Newcastle disease virus-vectored West Nile fever vaccine is immunogenic in mammals and poultry
Virology Journal volume 13, Article number: 109 (2016)
West Nile virus (WNV) is an emerging zoonotic pathogen which is harmful to human and animal health. Effective vaccination in susceptible hosts should protect against WNV infection and significantly reduce viral transmission between animals and from animals to humans. A versatile vaccine suitable for different species that can be delivered via flexible routes remains an essential unmet medical need. In this study, we developed a recombinant avirulent Newcastle disease virus (NDV) LaSota strain expressing WNV premembrane/envelope (PrM/E) proteins (designated rLa-WNV-PrM/E) and evaluated its immunogenicity in mice, horses, chickens, ducks and geese.
Mouse immunization experiments disclosed that rLa-WNV-PrM/E induces significant levels of WNV-neutralizing antibodies and E protein-specific CD4+ and CD8+ T-cell responses. Moreover, recombinant rLa-WNV-PrM/E elicited significant levels of WNV-specific IgG in horses upon delivery via intramuscular immunization, and in chickens, ducks and geese via intramuscular, oral or intranasal immunization.
Our results collectively support the utility of rLa-WNV-PrM/E as a promising WNV veterinary vaccine candidate for mammals and poultry.
West Nile virus (WNV) is the causative agent of West Nile fever (WNF), a major emerging zoonotic disease shown to have a significant negative impact on both human and animal health since the first recorded case in Uganda in 1937. WNV is a member of the genus Flavivirus belonging to the family Flaviviridae. The virus is one of the most widespread arthropod-transmitted pathogens, and is extensively distributed worldwide throughout Africa, Europe, Asia and North America. WNV has a broad host spectrum comprising several species of birds (including poultry), mammals, amphibians and reptiles. Culex mosquitoes play an important role as the primary global WNV transmission vector, and are responsible for the incidental infection of humans and horses, which are considered dead-end hosts of WNV [1–4].
Vaccination in sensitive host animals, especially those abundant in number and closely associated with humans, such as horses, poultry and other bird species, should protect against WNV infection and significantly reduce transmission between animals and from animals to humans. Currently, several injection-delivered vaccines [5–8] are licensed for horses, but not other sensitive host animals. A versatile vaccine suitable for different species that can be delivered via flexible administration routes therefore remains an unmet medical requirement.
Newcastle disease virus (NDV) has been actively developed and evaluated as a vaccine vector for the control of human and animal diseases [9–16]. NDV vector vaccines can be effectively delivered via intramuscular or intratracheal inoculation in mammals and intramuscular, intranasal or oral (through water or feed) inoculation in poultry [11, 12, 17–21]. In the current study, we generated a recombinant nonvirulent NDV LaSota virus strain expressing WNV pre-membrane (PrM) and envelope protein (E), two surface glycoproteins that form a heterodimer on the viral surface  and are responsible for eliciting the majority of protective immune responses . Immunogenicity of the recombinant NDV in mammals and poultry delivered via different immunization routes was further evaluated.
Construction of recombinant NDV LaSota virus
The chemically synthesized mammalian codon-optimized WNV PrM/E gene (strain NY99, GenBank No. DQ211652.1) was cloned and inserted into the Pme I site between the P and M genes of full-length genomic cDNA of NDV LaSota . The resultant plasmid was co-transfected with eukaryotic plasmids expressing NDV nucleoprotein (NP), phosphate protein (P) and large polymerase protein (L), following an established protocol . The rescued recombinant virus was designated rLa-WNV-PrM/E. Expression of WNV PrM and E proteins was confirmed via indirect immunofluorescence and western blot assays. Mouse anti-WNV E monoclonal antibody (developed in our laboratory), mouse anti-PrM monoclonal antibody  and chicken anti-NDV serum  was used as primary antibodies. Fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse antibody (Sigma, St. Louis, MO) and Tetramethylrhodamine (TRITC)-conjugated rabbit anti-chicken antibody (Sigma, St. Louis, MO) was used as secondary antibodies for immunofluorescence assay. Chicken anti-NDV serum and mouse anti-WNV serum (developed in our laboratory) were used as primary antibodies, horseradish-peroxidase (HRP)-conjugated goat anti-chicken IgG and goat anti-mouse IgG (SouthernBiotech, Birmingham, AL) were used as secondary antibodies for western blot assay.
To determine the pathogenicity of rLa-WNV-PrM/E in poultry, mean death time, intracerebral pathogenicity index, and intravenous pathogenicity index were determined in embryonated specific pathogen-free (SPF) chickens or eggs according to the OIE Manual . To assess pathogenicity in mouse, ten 6-week-old female C57BL/6 mice (Vital River, Beijing, China) were inoculated intramuscularly with 0.1 ml diluted allantoic fluid containing 1 × 108 EID50 (50 % Embryo Infectious Dose) rLa-WNV-PrM/E and intranasally with 0.03 ml diluted allantoic fluid containing 3 × 107 EID50 rLa-WNV-PrM/E. Mice were examined daily for 3 weeks for signs of illness, weight loss or death.
