- freely available
Viruses 2012, 4(10), 2097-2114; doi:10.3390/v4102097
Abstract: The family Arenaviridae, genus Arenavirus, consists of two phylogenetically independent groups: Old World (OW) and New World (NW) complexes. The Lassa and Lujo viruses in the OW complex and the Guanarito, Junin, Machupo, Sabia, and Chapare viruses in the NW complex cause viral hemorrhagic fever (VHF) in humans, leading to serious public health concerns. These viruses are also considered potential bioterrorism agents. Therefore, it is of great importance to detect these pathogens rapidly and specifically in order to minimize the risk and scale of arenavirus outbreaks. However, these arenaviruses are classified as BSL-4 pathogens, thus making it difficult to develop diagnostic techniques for these virus infections in institutes without BSL-4 facilities. To overcome these difficulties, antibody detection systems in the form of an enzyme-linked immunosorbent assay (ELISA) and an indirect immunofluorescence assay were developed using recombinant nucleoproteins (rNPs) derived from these viruses. Furthermore, several antigen-detection assays were developed. For example, novel monoclonal antibodies (mAbs) to the rNPs of Lassa and Junin viruses were generated. Sandwich antigen-capture (Ag-capture) ELISAs using these mAbs as capture antibodies were developed and confirmed to be sensitive and specific for detecting the respective arenavirus NPs. These rNP-based assays were proposed to be useful not only for an etiological diagnosis of VHFs, but also for seroepidemiological studies on VHFs. We recently developed arenavirus neutralization assays using vesicular stomatitis virus (VSV)-based pseudotypes bearing arenavirus recombinant glycoproteins. The goal of this article is to review the recent advances in developing laboratory diagnostic assays based on recombinant viral proteins for the diagnosis of VHFs and epidemiological studies on the VHFs caused by arenaviruses.
The virus family Arenaviridae consists of only one genus, but most viruses within this genus can be divided into two different groups: the Old World arenaviruses and the New World arenaviruses (also known as the Tacaribe complex) [1,2]. The differences between the two groups have been established through the use of serological assays. Most of the arenaviruses cause persistent infection in rodents without any symptoms, and humans acquire a variety of diseases when zoonotically infected. Lymphocytic choriomeningitis virus (LCMV) is the only arenavirus to exhibit a worldwide distribution, and causes illnesses such as meningitis [3,4]. Congenital LCMV infections have also been reported [4,5]. Most importantly, viral hemorrhagic fever (VHF) can be caused by several arenaviruses. Lassa fever, caused by the Lassa virus (LASV), an Old World arenavirus, is one of the most devastating VHFs in humans . Hemorrhaging and organ failure occur in a subset of patients infected with this virus, and it is associated with high mortality. Many cases of Lassa fever occur in Western Africa in countries such as Guinea, Sierra Leone, and Nigeria [7,8,9,10,11,12,13]. Tacaribe complex lineage B of the New World arenaviruses consists of the Junin virus (JUNV), Guanarito virus (GUNV), Sabia virus (SABV) and Machupo virus (MACV), the etiological agents of Argentine, Venezuelan, Brazilian, and Bolivian hemorrhagic fevers, respectively [14,15]. Although genetically distinct from one another, they appear to produce similar symptoms, accompanied by hemorrhaging in humans [14,15]. These pathogenic New World arenavirus species are closely associated with a specific rodent species .
Humans are usually infected with pathogenic arenaviruses through direct contact with tissue or blood, or after inhaling aerosolized particles from urine, feces, and saliva of infected rodents. After an incubation period of 1–3 weeks, infected individuals abruptly develop fever, retrosternal pain, sore throat, back pain, cough, abdominal pain, vomiting, diarrhea, conjunctivitis, facial swelling, proteinuria, and mucosal bleeding. Neurological problems have also been described, including hearing loss, tremors, and encephalitis. Because the symptoms of pathogenic arenavirus-related illness are varied and nonspecific, the clinical diagnosis is often difficult [14,16]. Human-to-human transmission may occur via mucosal or cutaneous contact, or through nosocomial contamination [14,16]. These viruses are also considered to be potential bioterrorism agents .
