2004;22(5–6):643–9. pmid:14741155
  41. 41. Kappeler A, LutzWallace C, Sapp T, Sidhu M. Detection of bovine polyomavirus contamination in fetal bovine sera and modified live viral vaccines using polymerase chain reaction. Biologicals. 1996;24(2):131–5. pmid:8889060
  42. 42. Roth JA, Kaeberle ML. Suppression of neutrophil and lymphocyte function induced by a vaccinal strain of bovine viral diarrhea virus with and without the administration of ACTH. Am J Vet Res. 1983;44(12):2366–72. Epub 1983/12/01. pmid:6318614
  43. 43. Grooms DL, Brock KV, Ward LA. Detection of cytopathic bovine viral diarrhea virus in the ovaries of cattle following immunization with a modified live bovine viral diarrhea virus vaccine. J Vet Diagn Invest. 1998;10(2):130–4. pmid:9576338
  44. 44. Foucras G, Corbière F, Tasca C, Pichereaux C, Caubet C, Trumel C, et al. Alloantibodies against MHC Class I: A Novel Mechanism of Neonatal Pancytopenia Linked to Vaccination. The Journal of Immunology. 2011;187(12):6564–70. pmid:22084436
  45. 45. Bell CR, MacHugh ND, Connelley TK, Degnan K, Morrison WI. Haematopoietic depletion in vaccine-induced neonatal pancytopenia depends on both the titre and specificity of alloantibody and levels of MHC I expression. Vaccine. 2015;33(30):3488–96. http://dx.doi.org/10.1016/j.vaccine.2015.05.069. pmid:26055292
  46. 46. Bell CR, Rocchi MS, Dagleish MP, Melzi E, Ballingall KT, Connelly M, et al. Reproduction of bovine neonatal pancytopenia (BNP) by feeding pooled colostrum reveals variable alloantibody damage to different haematopoietic lineages. Vet Immunol Immunopathol. 2013;151(3–4):303–14. pmid:23273932
  47. 47. Korber BT, Letvin NL, Haynes BF. T-cell vaccine strategies for human immunodeficiency virus, the virus with a thousand faces. J Virol. 2009;83(17):8300–14. pmid:19439471
  48. 48. Santra S, Liao HX, Zhang RJ, Muldoon M, Watson S, Fischer W, et al. Mosaic vaccines elicit CD8(+) T lymphocyte responses that confer enhanced immune coverage of diverse HIV strains in monkeys. Nat Med. 2010;16(3):324–U122. pmid:20173754
  49. 49. Kamlangdee A, Kingstad-Bakke B, Anderson TK, Goldberg TL, Osorio JE. Broad protection against avian influenza virus by using a modified vaccinia Ankara virus expressing a mosaic hemagglutinin gene. J Virol. 2014;88(22):13300–9. pmid:25210173
  50. 50. van Oirschot JT, Bruschke CJ, van Rijn PA. Vaccination of cattle against bovine viral diarrhoea. Vet Microbiol. 1999;64(2–3):169–83. pmid:10028171
  51. 51. Deregt D, Dubovi EJ, Jolley ME, Nguyen P, Burton KM, Gilbert SA. Mapping of two antigenic domains on the NS3 protein of the pestivirus bovine viral diarrhea virus. Vet Microbiol. 2005;108(1–2):13–22. pmid:15917132
  52. 52. Hegde NR, Srikumaran S. The use of bovine MHC class I allele-specific peptide motifs and proteolytic cleavage specificities for the prediction of potential cytotoxic T lymphocyte epitopes of bovine viral diarrhea virus. Virus Genes. 1997;14(2):111–21. pmid:9237350
  53. 53. Howard CJ, Clarke M. C., Sopp P. and Brownlie J.. Immunity to bovine viral diarrhoea virus in calves: the role of different T cell subpopulations analysed by specific depletion in vivo with monoclonal antibodies. Vet Immunol Immunopathol 1992.;32:303–14. pmid:1321529
  54. 54. Moraes MP, Segundo FD, Dias CC, Pena L, Grubman MJ. Increased efficacy of an adenovirus-vectored foot-and-mouth disease capsid subunit vaccine expressing nonstructural protein 2B is associated with a specific T cell response. Vaccine. 2011;29(51):9431–40. pmid:22027486
  55. 55. Grubman MJ, Segundo FDS, Dias CCA, Moraes MP, Perez-Martin E, de los Santos T. Use of replication-defective adenoviruses to develop vaccines and biotherapeutics against foot-and-mouth disease. Future Virol. 2012;7(8):767–78.
