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Vaccination with virus-like particles protects mice from lethal infection of Rift Valley Fever Virus
Virology 385 (2009) 409–415
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Virology j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / y v i r o
Vaccination with virus-like particles protects mice from lethal infection of Rift Valley Fever Virus Jonas Näslund a,b,c,1, Nina Lagerqvist a,b,d,1, Matthias Habjan e, Åke Lundkvist d, Magnus Evander c, Clas Ahlm b, Friedemann Weber e,⁎, Göran Bucht a,f,⁎ a
Swedish Defence Research Agency, Department of CBRN Defence and Security, SE-901 82 Umeå, Sweden Division of Infectious Diseases, Department of Clinical Microbiology Umeå University, SE-901 85 Umeå, Sweden Division of Virology, Department of Clinical Microbiology Umeå University, SE-901 85 Umeå, Sweden d Swedish Institute for Infectious Disease Control, SE-171 82 Solna, Sweden e Department of Virology, University of Freiburg, D-79008 Freiburg, Germany f National Environment Agency, Environmental Health Institute, 11 Biopolis Way, 06-05/08, Helios Block, Singapore 138667, Singapore b c
a r t i c l e
i n f o
Article history: Received 24 September 2008 Returned to author for revision 30 October 2008 Accepted 8 December 2008 Available online 20 January 2009 Keywords: Rift Valley Fever Mouse model Virus-like particle Vaccine Challenge
a b s t r a c t Rift Valley Fever virus (RVFV) regularly accounts for severe and often lethal outbreaks among livestock and humans in Africa. Safe and effective veterinarian and human vaccines are highly needed. We present evidence that administration of RVF virus-like particles (VLPs) induces protective immunity in mice. In an accompanying paper, (Habjan, M., Penski, N., Wagner, V., Spiegel, M., Överby, A.K., Kochs, G., Huiskonen, J., Weber, F., 2009. Efficient production of Rift Valley fever virus-like particles: the antiviral protein MxA can inhibit primary transcription of Bunyaviruses. Virology 385, 400–408) we report the production of these VLPs in mammalian cells. After three subsequent immunizations with 1 × 106 VLPs/dose, high titers of virusneutralizing antibodies were detected; 11 out of 12 mice were protected from challenge and only 1 out of 12 mice survived infection in the control groups. VLP vaccination efficiently suppressed replication of the challenge virus, whereas in the control animals high RNA levels and increasing antibody titers against the nucleocapsid protein indicated extensive viral replication. Our study demonstrates that the RVF VLPs are highly immunogenic and confer protection against RVFV infection in mice. In the test groups, the vaccinated mice did not exhibit any side effects, and the lack of anti-nucleocapsid protein antibodies serologically distinguished vaccinated animals from experimentally infected animals. © 2008 Elsevier Inc. All rights reserved.
Introduction Rift Valley Fever (RVF) is an emerging infectious disease and considered as one of the most important viral zoonoses in Africa. The disease was restricted to the sub-Saharan region of Africa until the middle of the 1970s. Since then, RVF has become endemic throughout Africa and more recently RVF has been found also on Arabian Peninsula (Balkhy and Memish, 2003; Gerdes, 2004). Epizootics of RVF are characterized by increasing numbers of abortions and high neonatal mortality. Among adult animals, the clinical manifestations range from asymptomatic to more severe conditions like hepatitis and death, although the progression and ⁎ Corresponding authors. F. Weber is to be contacted at Fax: +49 761 203 6634. G. Bucht, Fax: +65 6777 8029. E-mail addresses: [email protected] (J. Näslund), [email protected] (N. Lagerqvist), [email protected] (M. Habjan), [email protected] (Å. Lundkvist), [email protected] (M. Evander), [email protected] (C. Ahlm), [email protected] (F. Weber), [email protected] (G. Bucht). 1 These authors contributed equally to this work. 0042-6822/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.virol.2008.12.012
severity of the disease varies considerably between different animal species (Elliott, 1996). In humans, RVF is considered as a temporarily incapacitating illness with fever, myalgia, and headache. However, more severe clinical symptoms—such as encephalitis, ocular disease, hepatitis, and hemorrhagic fever—have been observed (Al-Hazmi et al., 2003; Siam et al., 1980). Presently, treatments are symptomatic combined with an occasional need of intensive care. The RVF virus (RVFV), a member of the Phlebovirus genera in the Bunyaviridae family, is transmitted by several different mosquito species and has an unusual broad range of susceptible animal hosts (Flick and Bouloy, 2005). As for other members of this family of viruses, the genetic information is divided into three negative stranded RNA segments: (S)mall, (M)edium and (L)arge. The L-segment encodes for the RNA polymerase, and the M- and S-segments encode for two glycoproteins (GN/GC), the 78 kDa protein and the nucleocapsid (N) protein, respectively. Furthermore, the RVFV genome encodes for two non-structural proteins: NSs and NSm of the S and M segment, correspondingly (Elliott, 1996). The NSs protein is involved in disruption of the host antiviral response (Billecocq et al., 2004; Le May et al., 2004) and the NSm protein is reported to suppress apoptotic pathways in host cells after infection (Won et al., 2007).
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The recent geographical expansion of RVF, combined with the ability of the virus to maintain in newly conquered areas due to the wide range of mosquito vectors and susceptible hosts, imply that novel therapeutics and vaccines are necessary. Currently, there are no registered vaccines for human use. However, a formalin inactivated virus vaccine for humans has been developed (Kark et al., 1982; Pittman et al., 1999), although the distribution has been limited to workers at high-risk occupations. The live attenuated Smithburn neurotropic strain (Smithburn, 1949) and formalin inactivated virus vaccines produced in South Africa and Egypt (Metwally, 2008) are approved for veterinarian use. The Smithburn vaccine is highly immunogenic, but is teratogenic in pregnant cattle and sheep (Botros et al., 2006; Coetzer and Barnard, 1977). The formalin inactivated vaccines are safe, but less immunogenic (Lubroth et al., 2007). Other vaccine candidates under evaluation are Clone 13 and MP12. Both have shown promising results in animal trials. Clone 13 is a natural attenuated isolate from a benign human case in the Central African Republic (Muller et al., 1995), and MP12 is a chemically attenuated virus derived from the Egyptian wild-type isolate ZH548 (Caplen et al., 1985; Vialat et al., 1997). The immunogenicity and pathogenicity of these candidates have been evaluated in various animal species (Muller et al., 1995; Morrill et al., 1997). The most extensively tested, MP12, induced foetal malformations during the first trimester (Hunter et al., 2002). More recently, reverse genetic systems have been established for RVFV that allow genetic engineering of the virus genomes and production of recombinant virus particles (Billecocq et al., 2008; Gerrard et al., 2007; Habjan et al., 2008; Ikegami et al., 2006). Such techniques have been used to produce attenuated RVFV particles which have displayed promising results as a vaccine in a rodent model (Bird et al., 2008). However, inherent problems of live vaccines are the expensive production due to high biosafety precautions and the risk of side effects such as teratogenicity and infections in immuno-suppressed individuals. Moreover, replicating RVFV vaccines could theoretically be spread by mosquitoes. Another approach for vaccine development is to use virus-like particles (VLPs) that are formed when the structural (envelope and/or nucleocapsid) proteins self-assemble to replication-deficient particles. In comparison to component vaccines based on recombinant or subunit proteins, VLPs are in general more immunogenic and more natural in presenting conformational epitopes, and hence, more likely to induce neutralizing antibodies (Grgacic and Anderson, 2006). In an accompanying paper, we describe the production of RVF VLPs in mammalian cells by expressing the viral structural genes along with a Renilla luciferase reporter minigenome (Habjan et al., 2009). These VLPs have the size, morphology, and protein composition resembling authentic RVFV particles. They contain nucleocapsids consisting of N and the reporter RNA minigenome and display the envelope (GN and GC) proteins at their surface. Moreover, the VLPs can enter target cells and carry out transcription of the reporter gene, but they cannot replicate or produce progeny. They can be neutralized by RVFVspecific antisera. Here, we report the use of these VLPs as a vaccine against RVF. The safety and immunogenicity were evaluated in mice and the results indicate that immunization with VLPs confers protection against challenge with lethal doses of RVFV. These VLPs offer a promising alternative to attenuated virus mutants as an efficient and safe vaccine against RVF. Results The RVF VLPs are immunogenic and induce virus-neutralizing antibodies Thirty mice were immunized three times with two weeks intermission with either 1 × 105 VLPs/dose, 1 × 106 VLPs/dose, sterile PBS or supernatant of cells transfected with all VLP constructs except the glycoprotein plasmid (Habjan et al., 2009). The two latter groups
constituted the negative controls. No clinical manifestations derived from the vaccine were observed in any of these mice. After three immunizations, the sera were analyzed for the presence of anti-RVFV specific antibodies. Immunofluorescense analysis (IFA) performed on virus-infected mammalian cells showed that all VLP-vaccinated mice sero-converted and antibodies were detectable at serum dilutions of up to 1:2700 (Fig. 1A). To analyze the antibody response in more detail, IFA was performed on mammalian cells transfected with either cDNA encoding the N protein or the GN/GC proteins (data not shown). VLP-vaccination induced antibodies towards the GN/GC protein among all mice, with titers ranging from 300 up to 900. On the other hand, no detectable N protein specific antibodies (total Ig) were observed by IFA after three subsequent VLP-vaccinations (data not shown). The virus neutralizing antibody titers were determined by a plaque reduction neutralization test (PRNT) (Fig. 1B). In the group of mice administered with 1 × 105 VLPs/dose, three animals (3/6) showed low titers, equal to or below 50, the remaining three animals were found to have neutralizing titers of 250. When the high-dose group administered with 1 × 106 VLPs/dose were analyzed, high titers of neutralizing antibodies were found; in this dose group, the titer values ranged from 250 to 1250 (Fig. 1B). However, one animal of this group did not respond to the vaccination, and the neutralizing titer was below 50. No sero-conversion was detected among the control mice. No apparent differences in the immune responses, viral replication, clinical signs, or survival proportions were observed between mice from the two control groups having received either sterile PBS or a cell
Fig. 1. Sero-conversion in mice after VLP vaccination determined by immunofluorescence assay and plaque reduction neutralization tests. Mice sera were collected after three subsequent VLP immunizations and analyzed for panel A antibody reactivity towards RVFV-infected mammalian cells and panel B titer of virus neutralizing antibodies. Two dose groups are represented: mice vaccinated with 1 × 105 or 1 × 106 VLPs/dose. No sero-conversion was observed among the control mice and are therefore excluded from these graphs. The antibody titer was determined as the reciprocal of the highest positive serum dilution of (panel A) three-fold and (panel B) five-fold serially diluted sera, respectively.