A number of arenavirus species have been recently discovered as a result of both rodent surveys and disease outbreaks [17,18,19,20,21,22,23,24,25,26]. A novel pathogenic New World arenavirus, Chapare virus (CHPV), has been isolated from a fatal case of VHF in Bolivia . In addition, five cases of VHF have been reported in South Africa, and a novel arenavirus, named Lujo virus, was isolated from a patient . The Lujo virus is most distantly related to the other Old World arenaviruses . To date, there is no information concerning the vertebrate host for the Chapare and Lujo viruses.
There is some evidence of endemicity of the Lassa virus in neighboring countries [27,28]. However, as the magnitude of international trade and travel is continuously increasing, and the perturbation of the environment (due either to human activity or natural ecological changes) may result in behavioral changes of reservoir rodents, highly pathogenic arenaviruses could be introduced to virus-free countries from endemic areas. In fact, more than twenty cases of Lassa fever have been reported outside of the endemic region in areas such as the USA, Canada, Europe, and Japan [29,30,31,32,33]. It is of great importance to detect these pathogens rapidly and specifically in order to minimize the risk and scale of outbreaks of VHFs caused by arenaviruses. However, these arenaviruses are classified as biosafety level (BSL)-4 pathogens, making it difficult to develop diagnostic techniques for these virus infections in laboratories without BSL-4 facilities. To overcome these difficulties, we have established recombinant viral nucleoproteins (rNPs)-based serological assays, such as IgG-enzyme-linked immunosorbent assay (ELISA), indirect immunofluorescence assay (IFA), and antigen (Ag)-capture ELISA for the diagnosis of VHFs caused by highly pathogenic arenaviruses. Furthermore, virus neutralization assays using pseudotype virus-bearing arenavirus GPs have been developed. In this review, we describe the usefulness of such recombinant protein-based diagnostic assays for diagnosing VHFs caused by arenaviruses.
2. Currently Used Diagnostic Techniques for VHFs
In outbreaks of VHFs, infections are confirmed by various laboratory diagnostic methods. Virus detection is performed by virus isolation, reverse transcription-polymerase chain reaction (RT-PCR), and antigen-capture ELISA. It has been shown that monoclonal antibody panels against pathogenic arenaviruses are useful for detecting viral antigens on the virus-infected cells as well as for investigating of antigenic relationships of arenaviruses [34,35,36]. Detection of the virus genome is suitable for a rapid and sensitive diagnosis of VHF patients in the early stage of illness, and extensive reviews of such RT-PCR assays have been described [37,38]. More recently, progress in the RT-PCR method covering genetic variations of the hemorrhagic fever viruses (HFVs) [39,40] and a multiplexed oligonucleotide microarray for the differential diagnosis of VHFs have also been reported . On the other hand, antibodies against these viruses can be detected by the indirect immunofluorescence assay (IFA), or IgG- and IgM-ELISA. An IFA detects the antibody in the serum, which is able to bind to the fixed monolayer of the virus-infected cells. Although the interpretation of immunofluorescence results requires experience, the assay has advantages over other methods, since each virus generates a characteristic fluorescence pattern that adds specificity to the assay compared to a simple ELISA readout. A serological diagnosis by the detection of specific IgM and IgG antibodies to the HFVs must be sensitive, specific and reliable, because a misdiagnosis can lead to panic in the general population. An IgM-specific ELISA is suitable for detecting recent infection, but the relevance of IgM testing for acute VHF depends on the virus and the duration of illness; specific IgM is not often present in the very early stage of illness, and patients who die of VHF often fail to seroconvert at all. An IgG-specific ELISA is efficacious, not only in the diagnosis of a large number of VHF cases, especially during convalescence, but also for epidemiological studies in the endemic regions. The detailed methods used for the IFA and IgG- and IgM-ELISAs for the diagnosis of VHF using authentic virus-antigens have been described in detail [42,43,44,45].