  56. 56. Fischer W, Perkins S, Theiler J, Bhattacharya T, Yusim K, Funkhouser R, et al. Polyvalent vaccines for optimal coverage of potential T-cell epitopes in global HIV-1 variants. Nat Med. 2007;13(1):100–6. pmid:17187074
  57. 57. van Rijn PA. A common neutralizing epitope on envelope glycoprotein E2 of different pestiviruses: implications for improvement of vaccines and diagnostics for classical swine fever (CSF)? Vet Microbiol. 2007;125(1–2):150–6. pmid:17561359
  58. 58. Njongmeta LM, Bray J, Davies CJ, Davis WC, Howard CJ, Hope JC, et al. CD205 antigen targeting combined with dendritic cell recruitment factors and antigen-linked CD40L activation primes and expands significant antigen-specific antibody and CD4+ T cell responses following DNA vaccination of outbred animals. Vaccine. 2012;30(9):1624–35. http://dx.doi.org/10.1016/j.vaccine.2011.12.110. pmid:22240344
  59. 59. Wang F, Ekiert DC, Ahmad I, Yu W, Zhang Y, Bazirgan O, et al. Reshaping antibody diversity. Cell. 2013;153(6):1379–93. pmid:23746848
  60. 60. Ridpath JF, Hietala SK, Sorden S, Neill JD. Evaluation of the reverse transcription-polymerase chain reaction/probe test of serum samples and immunohistochemistry of skin sections for detection of acute bovine viral diarrhea infections. J Vet Diagn Invest. 2002;14(4):303–7. pmid:12152809
  61. 61. Carbrey EA, Brown LN, Chow TL, Kahrs RF, McKercher DG, Smithies LK, et al., editors. Recommended standard laboratory techniques for diagnosing infectious bovine rhinotracheitis, bovine virus diarrhea, and shipping fever (parainfluenza-3). US Animal Health Association; 1971: USAHA.
  62. 62. Edwards S. The diagnosis of bovine virus diarrhoea-mucosal disease in cattle. Rev Sci Tech. 1990;9(1):115–30. pmid:1966717
  63. 63. Fulton RW, Saliki JT, Burge LJ, d'Offay JM, Bolin SR, Maes RK, et al. Neutralizing antibodies to type 1 and 2 bovine viral diarrhea viruses: detection by inhibition of viral cytopathology and infectivity by immunoperoxidase assay. Clin Diagn Lab Immunol. 1997;4(3):380–3. pmid:9144381
  64. 64. Reed LJ, Muench H. A simple method of estimating fifty per cent endpoints. American journal of epidemiology. 1938;27(3):493–7.
  65. 65. Fairbanks K, Schnackel J, Chase CC. Evaluation of a modified live virus type-1a bovine viral diarrhea virus vaccine (Singer strain) against a type-2 (strain 890) challenge. Vet Ther. 2003;4(1):24–34. pmid:12756633
  66. 66. Saliki JT, Fulton RW, Hull SR, Dubovi EJ. Microtiter virus isolation and enzyme immunoassays for detection of bovine viral diarrhea virus in cattle serum. J Clin Microbiol. 1997;35(4):803–7. pmid:9157132
  67. 67. Benjamin MM. Outline of veterinary clinical pathology: Iowa State University Press.; 1978.