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supernatant (see Materials and methods). Therefore, these two groups were consolidated and represent the control. Vaccination with RVF VLPs protects mice in challenge The mice were challenged two weeks after the third and final vaccination with a lethal dose (2.4 × 104 PFU) of RVFV. During the entire challenge experiment, the mice were monitored twice a day for signs of infection. The survival proportions among each vaccination group are shown in Fig. 2. Three out of six mice survived challenge in the low-dose group, immunized with 1 × 105 VLPs/dose. Mild clinical signs were observed among two of the three surviving animals around day 7 p.i., followed by complete recovery one to two days thereafter. Eleven out of twelve mice vaccinated with 1 × 106 VLPs/dose survived and no clinical manifestations were observed among the surviving animals during the experimental period. Interestingly, three mice vaccinated with the lower dose of VLP and one mouse of the high dose group showing the lowest titers of neutralizing antibodies, as shown in Fig. 1B, had to be sacrificed at days 9 and 10 due to severe clinical signs. One animal of twelve in the control group survived challenge without displaying any clinical manifestations. The remaining mice in the control group displayed severe clinical signs, in some cases as early as 4 to 5 days p.i., and two control animals developed clinical signs as late as 12 days p.i. Survival proportion was significantly higher in mice vaccinated with 1 × 106 VLPs/dose than in the control group (Chisquare test, p = 0.00004). However, the survival proportion was not significant for the mice in the low-dose group (Fisher exact test, p = 0.0833). VLP vaccination suppresses sero-conversion against the N protein after RVFV challenge Since vaccination with the RVF VLPs used in these experiments did not induce antibodies directed towards the N protein (pre-challenge sera obtained day 0 p.i., Fig. 3B, C), any detection of anti-N specific antibodies after challenge is an indication of viral replication in the animal hosts. When analyzing sera from the control animals after challenge (Fig. 3A), anti-N antibodies were clearly observed by ELISA from days 4 and 5 p.i., followed by increasing and highly individual titers. However, the VLP-vaccinated mice displayed no or a moderate rise in the antibody levels (Figs. 3B, C), and the curves are clearly separated from those of the control animals. The animals in the lowdose group with slightly elevated titer levels around day 5 p.i. and the high-dose animals with constantly low antibody titers indicate Fig. 3. Antibody response towards the N protein in vaccinated and control animals after challenge with RVFV. Serum samples were collected at different days after challenge and analysed by ELISA for the presence of specific antibodies directed against the N protein. The graphs represent antibody titers in serum samples of panel A control animals, panel B mice vaccinated with 1 × 105 VLPs/dose and panel C mice vaccinated with 1 × 106 VLPs/dose. The curves show the mean value from two separate measurements and the error bars represent the standard deviation. Incomplete curves correspond to samples taken from mice that were euthanized early due to a moribund condition.
negligible virus replication in the vaccinated animals after challenge. Only small variations in antibody levels were detected after day 12 p.i. (data not shown). VLP immunization inhibits RVFV replication in mice
Fig. 2. Percent survival of VLP immunized mice after RVFV challenge. The mice were vaccinated with either 1 × 105 or 1 × 106 VLPs/dose then challenged with 2.4 × 104 PFU of RVFV strain ZH548. Mice immunized with a cell suspension or PBS constitute the control group (Ctrl).
To further verify if vaccination with VLPs suppresses replication of the wild-type RVFV after challenge, viral RNA copy numbers in blood were quantified using a SYBR-green based qRT-PCR assay (Fig. 4). When analyzing serum samples of the control and the low-dose mice (Figs. 4A, B), a great divergence in the detected levels of RNA was observed between individual mice. A large proportion of the control
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Fig. 4. Quantification of RVFV RNA in blood samples of VLP-vaccinated and control animals after challenge. Blood samples were collected before infection (day 0) and on days three, five, seven, ten, and twelve after infection and analyzed for the presence of viral RNA by qRT-PCR. The graphs represent panel A control animals, panel B mice vaccinated with 1 × 105 VLPs/dose, and panel C mice vaccinated with 1 × 106 VLPs/dose. The curves illustrate RNA copies/ml blood and represent the mean value of two replicates. The error bars show the standard deviation between measurements.
mice showed high levels of viral RNA 3 days p.i (Fig. 4A), while animals of the low-dose group reached their maximum viral load 2 days later (Fig. 4B). Animals of the high-dose group also displayed a delayed onset of viral replication (Fig. 4C) and only low viral RNA levels, in comparison to the other groups, were found (Fig. 4). In conclusion, the RVF VLPs were found to be safe and highly immunogenic in vivo and protected mice from lethal challenge by RVFV. Discussion RVF is an emerging infectious disease characterized by high mortality rates among foetuses, neonates, and juvenile livestock and causes a severe febrile illness also in man. To prevent RVF epidemics and the concomitant serious socio-economic consequences, new and improved vaccine strategies are of utmost importance.
Recently, VLPs based on the RVFV envelope proteins have been developed (Habjan et al., 2009), and herein we present the use of these RVF VLPs as a vaccine in a mouse model. These VLPs, when analyzed by electron microscopy, have a similar morphology as the authentic RVFV particles. Moreover, these VLPs display correctly processed viral glycoproteins in their envelope, proteins that are recognized by neutralizing antisera and contain transcriptionally active minigenome-nucleocapsids, but are unable to replicate and produce progeny (Habjan et al., 2009). In general, VLPs are reported to have similar properties as the virulent strains of the corresponding agents since the predominant structural antigens are displayed to the immune system in a native conformation (Grgacic and Anderson, 2006). The capability of VLPs to confer protective immunity has previously been demonstrated for a number of viruses. VLPs assembled from recombinant human papilloma virus capsid proteins are used as a vaccine against cervical cancer (Harper et al., 2006; Kirnbauer et al., 1992; Mao et al., 2006; Villa et al., 2005; Zhou et al., 1991) and VLPs of the small surface antigen of hepatitis B virus (André and Safary, 1987; McAleer et al., 1984) are also successfully used. In addition, several preclinical studies have been performed targeting, e.g., influenza (Latham and Galarza, 2001; Pushko et al., 2005), Rotavirus (Bertolotti-Ciarlet et al., 2003; Crawford et al., 1994; Vieira et al., 2005), and filovirus infections (Swenson et al., 2005; Warfield et al., 2004). Antibody-mediated virus neutralization constitutes an essential part in the control of infectious agents (Reading and Dimmock, 2007). In the present study, virus neutralization titers of 250 or more were found to be protective and associated with survival after challenge. All mice but one obtained serum neutralizing titers of 250, or more, after vaccination with the high VLP dose. Similar titers were only found for half of the animals vaccinated with the lower dose of VLPs. An unexpected low neutralization titer was found in one animal of the high-dose group and was probably the cause for the serious clinical signs that later lead to euthanization of this animal after challenge. Surprisingly, two of the surviving animals from the low-dose group displayed mild clinical signs despite the fact that they acquired neutralizing titers of 250. Three mice of the low-dose group and one animal of the high dose-group with the lowest neutralizing titers (equal to or below 50) had to be sacrificed at days 9 and 10 due to severe clinical manifestations. In the control group, the majority of the animals developed severe clinical signs and high viremia early after challenge and a large proportion (92%) of the animals succumbed to infection. One control animal did not display any clinical signs of infection, although high viral load and anti-N antibodies indicated extensive RVFV replication. There was a strong association between the administered VLP dose, neutralizing antibody titers and the survival rate. Importantly, a significant protection (92%) was observed among the animals administered the high VLP dose in contrast to the animals in the low-dose group where only 50% survived infection. The surviving high-dose animals were also clearly distinguished from both the control and the low-dose animals in the context of viral RNA levels and anti-N antibody titers detected after infection. Interestingly, no mouse displayed detectable levels of anti-N specific antibodies after VLP vaccination. RVF VLPs do not replicate and the N proteins present in the interior of the VLP were apparently not enough to elicit an immune response against the N protein. However, all mice in this study sero-converted towards the glycoproteins after vaccination with the RVF VLPs. The N protein of RVFV is a major antigen in the early serological response to infection and is particularly useful during clinical diagnostics and surveillance. Due to the zoonotic nature of RVF, surveillance is of major importance in the international trade of animals and animal-related products. Marker vaccines make it possible to differentiate infected from vaccinated individuals with specific diagnostic tests, valuable tools in the framework of control programs (van Oirschot, 1999). To use the benefits of marker vaccines fully, it is essential to use easy and