4. Antigen-Capture ELISA
For the diagnosis of many viral infections, PCR assays have been shown to have an excellent analytical sensitivity, but the established techniques are limited by their requirement for expensive equipment and technical expertise. Moreover, the high degree of genetic variability of the RNA viruses, including arenavirus and bunyavirus, poses difficulties in selecting primers for RT-PCR assays that can detect all strains of the virus. Since the sensitivity of the Ag-capture ELISA is comparable to that of RT-PCR for several virus-mediated infectious diseases, including Lassa fever and filovirus hemorrhagic fever [51,71,72,73], the Ag-capture ELISA is a sophisticated approach that can be used for the diagnosis of viral infections. Ag-capture ELISAs detecting viral NP in viremic sera have been widely applied to detect various viruses, since they are the most abundant viral antigens and have highly conserved amino acid sequences [50,51,54,71,72,74,75]. Polyclonal anti-sera or a mixture of MAbs present in the ascetic fluids from animals immunized for HFVs have been used for capture-antibodies in the Ag-capture ELISA [36,76,77,78,79]. MAbs recognizing conserved epitopes of the rNP are also used as capture antibodies since they have a high specificity for the antigens, and an identification of the epitopes of these MAbs is of crucial importance for the assessment of the specificity and cross-reactivity of the assay system [50,51,53,54,71,75]. In order to develop a sensitive diagnostic test for Lassa fever and AHF, rNPs of LASV and JUNV (see above) have been prepared, and newly established MAbs against them have been characterized and used for Ag-capture ELISAs [50,51]. The Ag-capture ELISA using MAb 4A5 has been confirmed to be useful in the detection of authentic LASV antigen in sera serially collected from hamsters infected with LASV . The sensitivity of the MAb 4A5-based Ag-capture ELISA was similar to that of conventional RT-PCR, suggesting that the Ag-capture ELISA can be efficiently used in the diagnosis of Lassa fever . Therefore, the MAb 4A5- based Ag-capture ELISA is considered to be useful in the diagnosis of Lassa fever. Also, by using MAbs raised against the rNP of JUNV, Ag-capture ELISAs specific for JUNV and broadly reactive to human pathogenic New World arenaviruses have been developed . The Ag-capture ELISA using MAb E4-2 and C11-12 detected the Ags of all of the pathogenic New World arenaviruses tested, including JUNV. On the other hand, the Ag-capture ELISA using MAb C6-9 detects only the JUNV Ag. Considering that the symptoms of JUNV infection in humans are indistinguishable from those due to other pathogenic New World arenaviruses, the Ag capture ELISA using MAb C6-9 may be a useful diagnostic tool, especially for AHF .
5. Neutralization Assays Based on VSV Pseudotypes
The virus neutralization assay is accepted as the “gold standard” serodiagnostic assay to quantify the antibody response to infection and vaccination of a wide variety of viruses associated with human diseases [80,81,82,83,84,85,86]. The presence of neutralizing antibodies is a reliable indicator of protective immunity against VHF [87,88,89]. The most direct method for detection of neutralizing antibodies against HFVs is by plaque reduction neutralization tests using infectious viruses. However, because of the high pathogenicity of HFVs to humans and the strict regulation of select agents, only a limited number of laboratories are able to perform such neutralization tests. For many HFVs, replication-incompetent pseudotyped virus particles bearing viral envelope protein (GP) have been shown to mimic the respective HFV infections, thus, neutralization assays using the pseudotypes may be advantageous in some laboratory settings for the detection of antibodies to HFVs without the need for heightened biocontainment requirements.