Animal immunization studies
For mouse immunization, ten 6-week-old female C57BL/6 mice (Vital River, Beijing, China) were intramuscularly vaccinated with 0.1 ml diluted allantoic fluid containing 1 × 108 EID50 rLa-WNV-PrM/E twice with a 3-week interval. Splenocytes for assay of E protein-specific CD4+ and CD8+ T-cell responses were harvested 10 days after the first or second dose. Serum samples for the serological assay were prepared 2 weeks after each dose.
For horse immunization, five adult horses were intramuscularly inoculated with 2 ml diluted allantoic fluid containing 2 × 109 EID50 rLa-WNV-PrM/E, and five administered with 2 ml phosphate-buffered saline (PBS) as the control group. Three weeks after the first dose, a booster with the same vaccine was delivered using the same dosage and route. Serum samples were collected for serological assay 2 weeks after each immunization.
For poultry immunization, three groups (ten per group) of 4-week-old SPF chickens were assessed: intramuscular inoculation with 0.1 ml diluted allantoic fluid containing 1 × 108 EID50 rLa-WNV-PrM/E (Group One), oral inoculation with 10 ml diluted allantoic fluid containing 1 × 1010 EID50 rLa-WNV-PrM/E mixed with 500 g chicken feed and 300 ml water (Group Two), whereby feeding was stopped 5 h before inoculation, and intramuscular and oral inoculation with PBS (Group Three). Three groups (ten per group) of 4-week-old SPF ducks were immunized following the above procedure. For immunization of geese, four groups (15 per group) of 4-week-old birds were examined: intramuscular inoculation with 0.5 ml diluted allantoic fluid containing 5 × 108 EID50 rLa-WNV-PrM/E (Group One), intranasal inoculation with 0.5 ml diluted allantoic fluid containing 5 × 108 EID50 rLa-WNV-PrM/E via eye drops and nostril instillation (Group Two), oral inoculation with 0.5 ml diluted allantoic fluid containing 5 × 108 EID50 rLa-WNV-PrM/E via buccal cavity instillation (Group Three), and intramuscular inoculation with 0.5 ml PBS (Group Four). Three weeks after the first dose, chickens, ducks and geese were boosted with the vaccine using the same doses and routes. Serum samples were collected for serological assay 2 weeks after each immunization. All poultry were housed in the Experimental Animal Center of Harbin Veterinary Research Institute.
Analysis of WNV-specific IgG, neutralizing and NDV HI antibodies
Enzyme-linked immunosorbent assay (ELISA) for determining antigen-specific IgG in mouse serum was performed as described previously . Briefly, purified mammalian cells producing WNV virus-like particles (4 μg/ml, containing PrM and E proteins, unpublished) were used as coating antigen. Antibodies were detected using HRP-labeled goat anti-mouse IgG (SouthernBiotech, Birmingham, AL) secondary antibody. A standard curve was generated by coating with serially diluted mouse IgG (Southern Biotech, Birmingham, AL) at known concentrations. A linear equation was obtained based on the standard IgG concentration and their O.D values, thus the concentration of WNV-specific IgG was calculated according to the linear equation based on their O.D values and expressed as the amount of IgG per ml of serum (ng/ml). The above coating antigen was also used for ELISA detection of WNV-specific IgG in horse and poultry sera. HRP-labeled goat anti-horse IgG (SouthernBiotech, Birmingham, AL) and goat anti-chicken IgG (SouthernBiotech, Birmingham, AL) were used as secondary antibodies for horse and chicken serum detection, and mouse anti-duck IgG (AbD Serotec, Oxford, UK) and HRP-labeled goat anti-mouse IgG (SouthernBiotech, Birmingham, AL) for duck and goose serum detection. Due to the lack of purified IgG for these animals, results were expressed as O.D. values relative to negative controls.
Mouse serum neutralizing antibody levels were determined using the WNV plaque reduction neutralization test (PRNT) in the Biosafety Level 3 facility of Beijing Institute of Microbiology and Epidemiology. Briefly, 320 μl of 10-fold serially diluted mouse serum (heat-inactivated at 56 °C for 30 min before use) was mixed with 320 μl medium containing 150 plaque-forming units (PFU) of WNV (strain NY99) and incubated at 37 °C for 1 h. Next, the mixture was added to BHK-21 cells in the wells of a six-well plate and incubated at 37 °C for 1 h. Following removal of the mixture, cells washed three times with PBS. Cells were overlaid with 2 ml DMEM-agarose, and incubation continued at 37 °C. After 72 h, cells were fixed with 4 % paraformaldehyde and subsequently stained with 1.5 % crystal violet to visualize plaques. Neutralization titers were expressed as the reciprocal of the highest dilution of serum showing at least 50 % reduction in number of plaques, compared with the negative control. Neutralizing antibodies of chicken, duck, goose and horse sera were not assessed due to unavailability of the BSL-3 facility during the experimental period. NDV hemagglutinin inhibition (HI) antibodies of immunized animals were determined following a previously described protocol .