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assessable diagnostic tests. Immuno-fluorescence performed on infected or transfected cells, recombinant N protein based ELISA and neutralization tests are techniques widely used in laboratories that specialize in RVFV (Bird et al., 2008; Jansen van Vuren et al., 2007; Näslund et al., 2008; Zaki et al., 2006). Since VLP vaccination did not induce an anti-N specific antibody response, these already established techniques make it easy to monitor and evaluate future VLP vaccination programs. Currently, available RVF vaccines are based on attenuated or inactivated virus preparations. Because several disadvantages have been reported when using these vaccines (Lubroth et al., 2007), attempts to develop new vaccines is a priority. At the moment, the most advanced and promising live RVFV vaccine candidate is the recently produced rZH501-ΔNSs:GFP-ΔNSm, which lacks two RVFV virulence factors, NSs and NSm (Bird et al., 2008). Because there is a complete deletion of these genes involved in virulence, it is highly unlikely that the virus can revert to full virulence. However, other inherent drawbacks of live attenuated vaccines are -the need for an elevated biosafety level facility for production and the risk of using attenuated viruses in immunocompromised individuals - still remain. When vaccines based on the VLP strategy are used, the safety concerns and the logistical disadvantages are circumvented. Because available RVFV vaccines are associated with deleterious effects or low immunogenicity, there is an imperative need for the development of alternative RVF vaccine strategies. We demonstrate that RVF VLPs are safe and confer protective immunity in mice with respect to a lethal dose of RVFV. Furthermore, vaccinated individuals could readily be distinguished from infected with N-specific diagnostic tests. These results clearly show the potential of using VLPs as a future vaccine candidate against RVF. Materials and methods Cells and virus strain The RVFV strain ZH548 was originally isolated from an uncomplicated human case that occurred during the Egyptian outbreak in 1977 (Meegan, 1979; Sall et al., 1999). Low passage ZH548 stocks were propagated and titrated in confluent monolayers of Vero E6 cells (ATCC® Number CRL-1586) before used for challenge. Vero E6 cells were cultivated in Dulbecco's modified Eagle's medium (DMEM) (GIBCO, Invitrogen, Carlsbad, CA) supplemented with 10% FCS, 10 mM HEPES, and 1 mM sodium pyruvate and antibiotics (100 U penicillin/ml, 100 μg/ml streptomycin). For immunological methods, BHK-21 cells (ATCC® Number, CCL-10) were maintained in Glasgow MEM (GIBCO, Invitrogen) and supplemented with 5% FCS, 1.3 g/l Tryptose (Difco™, Becton, Dickinson and Company, Sparks, MD), 10 mM HEPES, and 1 mM sodium pyruvate and antibiotics (100 U penicillin/ml, 100 μg/ml streptomycin). Human 293T (ATCC® Number CRL-11268) cells were cultivated in DMEM and supplemented with 10% FCS, 0.4% sodium hydrogen carbonate, 1 mM sodium pyruvate, 0.53 mg/ml glutamine, and antibiotics (100 U penicillin/ml, 100 μg/ml streptomycin). All cells were cultivated at 5% CO2 at 37 °C. VLP production and purification Subconfluent monolayers of 293T cells in 90 mm dishes were transfected with 3 μg of plasmids pI.18_RVFV_N, pI.18_RVFV_L, pI.18_RVFV_M, and pHH21_RVFV_vMRen (Habjan et al., 2009) using Nanofectin (PAA Laboratories, Linz, Austria) transfection reagent. Two days post transfection, supernatants were collected and clarified from cell debris by centrifugation (6000 ×g, 10 min at 4 °C). The particles were then concentrated by ultracentrifugation at 24,000 rpm at 4 °C for 2 h in a Beckman SW41Ti rotor. The pellet
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was dried for 5 min before being re-suspended in phosphatebuffered saline (PBS). To define the titer, VLPs expressing GFP were generated and concentrated in parallel to the Ren-VLPs which were used for the vaccinations. BSR-T7/5 cells expressing RVFV L and N (Habjan et al., 2009) were incubated for 1 h with ten-fold serial dilutions of supernatants containing GFP-VLPs. Twenty-four hours later the number of GFP-positive (i.e. VLP-infected) cells was determined by fluorescence microscopy and the titer of the GFPVLP preparation was calculated accordingly. The titer of the RenVLPs was estimated to be in the same range. Western blot analysis confirmed that the amounts of GFP-VLPs and Ren-VLPs were comparable. VLP immunization This study used 33 female C57BL/6 mice (Scanbur BK) 6– 7 weeks old. The mice were kept in cages under BSL-2 conditions during the whole vaccination routine with free access to food and water. Two VLP doses were administered, 1 × 105 VLPs/dose (n = 6) and 1 × 106 VLPs/dose (n = 12). To serve as negative controls, 12 mice were inoculated with either sterile PBS (n = 5) or with a cell supernatant (n = 7), similar to what was used in the production of the RVF VLPs. Two independent vaccinations were performed with comparable immunization schedules and different batches of VLPs, although the second round did not include the lower dose group. All animals were vaccinated intraperitoneally (i.p.) three times (two weeks intermission) with the VLPs diluted in sterile PBS to a final volume of 200 μl. RVFV challenge Two weeks after the third vaccination, a challenge dose containing 2.4 × 104 PFU of RVFV strain ZH548 were administered i. p. During these procedures, the mice were kept individually in micro-isolator pans within an animal isolator (Bell Isolation Systems Ltd, Scotland, UK). All manipulations with infected mice and/or samples involving RVFV were performed strictly under BSL-3 conditions. The animals were examined twice daily for visual signs of illness until the experiments were ended, 12–14 days post infection (p.i.). Clinical signs observed among mice after infection ranged from distress, ruffled fur, and weight loss to fatigue, neurological manifestations, and a “hunchback-like” posture. Mice found in a moribund condition were instantly euthanized. Blood samples were collected before each immunization and every second or third day p.i. To ensure a sterile milieu, non-infected mice (n = 3) were kept under the same conditions during the entire experiment. This project was approved by The Animal Research Ethics Committee of Umeå University, Sweden. Plaque reduction neutralization test (PRNT) Heat inactivated sera, collected two weeks after the third immunization, were five-fold serially diluted in sterile PBS and incubated with a virus suspension containing 30 PFU of ZH548 before adsorption to a confluent monolayer of BHK-21 cells. After a 90 min incubation period and subsequent washing, the cells were covered with an overlay containing 1% CMC (Carboxy-Methyl Cellulose, Aquacide II, Calbiochem®, California, USA) prepared in maintenance DMEM and incubated for six days at 37 °C/5% CO2. After fixation with 10% formaldehyde, the plaques were visualized by counterstaining with 1% crystal violet in water containing 20% ethanol and 0.7% NaCl and counted after destaining with tap water. Sera of non-infected or infected mice constituted the negative and positive controls, respectively. The neutralizing antibody titers were determined as the reciprocal of the highest serum dilution that reduced the number of plaques by 80%.
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Immuno-fluorescence assay (IFA) and enzyme-linked immuno-sorbent assay (ELISA) For IFA, BHK-21 cells were infected with RVFV (strain ZH548) at a multiplicity of infection (MOI) of 1 or transfected and grown on cover slips. Transfection was performed using the eukaryotic expression vector (pcDNA3.1, Invitrogen) containing cDNA encoding the N protein or the open reading frame of the GN/GC proteins and FuGene 6 transfection reagent (Roche Inc.) according to the manufacturer's instructions. The cells were fixed in 4% paraformaldehyde and permeabilized with 0.2% Saponin in PBS 36 h p.i. or 48 h post transfection. After incubation with three-fold serial dilutions of the serum samples, collected two weeks after the third vaccination, infected or transfected cells were labelled using a goat anti-mouse Alexa Fluor™ 488 conjugate total Ig or a FITC conjugated rabbit anti-mouse total Ig, respectively, according to the instructions of the manufacturers (Molecular Probes, Invitrogen). ELISA was performed as described earlier (Näslund et al., 2008) using recombinant N-protein (GenBank accession no. AF134534) (Sall et al., 1999) of RVFV and constructed and purified as previously described (Le May et al., 2005; Lindkvist et al., 2007). The sera used were collected at the day of infection (day 0) then three, five, seven, ten and twelve days p.i. The serum samples were prepared as four-fold serial dilutions, and the antibody titers were calculated as the inverse dilution at the intersection of the straight line and the x-axis. Serum samples from pre-bleedings and non-infected mice were present on each plate to determine unspecific reactivity. Quantitative real time reverse transcriptase PCR (qRT-PCR) The viral load was measured in blood, collected at the day of infection (day 0) and day three, five, seven, ten, and twelve p.i, using a qRT-PCR as previously described (Näslund et al., 2008). Briefly, RNA was extracted and purified using Trizol LS (Invitrogen) and RNeasy Mini Kit (Qiagen Inc.) in accordance to the manufacturer's instructions. cDNA were synthesized using SuperScript™ III RT according to manufacturer's instructions (Invitrogen). qRT-PCR was performed in 24 μl reactions using SYBR-green (Biotools, B&M Labs, Madrid, Spain) with primers targeting the S-segment of RVFV (available from the authors upon request). Statistical methods Survival proportions were compared using Chi-square test and Fischer exact test for values under five (Epi Info™, Version 3.5). Quantitative variables were based on measurements of a minimum of two independent experiments containing duplicate samples. Variables are expressed as means, and the error bars correspond to the standard deviation between the replicates. Acknowledgments Ms. Karin Wallgren is greatly acknowledged for her excellent technical support during the immunizations and bleedings of the mice. This study was supported by the Swedish Defence Agency, the Medical Faculty of Umeå University, grants from the County Council of Västerbotten, and by the grant We 2616/5-1 from the Deutsche Forschungsgemeinschaft. This project was also partially supported by grants from the Swedish Research Council (project 12177) and the European Community (contract no. QLK2-CT-2002-01358). References Al-Hazmi, M., Ayoola, E.A., Abdurahman, M., Banzal, S., Ashraf, J., El-Bushra, A., Hazmi, A., Abdullah, M., Abbo, H., Elamin, A., Al-Sammani, el.-T., Gadour, M., Menon, C., Hamza, M., Rahim, I., Hafez, M., Jambavalikar, M., Arishi, H., Aqeel, A., 2003. Epidemic Rift Valley fever in Saudi Arabia: a clinical study of severe illness in humans. Clin. Infect. Dis. 36, 245–252.