  68. 68. Barouch DH, O'Brien KL, Simmons NL, King SL, Abbink P, Maxfield LF, et al. Mosaic HIV-1 vaccines expand the breadth and depth of cellular immune responses in rhesus monkeys. Nat Med. 2010;16(3):319–23. pmid:20173752
  69. 69. Barouch DH, Stephenson KE, Borducchi EN, Smith K, Stanley K, McNally AG, et al. Protective efficacy of a global HIV-1 mosaic vaccine against heterologous SHIV challenges in rhesus monkeys. Cell. 2013;155(3):531–9. pmid:24243013
  70. 70. Fischer W, Perkins S, Theiler J, Bhattacharya T, Yusim K, Funkhouser R, et al. Polyvalent vaccines for optimal coverage of potential T-cell epitopes in global HIV-1 variants. Nat Med. 2007;13(1):100–6. pmid:17187074
  71. 71. Walsh SR, Moodie Z, Fiore-Gartland AJ, Morgan C, Wilck MB, Hammer SM, et al. Vaccination With Heterologous HIV-1 Envelope Sequences and Heterologous Adenovirus Vectors Increases T-Cell Responses to Conserved Regions: HVTN 083. J Infect Dis. 2016;213(4):541–50. pmid:26475930
  72. 72. Yusim K, Dilan R, Borducchi E, Stanley K, Giorgi E, Fischer W, et al. Hepatitis C genotype 1 mosaic vaccines are immunogenic in mice and induce stronger T-cell responses than natural strains. Clin Vaccine Immunol. 2013;20(2):302–5. pmid:23221002
  73. 73. Reuter JD, Fang XQ, Ly CS, Suter KK, Gibbs D. Assessment of hazard risk associated with the intravenous use of viral vectors in rodents. Comparative Med. 2012;62(5):361–70.
  74. 74. Elahi SM, Shen SH, Talbot BG, Massie B, Harpin S, Elazhary Y. Recombinant adenoviruses expressing the E2 protein of bovine viral diarrhea virus induce humoral and cellular immune responses. FEMS microbiology letters. 1999;177(1):159–66. pmid:10436933
  75. 75. Couvreur B, Letellier C, Olivier F, Dehan P, Elouahabi A, Vandenbranden M, et al. Sequence-optimised E2 constructs from BVDV-1b and BVDV-2 for DNA immunisation in cattle. Veterinary research. 2007;38(6):819–34. pmid:17727807
  76. 76. Nelson G, Marconi P, Periolo O, La Torre J, Alvarez M. Immunocompetent truncated E2 glycoprotein of bovine viral diarrhea virus (BVDV) expressed in Nicotiana tabacum plants: a candidate antigen for new generation of veterinary vaccines. Vaccine. 2012;30(30):4499–504. pmid:22554468
  77. 77. Thomas C, Young N, Heaney J, Collins M, Brownlie J. Evaluation of efficacy of mammalian and baculovirus expressed E2 subunit vaccine candidates to bovine viral diarrhoea virus. Vaccine. 2009;27(17):2387–93. pmid:19428855
  78. 78. R El-Attar LM, Thomas C, Luke J, A Williams J, Brownlie J. Enhanced neutralising antibody response to bovine viral diarrhoea virus (BVDV) induced by DNA vaccination in calves. Vaccine. 2015;33(32):4004–12. pmid:26079613
  79. 79. Pecora A, Malacari DA, Perez Aguirreburualde MS, Bellido D, Nuñez MC, Dus Santos MJ, et al. Development of an APC-targeted multivalent E2-based vaccine against Bovine Viral Diarrhea Virus types 1 and 2. Vaccine. 2015;33(39):5163–71. pmid:26279338
  80. 80. Mody K, Mahony D, Cavallaro A, Zhang J, Zhang B, Mahony T, et al. Silica Vesicle Nanovaccine Formulations Stimulate Long-Term Immune Responses to the Bovine Viral Diarrhoea Virus E2 Protein. PLoS One. 2015;10(12):e0143507–e. pmid:26630001
  81. 81. Snider M, Garg R, Brownlie R, van den Hurk JV, van Drunen Littel-van den Hurk S. The bovine viral diarrhea virus E2 protein formulated with a novel adjuvant induces strong, balanced immune responses and provides protection from viral challenge in cattle. Vaccine. 2014;32(50):6758–64. pmid:25454860