Flow cytometric analysis of the mouse CD4+ and CD8+ T-cell response
The WNV E protein-specific CD4+ and CD8+ T-cell response in C57BL/6 mice was determined via flow cytometry using established protocols . rLa-WNV-PrM/E-immunized mice were sacrificed on day 10 after the first and second immunizations. Mouse splenocytes were prepared as documented by Ye et al. . Briefly, spleens were removed from euthanized mice, cut into small sections, and homogenized by gentle rubbing. After low-speed centrifugation, the supernatant was removed, cells gently re-suspended in red blood cell lysis buffer (Sigma), and incubated on ice for 1 min. Splenocytes (1 × 106) were stimulated with 20 μg/ml WNV E-specific CD4 peptide (PVGRLVTVNPFVSVA, H-2b, ) or CD8 peptide (LGMSNRDFL, H-2Db,  for 6 h in presence of 10 ng/ml Brefeldin A (eBioscience, San Diego, CA) to assess the CD4+ and CD8+ T-cell response, respectively. Cells were washed twice with PBS containing 3 % fetal calf serum and subsequently stained with Peridinin-Chlorophyll-Protein-Complex (PerCP)-conjugated rat anti-mouse CD4 (or CD8) and phycoerythrin (PE)-conjugated rat anti-mouse CD3 antibody (BD Pharmingen, San Diego, CA). Next, cells were fixed and permeabilized with Fix&Perm Buffer (eBioscience, San Diego, CA) and stained for intracellular interferon-gamma (IFN-γ) with an allophycocyanin (APC)-conjugated rat anti-mouse IFN-γ antibody (BD Pharmingen). The levels of CD4+ or CD8+ T-cell responses were determined using flow cytometry on a BD FACSAria Station (BD Immunocytometry Systems, San Jose, CA). Data were analyzed with FlowJo software (Treestar Inc, Ashland, OR).
Data on virus titers, antibody titers and mouse T cell responses were analyzed using two-tailed Student’s t test with the Excel program (Microsoft, Redmond, WA). To describe the p value significance, the following convention was used: not significant, p > 0.05; significant, p ≤ 0.05; highly significant, p ≤ 0.01.
Generation of rLa-WNV-PrM/E virus and in vitro characterization
Recombinant NDV expressing WNV PrM/E proteins was generated by inserting the PrM/E gene between the P and M genes in NDV genome cDNA (Fig. 1a). The presence of PrM/E was confirmed via RT-PCR. PrM/E protein expression was confirmed via western blot and indirect immunofluorescence staining of rLa-WNV-PrM/E-infected BHK-21 cells. Western blot detected the presence of both E (~45 kDa) and PrM (~25 kDa) proteins (Fig. 1b), which were further confirmed via indirect immunofluorescence with specific monoclonal antibodies against each protein. As expected, rLa-infected BHK-21 cells were not stained (Fig. 1c).
The growth titer of rLa-WNV-PrM/E in embryonated chicken eggs was comparable to that of parental rLaSota. Genetic stability of rLa-WNV-PrM/E was assessed by serial passage of the virus in SPF chicken eggs, and confirmed with RT-PCR and immunofluorescence (data not shown). Mean death time (>120 h), intracerebral pathogenicity index (=0), and intravenous pathogenicity index (=0) results demonstrated the lentogenic nature of rLa-WNV-PrM/E in poultry (data not shown). The genetic stability of WNV PrM/E gene within rLa-WNV-PrM/E was assessed by serially passage (at least 10 passages) of the virus in embryonated SPF chicken eggs, the presence and expression of PrM/E was confirmed by RT-PCR and indirect immunofluorescence assay. The results demonstrated the PrM/E gene can be stably maintained and expressed. Ten mice receiving intramuscular inoculation at a dose of 1 × 108 EID50 and intranasal inoculation of 3 × 107 EID50 rLa-WNV-PrM/E survived with no abnormalities during the 2-week observation period. No significant differences in weight gain were observed after inoculation. Our results indicate rLa-WNV-PrM/E is safe for mice (data not shown).
The recombinant virus induces significant WNV-specific humoral and T-cell responses in mice
WNV-specific IgG (Fig. 2a) was detected using ELISA. Notably, the IgG antibody level was significantly boosted after the second dose (p < 0.01). Serum neutralizing antibodies were analyzed with a WNV plaque reduction assay. As shown in Fig. 2b, WNV-neutralizing antibodies were detected after the first dose, and significantly boosted after the second dose (p < 0.01). NDV neutralizing antibodies were also detected after the first dose, and significantly boosted after the second dose (p < 0.01) (Fig. 2c).