André, F., Safary, A., 1987. Summary of clinical findings on Engerix-B, a genetically engineered yeast derived hepatitis B vaccine. Postgrad. Med. J. 63 (Suppl 2), 169–177. Balkhy, H., Memish, Z., 2003. Rift Valley fever: an uninvited zoonosis in the Arabian peninsula. Int. J. Antimicrob. Agents 21, 153–157. Bertolotti-Ciarlet, A., Ciarlet, M., Crawford, S., Conner, M., Estes, M., 2003. Immunogenicity and protective efficacy of rotavirus 2/6-virus-like particles produced by a dual baculovirus expression vector and administered intramuscularly, intranasally, or orally to mice. Vaccine 21, 3885–3900. Billecocq, A., Spiegel, M., Vialat, P., Kohl, A., Weber, F., Bouloy, M., Haller, O., 2004. NSs protein of Rift Valley fever virus blocks interferon production by inhibiting host gene transcription. J. Virol. 78, 9798–9806. Billecocq, A., Gauliard, N., Le May, N., Elliott, R., Flick, R., Bouloy, M., 2008. RNA polymerase I-mediated expression of viral RNA for the rescue of infectious virulent and avirulent Rift Valley fever viruses. Virology 378, 377–384. Bird, B., Albariño, C., Hartman, A., Erickson, B., Ksiazek, T., Nichol, S., 2008. Rift valley fever virus lacking the NSs and NSm genes is highly attenuated, confers protective immunity from virulent virus challenge, and allows for differential identification of infected and vaccinated animals. J. Virol. 82, 2681–2691. Botros, B., Omar, A., Elian, K., Mohamed, G., Soliman, A., Salib, A., Salman, D., Saad, M., Earhart, K., 2006. Adverse response of non-indigenous cattle of European breeds to live attenuated Smithburn Rift Valley fever vaccine. J. Med. Virol. 78, 787–791. Caplen, H., Peters, C.J., Bishop, D., 1985. Mutagen-directed attenuation of Rift Valley fever virus as a method for vaccine development. J. Gen. Virol. 66, 2217–2271. Coetzer, J., Barnard, B., 1977. Hydrops amnii in sheep associated with hydranencephaly and arthrogryposis with wesselsbron disease and rift valley fever viruses as aetiological agents. Onderstepoort J. Vet. Res. 44, 119–126. Crawford, S., Labbé, M., Cohen, J., Burroughs, M., Zhou, Y., Estes, M., 1994. Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. Elliott, R., 1996. In: Frankel-Conrat, H., Wagner, R.R. (Eds.), “The Bunyaviridae.” The Viruses. Plenum Press, New York. Flick, R., Bouloy, M., 2005. Rift Valley fever virus. Curr. Mol. Med. 5, 827–834. Gerdes, G., 2004. Rift Valley fever. Rev. Sci. Tech. 23, 613–623. Gerrard, S., Bird, B., Albariño, C., Nichol, S., 2007. The NSm proteins of Rift Valley fever virus are dispensable for maturation, replication and infection. Virology 359, 459–465. Grgacic, E., Anderson, D., 2006. Virus-like particles: passport to immune recognition. Methods 40, 60–65. Habjan, M., Penski, N., Spiegel, M., Weber, F., 2008. T7 RNA polymerase-dependent and -independent systems for cDNA-based rescue of Rift Valley fever virus. J. Gen. Virol. 89, 2157–2166. Habjan, M., Penski, N., Wagner, V., Spiegel, M., Överby, A.K., Kochs, G., Huiskonen, J., Weber, F., 2009. Efficient production of Rift Valley fever virus-like particles: the antiviral protein MxA can inhibit primary transcription of Bunyaviruses. Virology 385, 400–408. Harper, D., Franco, E., Wheeler, C., Moscicki, A., Romanowski, B., Roteli-Martins, C., Jenkins, D., Schuind, A., Costa Clemens, S., Dubin, G., 2006. Sustained efficacy up to 4.5 years of a bivalent L1 virus-like particle vaccine against human papillomavirus types 16 and 18: follow-up from a randomised control trial. Lancet 367, 1247–1255. Hunter, P., Erasmus, B., Vorster, J., 2002. Teratogenicity of a mutagenised Rift Valley fever virus (MVP 12) in sheep. Onderstepoort J. Vet. Res. 69, 95–98. Ikegami, T., Won, S., Peters, C.J., Makino, S., 2006. Rescue of infectious rift valley fever virus entirely from cDNA, analysis of virus lacking the NSs gene, and expression of a foreign gene. J. Virol. 80, 2933–2940. Jansen van Vuren, P., Potgieter, A., Paweska, J., van Dijk, A., 2007. Preparation and evaluation of a recombinant Rift Valley fever virus N protein for the detection of IgG and IgM antibodies in humans and animals by indirect ELISA. J. Virol. Methods 140, 106–114. Kark, J., Aynor, Y., Peters, C.J., 1982. A Rift Valley fever vaccine trial. I. Side effects and serologic response over a six-month follow-up. Am. J. Epidemiol. 116, 808–820. Kirnbauer, R., Booy, F., Cheng, N., Lowy, D., Schiller, J., 1992. Papillomavirus L1 major capsid protein self-assembles into virus-like particles that are highly immunogenic. Proc. Natl. Acad. Sci. U.S.A. 89, 12180–12184. Latham, T., Galarza, J., 2001. Formation of wild-type and chimeric influenza virus-like particles following simultaneous expression of only four structural proteins. J. Virol. 75, 6154–6165. Le May, N., Dubaele, S., Proietti De Santis, L., Billecocq, A., Bouloy, M., Egly, J., 2004. TFIIH transcription factor, a target for the Rift Valley hemorrhagic fever virus. Cell 116, 541–550. Le May, N., Gauliard, N., Billecocq, A., Bouloy, M., 2005. The N terminus of Rift Valley fever virus nucleoprotein is essential for dimerization. J. Virol. 79, 11974–11980. Lindkvist, M., Lahti, K., Lilliehöök, B., Holmström, A., Ahlm, C., Bucht, G., 2007. Crossreactive immune responses in mice after genetic vaccination with cDNA encoding hantavirus nucleocapsid proteins. Vaccine 25, 1690–1699. Lubroth, J., Rweyemamu, M., Viljoen, G., Diallo, A., Dungu, B., Amanfu, W., 2007. Veterinary vaccines and their use in developing countries. Rev. Sci. Tech. 26, 179–201. Mao, C., Koutsky, L., Ault, K., Wheeler, C., Brown, D., Wiley, D., Alvarez, F., Bautista, O., Jansen, K., Barr, E., 2006. Efficacy of human papillomavirus-16 vaccine to prevent cervical intraepithelial neoplasia: a randomized controlled trial. Obstet. Gynecol. 107, 18–27. McAleer, W., Buynak, E., Maigetter, R., Wampler, D., Miller, W., Hilleman, M., 1984. Human hepatitis B vaccine from recombinant yeast. Nature 307, 178–180. Meegan, J., 1979. The Rift Valley fever epizootic in Egypt 1977–78. 1. Description of the epizootic and virological studies. Trans. R. Soc. Trop. Med. Hyg. 73, 618–623.
J. Näslund et al. / Virology 385 (2009) 409–415 Metwally, S., 2008. In: Brown, C., Torres, A. (Eds.), Foreign Animal Diseases, 7th ed. Boca Publications Group, Inc., Boca Raton, FL, pp. 369–376. Morrill, J.C., Mebus, C.A., Peters, C.J., 1997. Safety and efficacy of a mutagen-attenuated Rift Valley fever virus vaccine in cattle. Am J Vet Res. 58 (10), 1104–1109 PMID: 9328662. Muller, R., Saluzzo, J., Lopez, N., Dreier, T., Turell, M., Smith, J., Bouloy, M., 1995. Characterization of clone 13, a naturally attenuated avirulent isolate of Rift Valley fever virus, which is altered in the small segment. Am. J. Trop. Med. Hyg. 53, 405–411. Näslund, J., Lagerqvist, N., Lundkvist, Å., Evander, M., Ahlm, C., Bucht, G., 2008. Kinetics of Rift Valley Fever Virus in experimentally infected mice using quantitative realtime RT-PCR. J. Virol. Methods 151, 277–282. Pittman, P., Liu, C., Cannon, T., Makuch, R., Mangiafico, J., Gibbs, P., Peters, C.J., 1999. Immunogenicity of an inactivated Rift Valley fever vaccine in humans: a 12-year experience. Vaccine 18, 181–189. Pushko, P., Tumpey, T., Bu, F., Knell, J., Robinson, R., Smith, G., 2005. Influenza virus-like particles comprised of the HA, NA, and M1 proteins of H9N2 influenza virus induce protective immune responses in BALB/c mice. Vaccine 23, 5751–5759. Reading, S., Dimmock, N., 2007. Neutralization of animal virus infectivity by antibody. Arch. Virol. 152, 1047–1059. Sall, A., Zanotto, P., Sene, O., Zeller, H., Digoutte, J., Thiongane, Y., Bouloy, M., 1999. Genetic reassortment of Rift Valley fever virus in nature. J. Virol. 73, 8196–8200. Siam, A., Meegan, J., Gharbawi, K., 1980. Rift Valley fever ocular manifestations: observations during the 1977 epidemic in Egypt. Br. J. Ophthalmol. 64, 366–374. Smithburn, K.C., 1949. Rift Valley Fever: the neurotropic adaptation of the virus and the experimental use of this modified virus as a vaccine. Br. J. Exp. Pathol. 1, 1–16. Swenson, D., Warfield, K., Negley, D., Schmaljohn, A., Aman, M., Bavari, S., 2005. Viruslike particles exhibit potential as a pan-filovirus vaccine for both Ebola and Marburg viral infections. Vaccine 23, 3033–3042.
415
van Oirschot, J., 1999. Diva vaccines that reduce virus transmission. J. Biotechnol. 73, 195–205. Vialat, P., Muller, R., Vu, T., Prehaud, C., Bouloy, M., 1997. Mapping of the mutations present in the genome of the Rift Valley fever virus attenuated MP12 strain and their putative role in attenuation. Virus Res. 52, 43–50. Vieira, H., Estêvão, C., Roldão, A., Peixoto, C., Sousa, M., Cruz, P., Carrondo, M., Alves, P., 2005. Triple layered rotavirus VLP production: kinetics of vector replication, mRNA stability and recombinant protein production. J. Biotechnol. 120, 72–82. Villa, L., Costa, R., Petta, C., Andrade, R., Ault, K., Giuliano, A., Wheeler, C., Koutsky, L., Malm, C., Lehtinen, M., Skjeldestad, F., Olsson, S., Steinwall, M., Brown, D., Kurman, R., Ronnett, B., Stoler, M., Ferenczy, A., Harper, D., Tamms, G., Yu, J., Lupinacci, L., Railkar, R., Taddeo, F., Jansen, K., Esser, M., Sings, H., Saah, A., Barr, E., 2005. Prophylactic quadrivalent human papillomavirus (types 6, 11, 16, and 18) L1 viruslike particle vaccine in young women: a randomised double-blind placebocontrolled multicentre phase II efficacy trial. Lancet Oncol. 6, 271–278. Warfield, K., Swenson, D., Negley, D., Schmaljohn, A., Aman, M., Bavari, S., 2004. Marburg virus-like particles protect guinea pigs from lethal Marburg virus infection. Vaccine 22, 3495–3502. Won, S., Ikegami, T., Peters, C.J., Makino, S., 2007. NSm protein of Rift Valley fever virus suppresses virus-induced apoptosis. J. Virol. 81, 13335–13345. Zaki, A., Coudrier, D., Yousef, A., Fakeeh, M., Bouloy, M., Billecocq, A., 2006. Production of monoclonal antibodies against Rift Valley fever virus Application for rapid diagnosis tests (virus detection and ELISA) in human sera. J. Virol. Methods 131, 34–40. Zhou, J., Sun, X., Stenzel, D., Frazer, I., 1991. Expression of vaccinia recombinant HPV 16 L1 and L2 ORF proteins in epithelial cells is sufficient for assembly of HPV virion-like particles. Virology 185, 251–257.