The VSV-based vector has already been used to generate replication-competent recombinant VSVs to study of the role of GPs of various viruses [90,91,92]. Recent advances in producing pseudotype virus particles have enabled the investigation of the virus cell entry, viral tropism, and effect of entry inhibitors, as well as measurement of the neutralization titers, by using human immunodeficiency virus-, feline immunodeficiency virus-, murine leukemia virus-, or VSV-based vectors [86,93,94,95,96,97,98,99,100,101,102,103]. Pseudotypes based on VSV have advantages compared with other pseudotypes based on retroviruses for the following reasons. First, the pseudotype virus titer obtained with the VSV system is generally higher than that of the pseudotyped retrovirus system . Second, the infection of target cells with a VSV pseudotype can be readily detected as green fluorescent protein (GFP)-positive cells at 7–16 h post-infection because of the high level of GFP expression in the VSV system [104,105]. In contrast, the time required for infection in the pseudotyped retrovirus system is 48 h [106,107], which is similar to the time required for infectious viruses to replicate to a level that results in plaque-forming or cytopathic effects in infected cells. A high-throughput assay for determining neutralizing antibody titers using VSV pseudotypes expressing secreted alkaline phosphatase [108,109] or luciferase (Figure 3) has also been developed.
We have recently developed a VSV-based pseudotype bearing Lassa virus GP (VSV-LAS-GP) for the detection of neutralizing antibodies in the sera obtained from a Lassa fever patient. An example of the LASV neutralization assay using the VSV pseudotype is shown (Figure 4). In the presence of serum from Lassa fever patients, the number of GFP-positive cells (infectivity of VSV-LAS-GP) is significantly reduced compared with the number in the absence of the patient’s serum (Figure 4A). The control VSV pseudotype bearing VSV GP (VSV-VSV-G) is not neutralized by any sera. When the cut‑off serum dilution is set at 50% inhibition of infectivity compared with the infectivity in the absence of the test serum, the neutralization titer of this patient’s serum for VSV-LAS-GP is calculated to be 75 (Figure 4B). Likewise, a VSV-based pseudotype bearing the Junin virus GP has been developed for the detection of neutralizing antibodies from AHF patients’ sera. The accuracy of the results of VSV-based neutralization assays has been confirmed by comparison with the results of the neutralization assay using live Junin virus .
The Lujo virus is a new member of the hemorrhagic fever-associated arenavirus family from Zambia and southern Africa, and the virus is classified as a BSL-4 pathogen . The genome sequence analysis of the Lujo virus suggests that the virus is genetically distinct from previously characterized arenaviruses. In order to study the infectivity of this newly identified arenavirus, we have recently developed a luciferase-expressing VSV pseudotype bearing Lujo virus GPC (VSV-Lujo-GP). As shown in Figure 3, infection with VSV-Lujo-GPC is specifically neutralized by rabbit anti-Lujo GPC serum. Thus, the VSV-Lujo-GP may be a useful tool not only for determining the neutralizing antibody titer within the serum, but also for exploring yet-to-be-defined cellular receptor(s) for Lujo virus infection or for screening inhibitors of the Lujo virus GP-mediated cell entry.
Hemorrhagic fever outbreaks caused by pathogenic arenaviruses result in high fatality rates. A rapid and accurate diagnosis is a critical first step in any outbreak. Serologic diagnostic methods for VHFs most often employ an ELISA, IFA, and/or virus neutralization assay. Diagnostic methods using recombinant viral proteins have been developed and their utilities for diagnosing of VHF have been reviewed. IgG- and IgM- ELISAs and IFAs using rNPs as antigens are useful for the detection of antibodies induced in the patients’ sera. These methods are also useful for seroepidemiological surveys for HFVs. Ag-capture ELISAs using MAbs to the arenavirus rNPs are specific for the virus species or can be broadly reactive for New World arenaviruses, depending on the MAb used. Furthermore, the VSV-based pseudotype system provides a safe and rapid tool for measuring virus neutralizing antibody titers, as well as a model to analyze the entry of the respective arenavirus in susceptible cells without using live arenaviruses. Recent discoveries of novel arenavirus species [17,26,110] and their potential to evolve predominantly via host switching, rather than with their hosts [110,111], suggest that an unknown pathogenic arenavirus may emerge in the future, and that the diagnostic methods for VHF caused by arenaviruses should thus be further developed and improved.
Conflict of Interest
The authors declare no conflict of interest.
We thank the staff of the Special Pathogens Laboratory, Department of Virology 1, NIID, I. Kurane, T. Mizutani, M. Niikura, A. Maeda, T. Ikegami, and M. Ogata, for their contributions to these studies on the development of diagnostic methods. We also thank M. Whitt for providing the VSV pseudotype system. This work was supported in part by a grant-in aid from the Ministry of Health, Labor and Welfare of Japan (H22-Shinkou-Ippan-006 and H24-Shinkou-Ippan-013).