The WNV E protein-specific CD4 epitope (PVGRLVTVNPFVSVA, H-2b,  and CD8 epitope (LGMSNRDFL, H-2Db,  were used to assess rLa-WNV-PrM/E-induced specific T-cell responses in mice. As shown in Fig. 3, rLa-WNV-PrM/E immunization induced significant epitope-specific IFN-γ-producing CD4+ and CD8+ T-cell responses after the first dose, which were significantly boosted after the second dose (p < 0.05).
rLa-WNV-PrM/E administered via different immunization routes induces significant WNV IgG antibody production in horses and poultry
Given that rLa-WNV-PrM/E induces good humoral responses in mice, we performed horse immunization with the vaccine. Horses received two doses of the vaccine via the intramuscular route with a 3-week interval. WNV-specific IgG was detected after the first dose, and significantly boosted after the second dose (p < 0.01) (Fig. 4a). HI antibodies against NDV were also detected in horse after the first dose, and significantly boosted after the second dose (p < 0.01) (Fig. 4b). To determine whether rLa-WNV-PrM/E induces an immune response in poultry, SPF chickens were intramuscularly or orally inoculated with the recombinant virus twice with a 3-week interval. In intramuscularly immunized chickens, the WNV-specific IgG was detected after the first dose, and significantly boosted after the second dose (p < 0.05) (Fig. 5a I). In orally immunized chickens, IgG was also detected after the first dose, but only slightly boosted after the second dose (Fig. 5a II). NDV HI antibodies was detected after the first dose, and significantly boosted after the second dose (p < 0.05) in intramuscularly immunized chickens (Fig. 5b I). In orally immunized chickens, NDV HI antibody was also detected after the first dose, but only slightly boosted after the second dose with no statistical significance (Fig. 5b II). The same immunization procedure was performed for ducks. WNV IgG in intramuscularly (Fig. 6a I) and orally (Fig. 6a II) immunized duck sera was induced after the first dose, and boosted significantly after the second dose (p < 0.05). NDV HI antibodies were detected after the first dose, and significantly boosted after the second dose in intramuscularly (Fig. 6b I) and orally immunized ducks (p < 0.01) (Fig. 6b II). Groups of outbred geese were either intramuscularly, intranasally or orally inoculated with rLa-WNV-PrM/E. Intramuscularly immunized geese produced detectable WNV IgG after the first dose, which was significantly boosted after the second dose (p < 0.05) (Fig. 7a I). Intranasally (Fig. 7a II) and orally (Fig. 7a III) immunized geese also produced a detectable level of IgG after the first dose, which was only slightly boosted after the second dose (p > 0.05). NDV HI antibodies were detected after the first dose, and significantly boosted after the second dose in intramuscularly (Fig. 7b I), intranasally (Fig. 7b II) and orally (Fig. 7b III) immunized geese (p < 0.01).
WNV is an important zoonotic pathogen widely distributed geographically, with emergence of increasingly neuroevasive strains. Here, a recombinant NDV LaSota virus expressing WNV PrM and E proteins, rLa-WNV-PrM/E, was constructed as a candidate veterinary vaccine for WNV prevention and control. rLa-WNV-PrM/E elicited significant levels of neutralizing antibodies and WNV-specific T-cell responses in mice, as well as WNV-specific IgG in horses, chickens, ducks, and geese, support the immunogenicity of the newly generated recombinant virus in mammals and poultry.
Mice are sensitive to WNV infection, and thus commonly used as the model animal for WNV vaccine evaluation and other related studies. In our experiments, mice intramuscularly inoculated with rLa-WNV-PrM/E produced significant WNV-neutralizing antibodies and specific IgG. Neutralizing antibodies play a crucial role in WNV control and clearance . We used the 50 % plaque reduction assay for determining the levels of neutralizing antibody against WNV NY99 in mice sera. This method is recommended by WHO for testing the potency of Japanese encephalitis vaccine. The cut-off value for testing serum seroprotection is 1 log10 (a ten-fold dilution of serum that reduces plaque formation by 50 % is sufficient for protection against viral challenge) . In an earlier study, mice immunized actively or passively that possessed WNV-neutralizing antibodies higher than 1 log10 were protected against the lethal WNV challenge . In our experiments, the neutralizing antibody titer of rLa-WNV-PrM/E-immunized mice reached up to 1.3 log10 after the first dose. After administration of the second dose, the neutralizing antibody titer was significantly boosted (~2.2 log10), implying that rLa-WNV-PrM/E confers robust protection against lethal WNV infection in mice. In sera of mice, high levels of anti-WNV E IgG were elicited after the first dose, which were significantly boosted after the second dose. IgG findings were in accordance with the neutralizing antibody pattern, indicating a linear correlation in rLa-WNV-PrM/E-immunized animals. T-cell responses are additionally critical in controlling WNV infection. CD4+ T-cells play a dominant protective role in viral clearance via facilitating antibody responses and sustaining WNV-specific CD8+ T-cell responses in the central nervous system (CNS) . The efficacy of CD8+ T cells in controlling WNV infection has also been characterized. An earlier study showed that while neutralizing antibodies play a central role in terminating WNV viremia, CD8+ T-cells are essential for preventing sustained WNV infection in peripheral and CNS compartments . Live vector vaccines have a significant advantage in that they effectively elicit cellular responses [36–38]. In our experiments, rLa-WNV-PrM/E induced high levels of WNV-specific CD4+ and CD8+ T-cell responses. Although a challenge study was not conducted at this time due to the unavailability of the BSL-3 laboratory, given the neutralizing antibody and T-cell response results, we presume that the rLa-WNV-PrM/E could confer protection in mice. Intramuscularly immunized horses produced WNV specific IgG following the first dose, which was significantly boosted after the second dose. Considering the close association between IgG and neutralizing antibody levels, we propose that horses acquire protective immunity after vaccination. Further neutralization assays and challenge studies are required to confirm the efficacy of the vaccine in horses.