Contents lists available at ScienceDirect
Virology j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / y v i r o
Vaccination with virus-like particles protects mice from lethal infection of Rift Valley Fever Virus Jonas Näslund a,b,c,1, Nina Lagerqvist a,b,d,1, Matthias Habjan e, Åke Lundkvist d, Magnus Evander c, Clas Ahlm b, Friedemann Weber e,⁎, Göran Bucht a,f,⁎ a
Swedish Defence Research Agency, Department of CBRN Defence and Security, SE-901 82 Umeå, Sweden Division of Infectious Diseases, Department of Clinical Microbiology Umeå University, SE-901 85 Umeå, Sweden Division of Virology, Department of Clinical Microbiology Umeå University, SE-901 85 Umeå, Sweden d Swedish Institute for Infectious Disease Control, SE-171 82 Solna, Sweden e Department of Virology, University of Freiburg, D-79008 Freiburg, Germany f National Environment Agency, Environmental Health Institute, 11 Biopolis Way, 06-05/08, Helios Block, Singapore 138667, Singapore b c
a r t i c l e
i n f o
Article history: Received 24 September 2008 Returned to author for revision 30 October 2008 Accepted 8 December 2008 Available online 20 January 2009 Keywords: Rift Valley Fever Mouse model Virus-like particle Vaccine Challenge
a b s t r a c t Rift Valley Fever virus (RVFV) regularly accounts for severe and often lethal outbreaks among livestock and humans in Africa. Safe and effective veterinarian and human vaccines are highly needed. We present evidence that administration of RVF virus-like particles (VLPs) induces protective immunity in mice. In an accompanying paper, (Habjan, M., Penski, N., Wagner, V., Spiegel, M., Överby, A.K., Kochs, G., Huiskonen, J., Weber, F., 2009. Efficient production of Rift Valley fever virus-like particles: the antiviral protein MxA can inhibit primary transcription of Bunyaviruses. Virology 385, 400–408) we report the production of these VLPs in mammalian cells. After three subsequent immunizations with 1 × 106 VLPs/dose, high titers of virusneutralizing antibodies were detected; 11 out of 12 mice were protected from challenge and only 1 out of 12 mice survived infection in the control groups. VLP vaccination efficiently suppressed replication of the challenge virus, whereas in the control animals high RNA levels and increasing antibody titers against the nucleocapsid protein indicated extensive viral replication. Our study demonstrates that the RVF VLPs are highly immunogenic and confer protection against RVFV infection in mice. In the test groups, the vaccinated mice did not exhibit any side effects, and the lack of anti-nucleocapsid protein antibodies serologically distinguished vaccinated animals from experimentally infected animals. © 2008 Elsevier Inc. All rights reserved.
Introduction Rift Valley Fever (RVF) is an emerging infectious disease and considered as one of the most important viral zoonoses in Africa. The disease was restricted to the sub-Saharan region of Africa until the middle of the 1970s. Since then, RVF has become endemic throughout Africa and more recently RVF has been found also on Arabian Peninsula (Balkhy and Memish, 2003; Gerdes, 2004). Epizootics of RVF are characterized by increasing numbers of abortions and high neonatal mortality. Among adult animals, the clinical manifestations range from asymptomatic to more severe conditions like hepatitis and death, although the progression and ⁎ Corresponding authors. F. Weber is to be contacted at Fax: +49 761 203 6634. G. Bucht, Fax: +65 6777 8029. E-mail addresses: [email protected] (J. Näslund), [email protected] (N. Lagerqvist), [email protected] (M. Habjan), [email protected] (Å. Lundkvist), [email protected] (M. Evander), [email protected] (C. Ahlm), [email protected] (F. Weber), [email protected] (G. Bucht). 1 These authors contributed equally to this work. 0042-6822/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.virol.2008.12.012
severity of the disease varies considerably between different animal species (Elliott, 1996). In humans, RVF is considered as a temporarily incapacitating illness with fever, myalgia, and headache. However, more severe clinical symptoms—such as encephalitis, ocular disease, hepatitis, and hemorrhagic fever—have been observed (Al-Hazmi et al., 2003; Siam et al., 1980). Presently, treatments are symptomatic combined with an occasional need of intensive care. The RVF virus (RVFV), a member of the Phlebovirus genera in the Bunyaviridae family, is transmitted by several different mosquito species and has an unusual broad range of susceptible animal hosts (Flick and Bouloy, 2005). As for other members of this family of viruses, the genetic information is divided into three negative stranded RNA segments: (S)mall, (M)edium and (L)arge. The L-segment encodes for the RNA polymerase, and the M- and S-segments encode for two glycoproteins (GN/GC), the 78 kDa protein and the nucleocapsid (N) protein, respectively. Furthermore, the RVFV genome encodes for two non-structural proteins: NSs and NSm of the S and M segment, correspondingly (Elliott, 1996). The NSs protein is involved in disruption of the host antiviral response (Billecocq et al., 2004; Le May et al., 2004) and the NSm protein is reported to suppress apoptotic pathways in host cells after infection (Won et al., 2007).
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The recent geographical expansion of RVF, combined with the ability of the virus to maintain in newly conquered areas due to the wide range of mosquito vectors and susceptible hosts, imply that novel therapeutics and vaccines are necessary. Currently, there are no registered vaccines for human use. However, a formalin inactivated virus vaccine for humans has been developed (Kark et al., 1982; Pittman et al., 1999), although the distribution has been limited to workers at high-risk occupations. The live attenuated Smithburn neurotropic strain (Smithburn, 1949) and formalin inactivated virus vaccines produced in South Africa and Egypt (Metwally, 2008) are approved for veterinarian use. The Smithburn vaccine is highly immunogenic, but is teratogenic in pregnant cattle and sheep (Botros et al., 2006; Coetzer and Barnard, 1977). The formalin inactivated vaccines are safe, but less immunogenic (Lubroth et al., 2007). Other vaccine candidates under evaluation are Clone 13 and MP12. Both have shown promising results in animal trials. Clone 13 is a natural attenuated isolate from a benign human case in the Central African Republic (Muller et al., 1995), and MP12 is a chemically attenuated virus derived from the Egyptian wild-type isolate ZH548 (Caplen et al., 1985; Vialat et al., 1997). The immunogenicity and pathogenicity of these candidates have been evaluated in various animal species (Muller et al., 1995; Morrill et al., 1997). The most extensively tested, MP12, induced foetal malformations during the first trimester (Hunter et al., 2002). More recently, reverse genetic systems have been established for RVFV that allow genetic engineering of the virus genomes and production of recombinant virus particles (Billecocq et al., 2008; Gerrard et al., 2007; Habjan et al., 2008; Ikegami et al., 2006). Such techniques have been used to produce attenuated RVFV particles which have displayed promising results as a vaccine in a rodent model (Bird et al., 2008). However, inherent problems of live vaccines are the expensive production due to high biosafety precautions and the risk of side effects such as teratogenicity and infections in immuno-suppressed individuals. Moreover, replicating RVFV vaccines could theoretically be spread by mosquitoes. Another approach for vaccine development is to use virus-like particles (VLPs) that are formed when the structural (envelope and/or nucleocapsid) proteins self-assemble to replication-deficient particles. In comparison to component vaccines based on recombinant or subunit proteins, VLPs are in general more immunogenic and more natural in presenting conformational epitopes, and hence, more likely to induce neutralizing antibodies (Grgacic and Anderson, 2006). In an accompanying paper, we describe the production of RVF VLPs in mammalian cells by expressing the viral structural genes along with a Renilla luciferase reporter minigenome (Habjan et al., 2009). These VLPs have the size, morphology, and protein composition resembling authentic RVFV particles. They contain nucleocapsids consisting of N and the reporter RNA minigenome and display the envelope (GN and GC) proteins at their surface. Moreover, the VLPs can enter target cells and carry out transcription of the reporter gene, but they cannot replicate or produce progeny. They can be neutralized by RVFVspecific antisera. Here, we report the use of these VLPs as a vaccine against RVF. The safety and immunogenicity were evaluated in mice and the results indicate that immunization with VLPs confers protection against challenge with lethal doses of RVFV. These VLPs offer a promising alternative to attenuated virus mutants as an efficient and safe vaccine against RVF. Results The RVF VLPs are immunogenic and induce virus-neutralizing antibodies Thirty mice were immunized three times with two weeks intermission with either 1 × 105 VLPs/dose, 1 × 106 VLPs/dose, sterile PBS or supernatant of cells transfected with all VLP constructs except the glycoprotein plasmid (Habjan et al., 2009). The two latter groups
constituted the negative controls. No clinical manifestations derived from the vaccine were observed in any of these mice. After three immunizations, the sera were analyzed for the presence of anti-RVFV specific antibodies. Immunofluorescense analysis (IFA) performed on virus-infected mammalian cells showed that all VLP-vaccinated mice sero-converted and antibodies were detectable at serum dilutions of up to 1:2700 (Fig. 1A). To analyze the antibody response in more detail, IFA was performed on mammalian cells transfected with either cDNA encoding the N protein or the GN/GC proteins (data not shown). VLP-vaccination induced antibodies towards the GN/GC protein among all mice, with titers ranging from 300 up to 900. On the other hand, no detectable N protein specific antibodies (total Ig) were observed by IFA after three subsequent VLP-vaccinations (data not shown). The virus neutralizing antibody titers were determined by a plaque reduction neutralization test (PRNT) (Fig. 1B). In the group of mice administered with 1 × 105 VLPs/dose, three animals (3/6) showed low titers, equal to or below 50, the remaining three animals were found to have neutralizing titers of 250. When the high-dose group administered with 1 × 106 VLPs/dose were analyzed, high titers of neutralizing antibodies were found; in this dose group, the titer values ranged from 250 to 1250 (Fig. 1B). However, one animal of this group did not respond to the vaccination, and the neutralizing titer was below 50. No sero-conversion was detected among the control mice. No apparent differences in the immune responses, viral replication, clinical signs, or survival proportions were observed between mice from the two control groups having received either sterile PBS or a cell
Fig. 1. Sero-conversion in mice after VLP vaccination determined by immunofluorescence assay and plaque reduction neutralization tests. Mice sera were collected after three subsequent VLP immunizations and analyzed for panel A antibody reactivity towards RVFV-infected mammalian cells and panel B titer of virus neutralizing antibodies. Two dose groups are represented: mice vaccinated with 1 × 105 or 1 × 106 VLPs/dose. No sero-conversion was observed among the control mice and are therefore excluded from these graphs. The antibody titer was determined as the reciprocal of the highest positive serum dilution of (panel A) three-fold and (panel B) five-fold serially diluted sera, respectively.