References and Notes
- Pfau, C.J.; Bergold, G.H.; Casals, J.; Johnson, K.M.; Murphy, F.A.; Pedersen, I.R.; Rawls, W.E.; Rowe, W.P.; Webb, P.A.; Weissenbacher, M.C. Arenaviruses. Intervirology 1974, 4, 207–214. [Google Scholar] [CrossRef]
- Charrel, R.N.; de Lamballerie, X. Arenaviruses other than Lassa virus. Antivir. Res. 2003, 57, 89–100. [Google Scholar]
- Jahrling, P.B.; Peters, C.J. Lymphocytic choriomeningitis virus. A neglected pathogen of man. Arch. Pathol. Lab. Med. 1992, 116, 486–488. [Google Scholar]
- Barton, L.L.; Mets, M.B.; Beauchamp, C.L. Lymphocytic choriomeningitis virus: Emerging fetal teratogen. Am. J. Obstet. Gynecol. 2002, 187, 1715–1716. [Google Scholar] [CrossRef]
- Wright, R.; Johnson, D.; Neumann, M.; Ksiazek, T.G.; Rollin, P.; Keech, R.V.; Bonthius, D.J.; Hitchon, P.; Grose, C.F.; Bell, W.E.; et al. Congenital lymphocytic choriomeningitis virus syndrome: A disease that mimics congenital toxoplasmosis or Cytomegalovirus infection. Pediatrics 1997, 100, E9. [Google Scholar]
- Salazar-Bravo, J.; Ruedas, L.A.; Yates, T.L. Mammalian reservoirs of arenaviruses. Curr. Top. Microbiol. Immunol. 2002, 262, 25–63. [Google Scholar]
- Carey, D.E.; Kemp, G.E.; White, H.A.; Pinneo, L.; Addy, R.F.; Fom, A.L.; Stroh, G.; Casals, J.; Henderson, B.E. Lassa fever. Epidemiological aspects of the 1970 epidemic, Jos, Nigeria. Trans. R. Soc. Trop. Med. Hyg. 1972, 66, 402–408. [Google Scholar] [CrossRef]
- Lukashevich, I.S.; Clegg, J.C.; Sidibe, K. Lassa virus activity in Guinea: Distribution of human antiviral antibody defined using enzyme-linked immunosorbent assay with recombinant antigen. J. Med. Virol. 1993, 40, 210–217. [Google Scholar] [CrossRef]
- McCormick, J.B.; Webb, P.A.; Krebs, J.W.; Johnson, K.M.; Smith, E.S. A prospective study of the epidemiology and ecology of Lassa fever. J. Infect. Dis. 1987, 155, 437–444. [Google Scholar] [CrossRef]
- Monath, T.P. Lassa fever: Review of epidemiology and epizootiology. Bull. World Health Organ. 1975, 52, 577–592. [Google Scholar]
- Monath, T.P. Lassa fever: A new appraisal. Niger. Med. J. 1973, 3, 162–163. [Google Scholar]
- Monson, M.H.; Frame, J.D.; Jahrling, P.B.; Alexander, K. Endemic Lassa fever in Liberia. I. Clinical and epidemiological aspects at Curran Lutheran Hospital, Zorzor, Liberia. Trans. R. Soc. Trop. Med. Hyg. 1984, 78, 549–553. [Google Scholar] [CrossRef]
- Ehichioya, D.U.; Hass, M.; Becker-Ziaja, B.; Ehimuan, J.; Asogun, D.A.; Fichet-Calvet, E.; Kleinsteuber, K.; Lelke, M.; ter Meulen, J.; Akpede, G.O.; et al. Current molecular epidemiology of Lassa virus in Nigeria. J. Clin. Microbiol. 2011, 49, 1157–1161. [Google Scholar] [CrossRef]
- Peters, C.J. Human infection with arenaviruses in the Americas. Curr. Top. Microbiol. Immunol. 2002, 262, 65–74. [Google Scholar]
- Tesh, R.B. Viral hemorrhagic fevers of South America. Biomedica 2002, 22, 287–295. [Google Scholar]