Vaccination of sensitive hosts not only protects the animal itself but also prevents transmission of WNV from animals to humans. Several veterinary WNV vaccines are currently available, including inactivated whole virus vaccine [5, 39, 40], DNA vaccines [41–43], recombinant canarypox-vectored vaccine [6, 44] and recombinant Yellow Fever 17D vaccine [45, 46]. Notably, canarypox-vectored WNV vaccine has been shown to effectively elicit WNV-specific neutralizing antibodies and confer protection in horses, geese, cats and dogs against lethal WNV challenge [6, 44, 47, 48]. These vaccines require delivery via intramuscular inoculation. The current study demonstrated that rLa-WNV-PrM/E is immunogenic in not only mice, horses and poultry when administered intramuscularly, but also in poultry upon delivery via intranasal or oral inoculation. Based on the collective findings, we propose that rLa-WNV-PrM/E is a promising veterinary candidate vaccine for multiple mammalian and avian species that can be delivered via flexible inoculation routes.
Domestic poultry, such as chickens, ducks and geese, are susceptible to WNV and develop clinical signs and viremia (usually sufficient to infect mosquitoes), thus contributing to WNV transmission. Chickens are widely used as sentinel animals in the early warning of WNV prevalence [2, 49–52]. Several studies have additionally provided evidence of the susceptibility of domestic or captive ducks to WNV [3, 53, 54]. Ducks develop high-titer viremia in blood and are capable of shedding virus orally . Geese are also susceptible to WNV, especially young geese, and develop a variety of neurological signs, often resulting in a significant number of deaths [4, 56, 57]. Geese represent another experimental animal model for WNV vaccine evaluation [58–60]. Notably, WNV human infection and isolation of the virus has recently been reported in mosquitoes in the Xinjiang Uygur Autonomous Region in Northwest China [61, 62]. China has an estimated 12 billion or more poultry, including four billion ducks and geese. Many domestic birds, especially ducks and geese, are raised in backyards and open ranges under poor biosecurity conditions and live in high-density groups close to large human populations. Since these birds may serve as important amplifying hosts of WNV, control of WNV circulation in birds is important for public health. For economic reasons, poultry farmers are generally unwilling to pay additional vaccine and labor costs for vaccination solely against WNV. As NDV is one of the most lethal and economically important pathogens for poultry (at least chickens and geese), farmers use live vaccines, such as LaSota strain, to protect against infection. In our study, most domestic poultry showed NDV HI antibody titters higher than 4 log2 after vaccination, irrespective of the delivery route (Figs. 5b, 6b and 7b). A HI antibody titer higher than 3 log2 is usually sufficient to protect poultry from lethal challenge of NDV. In this scenario, farmers will not need to pay additional vaccine and labor costs to protect against WNV infection by using rLa-WNV-PrM/E instead of NDV live vaccine for routine vaccination. Moreover, oral and intranasal immunization routes are more convenient than intramuscular immunization for poultry as well as water fowl and migratory and resident wild birds.
In summary, rLa-WNV-PrM/E vaccination in susceptible animals is important for protection against WNV infection. Our findings demonstrate that rLa-WNV-PrM/E delivered via multiple immunization routes is immunogenic in both mammals and poultry.
Murray KO, Mertens E, Despres P. West Nile virus and its emergence in the United States of America. Vet Res. 2010;41:67.
Langevin SA, Bunning M, Davis B, Komar N. Experimental infection of chickens as candidate sentinels for West Nile virus. Emerg Infect Dis. 2001;7:726–9.
Himsworth CG, Gurney KE, Neimanis AS, Wobeser GA, Leighton FA. An outbreak of West Nile virus infection in captive lesser scaup (Aythya affinis) ducklings. Avian Dis. 2009;53:129–34.
Banet-Noach C, Simanov L, Malkinson M. Direct (non-vector) transmission of West Nile virus in geese. Avian Pathol. 2003;32:489–94.
Ng T, Hathaway D, Jennings N, Champ D, Chiang YW, Chu HJ. Equine vaccine for West Nile virus. Dev Biol (Basel). 2003;114:221–7.
El Garch H, Minke JM, Rehder J, Richard S, Edlund Toulemonde C, Dinic S, Andreoni C, Audonnet JC, Nordgren R, Juillard V. A West Nile virus (WNV) recombinant canarypox virus vaccine elicits WNV-specific neutralizing antibodies and cell-mediated immune responses in the horse. Vet Immunol Immunopathol. 2008;123:230–9.
Bowen RA, Bosco-Lauth A, Syvrud K, Thomas A, Meinert TR, Ludlow DR, Cook C, Salt J, Ons E. Protection of horses from West Nile virus Lineage 2 challenge following immunization with a whole, inactivated WNV lineage 1 vaccine. Vaccine. 2014;32:5455–9.