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supernatant (see Materials and methods). Therefore, these two groups were consolidated and represent the control. Vaccination with RVF VLPs protects mice in challenge The mice were challenged two weeks after the third and final vaccination with a lethal dose (2.4 × 104 PFU) of RVFV. During the entire challenge experiment, the mice were monitored twice a day for signs of infection. The survival proportions among each vaccination group are shown in Fig. 2. Three out of six mice survived challenge in the low-dose group, immunized with 1 × 105 VLPs/dose. Mild clinical signs were observed among two of the three surviving animals around day 7 p.i., followed by complete recovery one to two days thereafter. Eleven out of twelve mice vaccinated with 1 × 106 VLPs/dose survived and no clinical manifestations were observed among the surviving animals during the experimental period. Interestingly, three mice vaccinated with the lower dose of VLP and one mouse of the high dose group showing the lowest titers of neutralizing antibodies, as shown in Fig. 1B, had to be sacrificed at days 9 and 10 due to severe clinical signs. One animal of twelve in the control group survived challenge without displaying any clinical manifestations. The remaining mice in the control group displayed severe clinical signs, in some cases as early as 4 to 5 days p.i., and two control animals developed clinical signs as late as 12 days p.i. Survival proportion was significantly higher in mice vaccinated with 1 × 106 VLPs/dose than in the control group (Chisquare test, p = 0.00004). However, the survival proportion was not significant for the mice in the low-dose group (Fisher exact test, p = 0.0833). VLP vaccination suppresses sero-conversion against the N protein after RVFV challenge Since vaccination with the RVF VLPs used in these experiments did not induce antibodies directed towards the N protein (pre-challenge sera obtained day 0 p.i., Fig. 3B, C), any detection of anti-N specific antibodies after challenge is an indication of viral replication in the animal hosts. When analyzing sera from the control animals after challenge (Fig. 3A), anti-N antibodies were clearly observed by ELISA from days 4 and 5 p.i., followed by increasing and highly individual titers. However, the VLP-vaccinated mice displayed no or a moderate rise in the antibody levels (Figs. 3B, C), and the curves are clearly separated from those of the control animals. The animals in the lowdose group with slightly elevated titer levels around day 5 p.i. and the high-dose animals with constantly low antibody titers indicate Fig. 3. Antibody response towards the N protein in vaccinated and control animals after challenge with RVFV. Serum samples were collected at different days after challenge and analysed by ELISA for the presence of specific antibodies directed against the N protein. The graphs represent antibody titers in serum samples of panel A control animals, panel B mice vaccinated with 1 × 105 VLPs/dose and panel C mice vaccinated with 1 × 106 VLPs/dose. The curves show the mean value from two separate measurements and the error bars represent the standard deviation. Incomplete curves correspond to samples taken from mice that were euthanized early due to a moribund condition.
negligible virus replication in the vaccinated animals after challenge. Only small variations in antibody levels were detected after day 12 p.i. (data not shown). VLP immunization inhibits RVFV replication in mice
Fig. 2. Percent survival of VLP immunized mice after RVFV challenge. The mice were vaccinated with either 1 × 105 or 1 × 106 VLPs/dose then challenged with 2.4 × 104 PFU of RVFV strain ZH548. Mice immunized with a cell suspension or PBS constitute the control group (Ctrl).
To further verify if vaccination with VLPs suppresses replication of the wild-type RVFV after challenge, viral RNA copy numbers in blood were quantified using a SYBR-green based qRT-PCR assay (Fig. 4). When analyzing serum samples of the control and the low-dose mice (Figs. 4A, B), a great divergence in the detected levels of RNA was observed between individual mice. A large proportion of the control
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Fig. 4. Quantification of RVFV RNA in blood samples of VLP-vaccinated and control animals after challenge. Blood samples were collected before infection (day 0) and on days three, five, seven, ten, and twelve after infection and analyzed for the presence of viral RNA by qRT-PCR. The graphs represent panel A control animals, panel B mice vaccinated with 1 × 105 VLPs/dose, and panel C mice vaccinated with 1 × 106 VLPs/dose. The curves illustrate RNA copies/ml blood and represent the mean value of two replicates. The error bars show the standard deviation between measurements.
mice showed high levels of viral RNA 3 days p.i (Fig. 4A), while animals of the low-dose group reached their maximum viral load 2 days later (Fig. 4B). Animals of the high-dose group also displayed a delayed onset of viral replication (Fig. 4C) and only low viral RNA levels, in comparison to the other groups, were found (Fig. 4). In conclusion, the RVF VLPs were found to be safe and highly immunogenic in vivo and protected mice from lethal challenge by RVFV. Discussion RVF is an emerging infectious disease characterized by high mortality rates among foetuses, neonates, and juvenile livestock and causes a severe febrile illness also in man. To prevent RVF epidemics and the concomitant serious socio-economic consequences, new and improved vaccine strategies are of utmost importance.
Recently, VLPs based on the RVFV envelope proteins have been developed (Habjan et al., 2009), and herein we present the use of these RVF VLPs as a vaccine in a mouse model. These VLPs, when analyzed by electron microscopy, have a similar morphology as the authentic RVFV particles. Moreover, these VLPs display correctly processed viral glycoproteins in their envelope, proteins that are recognized by neutralizing antisera and contain transcriptionally active minigenome-nucleocapsids, but are unable to replicate and produce progeny (Habjan et al., 2009). In general, VLPs are reported to have similar properties as the virulent strains of the corresponding agents since the predominant structural antigens are displayed to the immune system in a native conformation (Grgacic and Anderson, 2006). The capability of VLPs to confer protective immunity has previously been demonstrated for a number of viruses. VLPs assembled from recombinant human papilloma virus capsid proteins are used as a vaccine against cervical cancer (Harper et al., 2006; Kirnbauer et al., 1992; Mao et al., 2006; Villa et al., 2005; Zhou et al., 1991) and VLPs of the small surface antigen of hepatitis B virus (André and Safary, 1987; McAleer et al., 1984) are also successfully used. In addition, several preclinical studies have been performed targeting, e.g., influenza (Latham and Galarza, 2001; Pushko et al., 2005), Rotavirus (Bertolotti-Ciarlet et al., 2003; Crawford et al., 1994; Vieira et al., 2005), and filovirus infections (Swenson et al., 2005; Warfield et al., 2004). Antibody-mediated virus neutralization constitutes an essential part in the control of infectious agents (Reading and Dimmock, 2007). In the present study, virus neutralization titers of 250 or more were found to be protective and associated with survival after challenge. All mice but one obtained serum neutralizing titers of 250, or more, after vaccination with the high VLP dose. Similar titers were only found for half of the animals vaccinated with the lower dose of VLPs. An unexpected low neutralization titer was found in one animal of the high-dose group and was probably the cause for the serious clinical signs that later lead to euthanization of this animal after challenge. Surprisingly, two of the surviving animals from the low-dose group displayed mild clinical signs despite the fact that they acquired neutralizing titers of 250. Three mice of the low-dose group and one animal of the high dose-group with the lowest neutralizing titers (equal to or below 50) had to be sacrificed at days 9 and 10 due to severe clinical manifestations. In the control group, the majority of the animals developed severe clinical signs and high viremia early after challenge and a large proportion (92%) of the animals succumbed to infection. One control animal did not display any clinical signs of infection, although high viral load and anti-N antibodies indicated extensive RVFV replication. There was a strong association between the administered VLP dose, neutralizing antibody titers and the survival rate. Importantly, a significant protection (92%) was observed among the animals administered the high VLP dose in contrast to the animals in the low-dose group where only 50% survived infection. The surviving high-dose animals were also clearly distinguished from both the control and the low-dose animals in the context of viral RNA levels and anti-N antibody titers detected after infection. Interestingly, no mouse displayed detectable levels of anti-N specific antibodies after VLP vaccination. RVF VLPs do not replicate and the N proteins present in the interior of the VLP were apparently not enough to elicit an immune response against the N protein. However, all mice in this study sero-converted towards the glycoproteins after vaccination with the RVF VLPs. The N protein of RVFV is a major antigen in the early serological response to infection and is particularly useful during clinical diagnostics and surveillance. Due to the zoonotic nature of RVF, surveillance is of major importance in the international trade of animals and animal-related products. Marker vaccines make it possible to differentiate infected from vaccinated individuals with specific diagnostic tests, valuable tools in the framework of control programs (van Oirschot, 1999). To use the benefits of marker vaccines fully, it is essential to use easy and