- McCormick, J.B.; Fisher-Hoch, S.P. Lassa fever. Curr. Top. Microbiol. Immunol. 2002, 262, 75–109. [Google Scholar]
- Briese, T.; Paweska, J.T.; McMullan, L.K.; Hutchison, S.K.; Street, C.; Palacios, G.; Khristova, M.L.; Weyer, J.; Swanepoel, R.; Egholm, M.; et al. Genetic detection and characterization of Lujo virus, a new hemorrhagic fever-associated arenavirus from southern Africa. PLoS Pathog. 2009, 5, e1000455. [Google Scholar] [CrossRef]
- Cajimat, M.N.; Milazzo, M.L.; Bradley, R.D.; Fulhorst, C.F. Catarina virus, an arenaviral species principally associated with Neotoma micropus (southern plains woodrat) in Texas. Am. J. Trop. Med. Hyg. 2007, 77, 732–736. [Google Scholar]
- Cajimat, M.N.; Milazzo, M.L.; Borchert, J.N.; Abbott, K.D.; Bradley, R.D.; Fulhorst, C.F. Diversity among Tacaribe serocomplex viruses (family Arenaviridae) naturally associated with the Mexican woodrat (Neotoma mexicana). Virus Res. 2008, 133, 211–217. [Google Scholar] [CrossRef]
- Delgado, S.; Erickson, B.R.; Agudo, R.; Blair, P.J.; Vallejo, E.; Albarino, C.G.; Vargas, J.; Comer, J.A.; Rollin, P.E.; Ksiazek, T.G.; et al. Chapare virus, a newly discovered arenavirus isolated from a fatal hemorrhagic fever case in Bolivia. PLoS Pathog. 2008, 4, e1000047. [Google Scholar] [CrossRef]
- Gunther, S.; Hoofd, G.; Charrel, R.; Roser, C.; Becker-Ziaja, B.; Lloyd, G.; Sabuni, C.; Verhagen, R.; van der Groen, G.; Kennis, J.; et al. Mopeia virus-related arenavirus in natal multimammate mice, Morogoro, Tanzania. Emerg. Infect. Dis. 2009, 15, 2008–2012. [Google Scholar] [CrossRef]
- Lecompte, E.; ter Meulen, J.; Emonet, S.; Daffis, S.; Charrel, R.N. Genetic identification of Kodoko virus, a novel arenavirus of the African pigmy mouse (Mus Nannomys minutoides) in West Africa. Virology 2007, 364, 178–183. [Google Scholar] [CrossRef]
- Milazzo, M.L.; Cajimat, M.N.; Haynie, M.L.; Abbott, K.D.; Bradley, R.D.; Fulhorst, C.F. Diversity among tacaribe serocomplex viruses (family Arenaviridae) naturally associated with the white-throated woodrat (Neotoma albigula) in the southwestern United States. Vector Borne Zoonotic Dis. 2008, 8, 523–450. [Google Scholar] [CrossRef]
- Palacios, G.; Druce, J.; Du, L.; Tran, T.; Birch, C.; Briese, T.; Conlan, S.; Quan, P.L.; Hui, J.; Marshall, J.; et al. A new arenavirus in a cluster of fatal transplant-associated diseases. N. Engl. J. Med. 2008, 358, 991–998. [Google Scholar] [CrossRef]
- Palacios, G.; Savji, N.; Hui, J.; Travassos da Rosa, A.; Popov, V.; Briese, T.; Tesh, R.; Lipkin, W.I. Genomic and phylogenetic characterization of Merino Walk virus, a novel arenavirus isolated in South Africa. J. Gen. Virol. 2010, 91, 1315–1324. [Google Scholar] [CrossRef]
- Ishii, A.; Thomas, Y.; Moonga, L.; Nakamura, I.; Ohnuma, A.; Hang'ombe, B.; Takada, A.; Mweene, A.; Sawa, H. Novel arenavirus, Zambia. Emerg. Infect. Dis. 2011, 17, 1921–1924. [Google Scholar] [CrossRef]