Minke JM, Siger L, Cupillard L, Powers B, Bakonyi T, Boyum S, Nowotny N, Bowen R. Protection provided by a recombinant ALVAC((R))-WNV vaccine expressing the prM/E genes of a lineage 1 strain of WNV against a virulent challenge with a lineage 2 strain. Vaccine. 2011;29:4608–12.
Bukreyev A, Skiadopoulos MH, Murphy BR, Collins PL. Nonsegmented negative-strand viruses as vaccine vectors. J Virol. 2006;80:10293–306.
Bukreyev A, Collins PL. Newcastle disease virus as a vaccine vector for humans. Curr Opin Mol Ther. 2008;10:46–55.
Ge J, Deng G, Wen Z, Tian G, Wang Y, Shi J, Wang X, Li Y, Hu S, Jiang Y, et al. Newcastle disease virus-based live attenuated vaccine completely protects chickens and mice from lethal challenge of homologous and heterologous H5N1 avian influenza viruses. J Virol. 2007;81:150–8.
Ge J, Tian G, Zeng X, Jiang Y, Chen H, Bua Z. Generation and evaluation of a Newcastle disease virus-based H9 avian influenza live vaccine. Avian Dis. 2010;54:294–6.
DiNapoli JM, Yang L, Samal SK, Murphy BR, Collins PL, Bukreyev A. Respiratory tract immunization of non-human primates with a Newcastle disease virus-vectored vaccine candidate against Ebola virus elicits a neutralizing antibody response. Vaccine. 2010;29:17–25.
DiNapoli JM, Nayak B, Yang L, Finneyfrock BW, Cook A, Andersen H, Torres-Velez F, Murphy BR, Samal SK, Collins PL, Bukreyev A. Newcastle disease virus-vectored vaccines expressing the hemagglutinin or neuraminidase protein of H5N1 highly pathogenic avian influenza virus protect against virus challenge in monkeys. J Virol. 2010;84:1489–503.
Kortekaas J, de Boer SM, Kant J, Vloet RP, Antonis AF, Moormann RJ. Rift Valley fever virus immunity provided by a paramyxovirus vaccine vector. Vaccine. 2010;28:4394–401.
Khattar SK, Collins PL, Samal SK. Immunization of cattle with recombinant Newcastle disease virus expressing bovine herpesvirus-1 (BHV-1) glycoprotein D induces mucosal and serum antibody responses and provides partial protection against BHV-1. Vaccine. 2010;28:3159–70.
Liu Q, Mena I, Ma J, Bawa B, Krammer F, Lyoo YS, Lang Y, Morozov I, Mahardika GN, Ma W, et al. Newcastle Disease Virus-Vectored H7 and H5 Live Vaccines Protect Chickens from Challenge with H7N9 or H5N1 Avian Influenza Viruses. J Virol. 2015;89:7401–8.
Zhao W, Spatz S, Zhang Z, Wen G, Garcia M, Zsak L, Zsak L, Yu Q. Newcastle disease virus (NDV) recombinants expressing infectious laryngotracheitis virus (ILTV) glycoproteins gB and gD protect chickens against ILTV and NDV challenges. J Virol. 2014;88:8397–406.
Marquardt WW, Snyder DB, Savage PK, Kadavil SK, Yancey FS. Antibody response to Newcastle disease virus given by two different routes as measured by ELISA and hemagglutination-inhibition test and associated tracheal immunity. Avian Dis. 1985;29:71–9.
Jayawardane GW, Spradbrow PB. Mucosal immunity in chickens vaccinated with the V4 strain of Newcastle disease virus. Vet Microbiol. 1995;46:69–77.
Wambura PN, Wilson C. Protective antibody response following oral vaccination of feral pigeons (Columba livia) with Newcastle disease vaccine (strain I-2) coated on oiled rice. Vet Res Commun. 2009;33:921–6.
Wengler G, Wengler G. Cell-associated West Nile flavivirus is covered with E + pre-M protein heterodimers which are destroyed and reorganized by proteolytic cleavage during virus release. J Virol. 1989;63:2521–6.
Sanchez MD, Pierson TC, McAllister D, Hanna SL, Puffer BA, Valentine LE, Murtadha MM, Hoxie JA, Doms RW. Characterization of neutralizing antibodies to West Nile virus. Virology. 2005;336:70–82.
Guo LP, Huo H, Wang XL, Bu ZG, Hua RH. Generation and characterization of a monoclonal antibody against prM protein of West Nile virus. Monoclon Antib Immunodiagn Immunother. 2014;33:438–43.
OIE. Manual of Diagnostic Tests and Vaccines for Terrestrial Animals 2011. Paris: Office International des Epizooties; 2011.
Kong D, Wen Z, Su H, Ge J, Chen W, Wang X, Wu C, Yang C, Chen H, Bu Z. Newcastle disease virus-vectored Nipah encephalitis vaccines induce B and T cell responses in mice and long-lasting neutralizing antibodies in pigs. Virol. 2012;432:327–35.
Wen Z, Ye L, Gao Y, Pan L, Dong K, Bu Z, Compans RW, Yang C. Immunization by influenza virus-like particles protects aged mice against lethal influenza virus challenge. Antiviral Res. 2009;84:215–24.