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assessable diagnostic tests. Immuno-fluorescence performed on infected or transfected cells, recombinant N protein based ELISA and neutralization tests are techniques widely used in laboratories that specialize in RVFV (Bird et al., 2008; Jansen van Vuren et al., 2007; Näslund et al., 2008; Zaki et al., 2006). Since VLP vaccination did not induce an anti-N specific antibody response, these already established techniques make it easy to monitor and evaluate future VLP vaccination programs. Currently, available RVF vaccines are based on attenuated or inactivated virus preparations. Because several disadvantages have been reported when using these vaccines (Lubroth et al., 2007), attempts to develop new vaccines is a priority. At the moment, the most advanced and promising live RVFV vaccine candidate is the recently produced rZH501-ΔNSs:GFP-ΔNSm, which lacks two RVFV virulence factors, NSs and NSm (Bird et al., 2008). Because there is a complete deletion of these genes involved in virulence, it is highly unlikely that the virus can revert to full virulence. However, other inherent drawbacks of live attenuated vaccines are -the need for an elevated biosafety level facility for production and the risk of using attenuated viruses in immunocompromised individuals - still remain. When vaccines based on the VLP strategy are used, the safety concerns and the logistical disadvantages are circumvented. Because available RVFV vaccines are associated with deleterious effects or low immunogenicity, there is an imperative need for the development of alternative RVF vaccine strategies. We demonstrate that RVF VLPs are safe and confer protective immunity in mice with respect to a lethal dose of RVFV. Furthermore, vaccinated individuals could readily be distinguished from infected with N-specific diagnostic tests. These results clearly show the potential of using VLPs as a future vaccine candidate against RVF. Materials and methods Cells and virus strain The RVFV strain ZH548 was originally isolated from an uncomplicated human case that occurred during the Egyptian outbreak in 1977 (Meegan, 1979; Sall et al., 1999). Low passage ZH548 stocks were propagated and titrated in confluent monolayers of Vero E6 cells (ATCC® Number CRL-1586) before used for challenge. Vero E6 cells were cultivated in Dulbecco's modified Eagle's medium (DMEM) (GIBCO, Invitrogen, Carlsbad, CA) supplemented with 10% FCS, 10 mM HEPES, and 1 mM sodium pyruvate and antibiotics (100 U penicillin/ml, 100 μg/ml streptomycin). For immunological methods, BHK-21 cells (ATCC® Number, CCL-10) were maintained in Glasgow MEM (GIBCO, Invitrogen) and supplemented with 5% FCS, 1.3 g/l Tryptose (Difco™, Becton, Dickinson and Company, Sparks, MD), 10 mM HEPES, and 1 mM sodium pyruvate and antibiotics (100 U penicillin/ml, 100 μg/ml streptomycin). Human 293T (ATCC® Number CRL-11268) cells were cultivated in DMEM and supplemented with 10% FCS, 0.4% sodium hydrogen carbonate, 1 mM sodium pyruvate, 0.53 mg/ml glutamine, and antibiotics (100 U penicillin/ml, 100 μg/ml streptomycin). All cells were cultivated at 5% CO2 at 37 °C. VLP production and purification Subconfluent monolayers of 293T cells in 90 mm dishes were transfected with 3 μg of plasmids pI.18_RVFV_N, pI.18_RVFV_L, pI.18_RVFV_M, and pHH21_RVFV_vMRen (Habjan et al., 2009) using Nanofectin (PAA Laboratories, Linz, Austria) transfection reagent. Two days post transfection, supernatants were collected and clarified from cell debris by centrifugation (6000 ×g, 10 min at 4 °C). The particles were then concentrated by ultracentrifugation at 24,000 rpm at 4 °C for 2 h in a Beckman SW41Ti rotor. The pellet
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was dried for 5 min before being re-suspended in phosphatebuffered saline (PBS). To define the titer, VLPs expressing GFP were generated and concentrated in parallel to the Ren-VLPs which were used for the vaccinations. BSR-T7/5 cells expressing RVFV L and N (Habjan et al., 2009) were incubated for 1 h with ten-fold serial dilutions of supernatants containing GFP-VLPs. Twenty-four hours later the number of GFP-positive (i.e. VLP-infected) cells was determined by fluorescence microscopy and the titer of the GFPVLP preparation was calculated accordingly. The titer of the RenVLPs was estimated to be in the same range. Western blot analysis confirmed that the amounts of GFP-VLPs and Ren-VLPs were comparable. VLP immunization This study used 33 female C57BL/6 mice (Scanbur BK) 6– 7 weeks old. The mice were kept in cages under BSL-2 conditions during the whole vaccination routine with free access to food and water. Two VLP doses were administered, 1 × 105 VLPs/dose (n = 6) and 1 × 106 VLPs/dose (n = 12). To serve as negative controls, 12 mice were inoculated with either sterile PBS (n = 5) or with a cell supernatant (n = 7), similar to what was used in the production of the RVF VLPs. Two independent vaccinations were performed with comparable immunization schedules and different batches of VLPs, although the second round did not include the lower dose group. All animals were vaccinated intraperitoneally (i.p.) three times (two weeks intermission) with the VLPs diluted in sterile PBS to a final volume of 200 μl. RVFV challenge Two weeks after the third vaccination, a challenge dose containing 2.4 × 104 PFU of RVFV strain ZH548 were administered i. p. During these procedures, the mice were kept individually in micro-isolator pans within an animal isolator (Bell Isolation Systems Ltd, Scotland, UK). All manipulations with infected mice and/or samples involving RVFV were performed strictly under BSL-3 conditions. The animals were examined twice daily for visual signs of illness until the experiments were ended, 12–14 days post infection (p.i.). Clinical signs observed among mice after infection ranged from distress, ruffled fur, and weight loss to fatigue, neurological manifestations, and a “hunchback-like” posture. Mice found in a moribund condition were instantly euthanized. Blood samples were collected before each immunization and every second or third day p.i. To ensure a sterile milieu, non-infected mice (n = 3) were kept under the same conditions during the entire experiment. This project was approved by The Animal Research Ethics Committee of Umeå University, Sweden. Plaque reduction neutralization test (PRNT) Heat inactivated sera, collected two weeks after the third immunization, were five-fold serially diluted in sterile PBS and incubated with a virus suspension containing 30 PFU of ZH548 before adsorption to a confluent monolayer of BHK-21 cells. After a 90 min incubation period and subsequent washing, the cells were covered with an overlay containing 1% CMC (Carboxy-Methyl Cellulose, Aquacide II, Calbiochem®, California, USA) prepared in maintenance DMEM and incubated for six days at 37 °C/5% CO2. After fixation with 10% formaldehyde, the plaques were visualized by counterstaining with 1% crystal violet in water containing 20% ethanol and 0.7% NaCl and counted after destaining with tap water. Sera of non-infected or infected mice constituted the negative and positive controls, respectively. The neutralizing antibody titers were determined as the reciprocal of the highest serum dilution that reduced the number of plaques by 80%.
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Immuno-fluorescence assay (IFA) and enzyme-linked immuno-sorbent assay (ELISA) For IFA, BHK-21 cells were infected with RVFV (strain ZH548) at a multiplicity of infection (MOI) of 1 or transfected and grown on cover slips. Transfection was performed using the eukaryotic expression vector (pcDNA3.1, Invitrogen) containing cDNA encoding the N protein or the open reading frame of the GN/GC proteins and FuGene 6 transfection reagent (Roche Inc.) according to the manufacturer's instructions. The cells were fixed in 4% paraformaldehyde and permeabilized with 0.2% Saponin in PBS 36 h p.i. or 48 h post transfection. After incubation with three-fold serial dilutions of the serum samples, collected two weeks after the third vaccination, infected or transfected cells were labelled using a goat anti-mouse Alexa Fluor™ 488 conjugate total Ig or a FITC conjugated rabbit anti-mouse total Ig, respectively, according to the instructions of the manufacturers (Molecular Probes, Invitrogen). ELISA was performed as described earlier (Näslund et al., 2008) using recombinant N-protein (GenBank accession no. AF134534) (Sall et al., 1999) of RVFV and constructed and purified as previously described (Le May et al., 2005; Lindkvist et al., 2007). The sera used were collected at the day of infection (day 0) then three, five, seven, ten and twelve days p.i. The serum samples were prepared as four-fold serial dilutions, and the antibody titers were calculated as the inverse dilution at the intersection of the straight line and the x-axis. Serum samples from pre-bleedings and non-infected mice were present on each plate to determine unspecific reactivity. Quantitative real time reverse transcriptase PCR (qRT-PCR) The viral load was measured in blood, collected at the day of infection (day 0) and day three, five, seven, ten, and twelve p.i, using a qRT-PCR as previously described (Näslund et al., 2008). Briefly, RNA was extracted and purified using Trizol LS (Invitrogen) and RNeasy Mini Kit (Qiagen Inc.) in accordance to the manufacturer's instructions. cDNA were synthesized using SuperScript™ III RT according to manufacturer's instructions (Invitrogen). qRT-PCR was performed in 24 μl reactions using SYBR-green (Biotools, B&M Labs, Madrid, Spain) with primers targeting the S-segment of RVFV (available from the authors upon request). Statistical methods Survival proportions were compared using Chi-square test and Fischer exact test for values under five (Epi Info™, Version 3.5). Quantitative variables were based on measurements of a minimum of two independent experiments containing duplicate samples. Variables are expressed as means, and the error bars correspond to the standard deviation between the replicates. Acknowledgments Ms. Karin Wallgren is greatly acknowledged for her excellent technical support during the immunizations and bleedings of the mice. This study was supported by the Swedish Defence Agency, the Medical Faculty of Umeå University, grants from the County Council of Västerbotten, and by the grant We 2616/5-1 from the Deutsche Forschungsgemeinschaft. This project was also partially supported by grants from the Swedish Research Council (project 12177) and the European Community (contract no. QLK2-CT-2002-01358). References Al-Hazmi, M., Ayoola, E.A., Abdurahman, M., Banzal, S., Ashraf, J., El-Bushra, A., Hazmi, A., Abdullah, M., Abbo, H., Elamin, A., Al-Sammani, el.-T., Gadour, M., Menon, C., Hamza, M., Rahim, I., Hafez, M., Jambavalikar, M., Arishi, H., Aqeel, A., 2003. Epidemic Rift Valley fever in Saudi Arabia: a clinical study of severe illness in humans. Clin. Infect. Dis. 36, 245–252.