Ye L, Bu Z, Vzorov A, Taylor D, Compans RW, Yang C. Surface stability and immunogenicity of the human immunodeficiency virus envelope glycoprotein: role of the cytoplasmic domain. J Virol. 2004;78:13409–19.
Brien JD, Uhrlaub JL, Nikolich-Zugich J. West Nile virus-specific CD4 T cells exhibit direct antiviral cytokine secretion and cytotoxicity and are sufficient for antiviral protection. J Immunol. 2008;181:8568–75.
Brien JD, Uhrlaub JL, Nikolich-Zugich J. Protective capacity and epitope specificity of CD8(+) T cells responding to lethal West Nile virus infection. Eur J Immunol. 2007;37:1855–63.
Diamond MS, Shrestha B, Marri A, Mahan D, Engle M. B cells and antibody play critical roles in the immediate defense of disseminated infection by West Nile encephalitis virus. J Virol. 2003;77:2578–86.
Ferguson M, Kurane I, Wimalaratne O, Shin J, Wood D, group WHOic. WHO informal consultation on the scientific basis of specifications for production and control of inactivated Japanese encephalitis vaccines for human use, Geneva, Switzerland, 1–2 June 2006. Vaccine. 2007;25:5233–43.
Muraki Y, Fujita T, Matsuura M, Fuke I, Manabe S, Ishikawa T, Okuno Y, Morita K. The efficacy of inactivated West Nile vaccine (WN-VAX) in mice and monkeys. J Virol. 2015;12:54.
Sitati EM, Diamond MS. CD4+ T-cell responses are required for clearance of West Nile virus from the central nervous system. J Virol. 2006;80:12060–9.
Shrestha B, Diamond MS. Role of CD8+ T cells in control of West Nile virus infection. J Virol. 2004;78:8312–21.
Mathew A, Terajima M, West K, Green S, Rothman AL, Ennis FA, Kennedy JS. Identification of murine poxvirus-specific CD8+ CTL epitopes with distinct functional profiles. J Immunol. 2005;174:2212–9.
Song GY, Srivastava T, Ishizaki H, Lacey SF, Diamond DJ, Ellenhorn JD. Recombinant modified vaccinia virus ankara (MVA) expressing wild-type human p53 induces specific antitumor CTL expansion. Cancer Invest. 2011;29:501–10.
Wang Z, La Rosa C, Lacey SF, Maas R, Mekhoubad S, Britt WJ, Diamond DJ. Attenuated poxvirus expressing three immunodominant CMV antigens as a vaccine strategy for CMV infection. J Clin Virol. 2006;35:324–31.
Samina I, Havenga M, Koudstaal W, Khinich Y, Koldijk M, Malkinson M, Simanov M, Perl S, Gijsbers L, Weverling GJ et al. Safety and efficacy in geese of a PER.C6-based inactivated West Nile virus vaccine. Vaccine. 2007;25:8338–45.
Lim CK, Takasaki T, Kotaki A, Kurane I. Vero cell-derived inactivated West Nile (WN) vaccine induces protective immunity against lethal WN virus infection in mice and shows a facilitated neutralizing antibody response in mice previously immunized with Japanese encephalitis vaccine. Virol. 2008;374:60–70.
Davis BS, Chang GJ, Cropp B, Roehrig JT, Martin DA, Mitchell CJ, Bowen R, Bunning ML. West Nile virus recombinant DNA vaccine protects mouse and horse from virus challenge and expresses in vitro a noninfectious recombinant antigen that can be used in enzyme-linked immunosorbent assays. J Virol. 2001;75:4040–7.
Seregin A, Nistler R, Borisevich V, Yamshchikov G, Chaporgina E, Kwok CW, Yamshchikov V. Immunogenicity of West Nile virus infectious DNA and its noninfectious derivatives. Virol. 2006;356:115–25.
Chang DC, Liu WJ, Anraku I, Clark DC, Pollitt CC, Suhrbier A, Hall RA, Khromykh AA. Single-round infectious particles enhance immunogenicity of a DNA vaccine against West Nile virus. Nat Biotechnol. 2008;26:571–7.
Karaca K, Bowen R, Austgen LE, Teehee M, Siger L, Grosenbaugh D, Loosemore L, Audonnet JC, Nordgren R, Minke JM. Recombinant canarypox vectored West Nile virus (WNV) vaccine protects dogs and cats against a mosquito WNV challenge. Vaccine. 2005;23:3808–13.
Monath TP, Liu J, Kanesa-Thasan N, Myers GA, Nichols R, Deary A, McCarthy K, Johnson C, Ermak T, Shin S, et al. A live, attenuated recombinant West Nile virus vaccine. Proc Natl Acad Sci U S A. 2006;103:6694–9.
Long MT, Gibbs EP, Mellencamp MW, Bowen RA, Seino KK, Zhang S, Beachboard SE, Humphrey PP. Efficacy, duration, and onset of immunogenicity of a West Nile virus vaccine, live Flavivirus chimera, in horses with a clinical disease challenge model. Equine Vet J. 2007;39:491–7.