André, F., Safary, A., 1987. Summary of clinical findings on Engerix-B, a genetically engineered yeast derived hepatitis B vaccine. Postgrad. Med. J. 63 (Suppl 2), 169–177. Balkhy, H., Memish, Z., 2003. Rift Valley fever: an uninvited zoonosis in the Arabian peninsula. Int. J. Antimicrob. Agents 21, 153–157. Bertolotti-Ciarlet, A., Ciarlet, M., Crawford, S., Conner, M., Estes, M., 2003. Immunogenicity and protective efficacy of rotavirus 2/6-virus-like particles produced by a dual baculovirus expression vector and administered intramuscularly, intranasally, or orally to mice. Vaccine 21, 3885–3900. Billecocq, A., Spiegel, M., Vialat, P., Kohl, A., Weber, F., Bouloy, M., Haller, O., 2004. NSs protein of Rift Valley fever virus blocks interferon production by inhibiting host gene transcription. J. Virol. 78, 9798–9806. Billecocq, A., Gauliard, N., Le May, N., Elliott, R., Flick, R., Bouloy, M., 2008. RNA polymerase I-mediated expression of viral RNA for the rescue of infectious virulent and avirulent Rift Valley fever viruses. Virology 378, 377–384. Bird, B., Albariño, C., Hartman, A., Erickson, B., Ksiazek, T., Nichol, S., 2008. Rift valley fever virus lacking the NSs and NSm genes is highly attenuated, confers protective immunity from virulent virus challenge, and allows for differential identification of infected and vaccinated animals. J. Virol. 82, 2681–2691. Botros, B., Omar, A., Elian, K., Mohamed, G., Soliman, A., Salib, A., Salman, D., Saad, M., Earhart, K., 2006. Adverse response of non-indigenous cattle of European breeds to live attenuated Smithburn Rift Valley fever vaccine. J. Med. Virol. 78, 787–791. Caplen, H., Peters, C.J., Bishop, D., 1985. Mutagen-directed attenuation of Rift Valley fever virus as a method for vaccine development. J. Gen. Virol. 66, 2217–2271. Coetzer, J., Barnard, B., 1977. Hydrops amnii in sheep associated with hydranencephaly and arthrogryposis with wesselsbron disease and rift valley fever viruses as aetiological agents. Onderstepoort J. Vet. Res. 44, 119–126. Crawford, S., Labbé, M., Cohen, J., Burroughs, M., Zhou, Y., Estes, M., 1994. Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. Elliott, R., 1996. In: Frankel-Conrat, H., Wagner, R.R. (Eds.), “The Bunyaviridae.” The Viruses. Plenum Press, New York. Flick, R., Bouloy, M., 2005. Rift Valley fever virus. Curr. Mol. Med. 5, 827–834. Gerdes, G., 2004. Rift Valley fever. Rev. Sci. Tech. 23, 613–623. Gerrard, S., Bird, B., Albariño, C., Nichol, S., 2007. The NSm proteins of Rift Valley fever virus are dispensable for maturation, replication and infection. Virology 359, 459–465. Grgacic, E., Anderson, D., 2006. Virus-like particles: passport to immune recognition. Methods 40, 60–65. Habjan, M., Penski, N., Spiegel, M., Weber, F., 2008. T7 RNA polymerase-dependent and -independent systems for cDNA-based rescue of Rift Valley fever virus. J. Gen. Virol. 89, 2157–2166. Habjan, M., Penski, N., Wagner, V., Spiegel, M., Överby, A.K., Kochs, G., Huiskonen, J., Weber, F., 2009. Efficient production of Rift Valley fever virus-like particles: the antiviral protein MxA can inhibit primary transcription of Bunyaviruses. Virology 385, 400–408. Harper, D., Franco, E., Wheeler, C., Moscicki, A., Romanowski, B., Roteli-Martins, C., Jenkins, D., Schuind, A., Costa Clemens, S., Dubin, G., 2006. Sustained efficacy up to 4.5 years of a bivalent L1 virus-like particle vaccine against human papillomavirus types 16 and 18: follow-up from a randomised control trial. Lancet 367, 1247–1255. Hunter, P., Erasmus, B., Vorster, J., 2002. Teratogenicity of a mutagenised Rift Valley fever virus (MVP 12) in sheep. Onderstepoort J. Vet. Res. 69, 95–98. Ikegami, T., Won, S., Peters, C.J., Makino, S., 2006. Rescue of infectious rift valley fever virus entirely from cDNA, analysis of virus lacking the NSs gene, and expression of a foreign gene. J. Virol. 80, 2933–2940. Jansen van Vuren, P., Potgieter, A., Paweska, J., van Dijk, A., 2007. Preparation and evaluation of a recombinant Rift Valley fever virus N protein for the detection of IgG and IgM antibodies in humans and animals by indirect ELISA. J. Virol. Methods 140, 106–114. Kark, J., Aynor, Y., Peters, C.J., 1982. A Rift Valley fever vaccine trial. I. Side effects and serologic response over a six-month follow-up. Am. J. Epidemiol. 116, 808–820. Kirnbauer, R., Booy, F., Cheng, N., Lowy, D., Schiller, J., 1992. Papillomavirus L1 major capsid protein self-assembles into virus-like particles that are highly immunogenic. Proc. Natl. Acad. Sci. U.S.A. 89, 12180–12184. Latham, T., Galarza, J., 2001. Formation of wild-type and chimeric influenza virus-like particles following simultaneous expression of only four structural proteins. J. Virol. 75, 6154–6165. Le May, N., Dubaele, S., Proietti De Santis, L., Billecocq, A., Bouloy, M., Egly, J., 2004. TFIIH transcription factor, a target for the Rift Valley hemorrhagic fever virus. Cell 116, 541–550. Le May, N., Gauliard, N., Billecocq, A., Bouloy, M., 2005. The N terminus of Rift Valley fever virus nucleoprotein is essential for dimerization. J. Virol. 79, 11974–11980. Lindkvist, M., Lahti, K., Lilliehöök, B., Holmström, A., Ahlm, C., Bucht, G., 2007. Crossreactive immune responses in mice after genetic vaccination with cDNA encoding hantavirus nucleocapsid proteins. Vaccine 25, 1690–1699. Lubroth, J., Rweyemamu, M., Viljoen, G., Diallo, A., Dungu, B., Amanfu, W., 2007. Veterinary vaccines and their use in developing countries. Rev. Sci. Tech. 26, 179–201. Mao, C., Koutsky, L., Ault, K., Wheeler, C., Brown, D., Wiley, D., Alvarez, F., Bautista, O., Jansen, K., Barr, E., 2006. Efficacy of human papillomavirus-16 vaccine to prevent cervical intraepithelial neoplasia: a randomized controlled trial. Obstet. Gynecol. 107, 18–27. McAleer, W., Buynak, E., Maigetter, R., Wampler, D., Miller, W., Hilleman, M., 1984. Human hepatitis B vaccine from recombinant yeast. Nature 307, 178–180. Meegan, J., 1979. The Rift Valley fever epizootic in Egypt 1977–78. 1. Description of the epizootic and virological studies. Trans. R. Soc. Trop. Med. Hyg. 73, 618–623.
J. Näslund et al. / Virology 385 (2009) 409–415 Metwally, S., 2008. In: Brown, C., Torres, A. (Eds.), Foreign Animal Diseases, 7th ed. Boca Publications Group, Inc., Boca Raton, FL, pp. 369–376. Morrill, J.C., Mebus, C.A., Peters, C.J., 1997. Safety and efficacy of a mutagen-attenuated Rift Valley fever virus vaccine in cattle. Am J Vet Res. 58 (10), 1104–1109 PMID: 9328662. Muller, R., Saluzzo, J., Lopez, N., Dreier, T., Turell, M., Smith, J., Bouloy, M., 1995. Characterization of clone 13, a naturally attenuated avirulent isolate of Rift Valley fever virus, which is altered in the small segment. Am. J. Trop. Med. Hyg. 53, 405–411. Näslund, J., Lagerqvist, N., Lundkvist, Å., Evander, M., Ahlm, C., Bucht, G., 2008. Kinetics of Rift Valley Fever Virus in experimentally infected mice using quantitative realtime RT-PCR. J. Virol. Methods 151, 277–282. Pittman, P., Liu, C., Cannon, T., Makuch, R., Mangiafico, J., Gibbs, P., Peters, C.J., 1999. Immunogenicity of an inactivated Rift Valley fever vaccine in humans: a 12-year experience. Vaccine 18, 181–189. Pushko, P., Tumpey, T., Bu, F., Knell, J., Robinson, R., Smith, G., 2005. Influenza virus-like particles comprised of the HA, NA, and M1 proteins of H9N2 influenza virus induce protective immune responses in BALB/c mice. Vaccine 23, 5751–5759. Reading, S., Dimmock, N., 2007. Neutralization of animal virus infectivity by antibody. Arch. Virol. 152, 1047–1059. Sall, A., Zanotto, P., Sene, O., Zeller, H., Digoutte, J., Thiongane, Y., Bouloy, M., 1999. Genetic reassortment of Rift Valley fever virus in nature. J. Virol. 73, 8196–8200. Siam, A., Meegan, J., Gharbawi, K., 1980. Rift Valley fever ocular manifestations: observations during the 1977 epidemic in Egypt. Br. J. Ophthalmol. 64, 366–374. Smithburn, K.C., 1949. Rift Valley Fever: the neurotropic adaptation of the virus and the experimental use of this modified virus as a vaccine. Br. J. Exp. Pathol. 1, 1–16. Swenson, D., Warfield, K., Negley, D., Schmaljohn, A., Aman, M., Bavari, S., 2005. Viruslike particles exhibit potential as a pan-filovirus vaccine for both Ebola and Marburg viral infections. Vaccine 23, 3033–3042.
415
van Oirschot, J., 1999. Diva vaccines that reduce virus transmission. J. Biotechnol. 73, 195–205. Vialat, P., Muller, R., Vu, T., Prehaud, C., Bouloy, M., 1997. Mapping of the mutations present in the genome of the Rift Valley fever virus attenuated MP12 strain and their putative role in attenuation. Virus Res. 52, 43–50. Vieira, H., Estêvão, C., Roldão, A., Peixoto, C., Sousa, M., Cruz, P., Carrondo, M., Alves, P., 2005. Triple layered rotavirus VLP production: kinetics of vector replication, mRNA stability and recombinant protein production. J. Biotechnol. 120, 72–82. Villa, L., Costa, R., Petta, C., Andrade, R., Ault, K., Giuliano, A., Wheeler, C., Koutsky, L., Malm, C., Lehtinen, M., Skjeldestad, F., Olsson, S., Steinwall, M., Brown, D., Kurman, R., Ronnett, B., Stoler, M., Ferenczy, A., Harper, D., Tamms, G., Yu, J., Lupinacci, L., Railkar, R., Taddeo, F., Jansen, K., Esser, M., Sings, H., Saah, A., Barr, E., 2005. Prophylactic quadrivalent human papillomavirus (types 6, 11, 16, and 18) L1 viruslike particle vaccine in young women: a randomised double-blind placebocontrolled multicentre phase II efficacy trial. Lancet Oncol. 6, 271–278. Warfield, K., Swenson, D., Negley, D., Schmaljohn, A., Aman, M., Bavari, S., 2004. Marburg virus-like particles protect guinea pigs from lethal Marburg virus infection. Vaccine 22, 3495–3502. Won, S., Ikegami, T., Peters, C.J., Makino, S., 2007. NSm protein of Rift Valley fever virus suppresses virus-induced apoptosis. J. Virol. 81, 13335–13345. Zaki, A., Coudrier, D., Yousef, A., Fakeeh, M., Bouloy, M., Billecocq, A., 2006. Production of monoclonal antibodies against Rift Valley fever virus Application for rapid diagnosis tests (virus detection and ELISA) in human sera. J. Virol. Methods 131, 34–40. Zhou, J., Sun, X., Stenzel, D., Frazer, I., 1991. Expression of vaccinia recombinant HPV 16 L1 and L2 ORF proteins in epithelial cells is sufficient for assembly of HPV virion-like particles. Virology 185, 251–257.