Seino KK, Long MT, Gibbs EP, Bowen RA, Beachboard SE, Humphrey PP, Dixon MA, Bourgeois MA. Comparative efficacies of three commercially available vaccines against West Nile Virus (WNV) in a short-duration challenge trial involving an equine WNV encephalitis model. Clin Vaccine Immunol. 2007;14:1465–71.
Minke JM, Siger L, Karaca K, Austgen L, Gordy P, Bowen R, Renshaw RW, Loosmore S, Audonnet JC, Nordgren B. Recombinant canarypoxvirus vaccine carrying the prM/E genes of West Nile virus protects horses against a West Nile virus-mosquito challenge. Arch Virol Suppl. 2004;18:221–30.
Blackmore CG, Stark LM, Jeter WC, Oliveri RL, Brooks RG, Conti LA, Wiersma ST. Surveillance results from the first West Nile virus transmission season in Florida, 2001. Am J Trop Med Hyg. 2003;69:141–50.
Chaskopoulou A, Dovas C, Chaintoutis S, Bouzalas I, Ara G, Papanastassopoulou M: Evidence of enzootic circulation of West Nile virus (Nea Santa-Greece-2010, lineage 2), Greece, May to July 2011. Euro Surveill. 2011: 16.
Fall AG, Diaite A, Seck MT, Bouyer J, Lefrancois T, Vachiery N, Aprelon R, Faye O, Konate L, Lancelot R. West Nile virus transmission in sentinel chickens and potential mosquito vectors, Senegal River Delta, 2008–2009. Int J Environ Res Public Health. 2013;10:4718–27.
Komar N. West Nile virus surveillance using sentinel birds. Ann N Y Acad Sci. 2001;951:58–73.
Meece JK, Kronenwetter-Koepel TA, Vandermause MF, Reed KD. West Nile virus infection in commercial waterfowl operation, Wisconsin. Emerg Infect Dis. 2006;12:1451–3.
Wojnarowicz C, Olkowski A, Schwean-Lardner K. First Canadian outbreak of West Nile virus disease in farmed domestic ducks in Saskatchewan. Can Vet J. 2007;48:1270–1.
Hofmeister E, Porter RE, Franson JC. Experimental susceptibility of wood ducks (Aix sponsa) for West Nile virus. J Wildl Dis. 2015;51:411–8.
Guy JS, Malkinson M. Arbovirus infections. In: Saif YM, editor. Diseases of Poultry. 11th ed. Ames: Iowa State University Press; 2003. p. 388–98.
Austin RJ, Whiting TL, Anderson RA, Drebot MA. An outbreak of West Nile virus-associated disease in domestic geese (Anser anser domesticus) upon initial introduction to a geographic region, with evidence of bird to bird transmission. Can Vet J. 2004;45:117–23.
Samina I, Khinich Y, Simanov M, Malkinson M. An inactivated West Nile virus vaccine for domestic geese-efficacy study and a summary of 4 years of field application. Vaccine. 2005;23:4955–8.
Sa ESM, Ellis A, Karaca K, Minke J, Nordgren R, Wu S, Swayne DE. Domestic goose as a model for West Nile virus vaccine efficacy. Vaccine. 2013;31:1045–50.
Jarvi SI, Lieberman MM, Hofmeister E, Nerurkar VR, Wong T, Weeks-Levy C. Protective efficacy of a recombinant subunit West Nile virus vaccine in domestic geese (Anser anser). Vaccine. 2008;26:5338–44.
Lu Z, Fu SH, Cao L, Tang CJ, Zhang S, Li ZX, Tusong M, Yao XH, Zhang HL, Wang PY, et al. Human infection with West Nile Virus, Xinjiang, China, 2011. Emerg Infect Dis. 2014;20:1421–3.
Li XL, Fu SH, Liu WB, Wang HY, Lu Z, Tong SX, Li ZX, Nasci RS, Kosoy O, Cui Y, Liang GD. West nile virus infection in Xinjiang, China. Vector Borne Zoonotic Dis. 2013;13:131–3.
This study was supported by The National Key Technology R&D Program (2013BAD12B05) and National High-Tech Research and Development Program of China (2011AA10A212).
ZB and ZW designed and oversaw the experiments and wrote the manuscript. JY and JW rescued and characterized the recombinant virus. JW and JY determined the mice IgG and HI antibodies, ZW carried out T cell response assays. JW, JG, XW, RH, RL, YS, ES and DW carried out the horse and poultry immunization assay. XL and CQ determined the mice neutralizing antibodies. All authors have read and approved the submitted manuscript.
The authors declare that they have no competing interests.
Ethics approval and consent to participate
The present study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the Ministry of Science and Technology of the People’s Republic of China. The protocol was approved by the Animal Research Ethics Committee of Harbin Veterinary Research Institute, Chinese Academy of Agricultural Sciences.
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Wang, J., Yang, J., Ge, J. et al. Newcastle disease virus-vectored West Nile fever vaccine is immunogenic in mammals and poultry. Virol J 13, 109 (2016). https://doi.org/10.1186/s12985-016-0568-5
- West Nile fever
- Newcastle disease virus vectored vaccine
- Neutralizing antibody
- T cell response