- Frame, J.D. Surveillance of Lassa fever in missionaries stationed in West Africa. Bull. World Health Organ. 1975, 52, 593–598. [Google Scholar]
- Richmond, J.K.; Baglole, D.J. Lassa fever: Epidemiology, clinical features, and social consequences. BMJ 2003, 327, 1271–1275. [Google Scholar] [CrossRef]
- Gunther, S.; Emmerich, P.; Laue, T.; Kuhle, O.; Asper, M.; Jung, A.; Grewing, T.; ter Meulen, J.; Schmitz, H. Imported lassa fever in Germany: Molecular characterization of a new lassa virus strain. Emerg. Infect. Dis. 2000, 6, 466–476. [Google Scholar] [CrossRef]
- Hirabayashi, Y.; Oka, S.; Goto, H.; Shimada, K.; Kurata, T.; Fisher-Hoch, S.P.; McCormick, J.B. An imported case of Lassa fever with late appearance of polyserositis. J. Infect. Dis. 1988, 158, 872–875. [Google Scholar]
- Jeffs, B. A clinical guide to viral haemorrhagic fevers: Ebola, Marburg and Lassa. Trop. Doct. 2006, 36, 1–4. [Google Scholar] [CrossRef]
- Macher, A.M.; Wolfe, M.S. Historical Lassa fever reports and 30-year clinical update. Emerg. Infect. Dis. 2006, 12, 835–837. [Google Scholar] [CrossRef]
- Mahdy, M.S.; Chiang, W.; McLaughlin, B.; Derksen, K.; Truxton, B.H.; Neg, K. Lassa fever: The first confirmed case imported into Canada. Can. Dis. Wkly. Rep. 1989, 15, 193–198. [Google Scholar]
- Buchmeier, M.J.; Lewicki, H.A.; Tomori, O.; Oldstone, M.B. Monoclonal antibodies to lymphocytic choriomeningitis and pichinde viruses: Generation, characterization, and cross-reactivity with other arenaviruse. Virology 1981, 113, 73–85. [Google Scholar] [CrossRef]
- Ruo, S.L.; Mitchell, S.W.; Kiley, M.P.; Roumillat, L.F.; Fisher-Hoch, S.P.; McCormick, J.B. Antigenic relatedness between arenaviruses defined at the epitope level by monoclonal antibodies. J. Gen. Virol. 1991, 72, 549–555. [Google Scholar] [CrossRef]
- Sanchez, A.; Pifat, D.Y.; Kenyon, R.H.; Peters, C.J.; McCormick, J.B.; Kiley, M.P. Junin virus monoclonal antibodies: Characterization and cross-reactivity with other arenaviruses. J. Gen. Virol. 1989, 70, 1125–1132. [Google Scholar] [CrossRef]
- Charrel, R.N.; de Lamballerie, X. Zoonotic aspects of arenavirus infections. Vet. Microbiol. 2010, 140, 213–220. [Google Scholar] [CrossRef]
- Drosten, C.; Kummerer, B.M.; Schmitz, H.; Gunther, S. Molecular diagnostics of viral hemorrhagic fevers. Antivir. Res. 2003, 57, 61–87. [Google Scholar]
- Ogawa, H.; Miyamoto, H.; Ebihara, H.; Ito, K.; Morikawa, S.; Feldmann, H.; Takada, A. Detection of all known filovirus species by reverse transcription-polymerase chain reaction using a primer set specific for the viral nucleoprotein gene. J. Virol. Meth. 2011, 171, 310–313. [Google Scholar] [CrossRef]
- Olschlager, S.; Lelke, M.; Emmerich, P.; Panning, M.; Drosten, C.; Hass, M.; Asogun, D.; Ehichioya, D.; Omilabu, S.; Gunther, S. Improved detection of Lassa virus by reverse transcription-PCR targeting the 5' region of S RNA. J. Clin. Microbiol. 2010, 48, 2009–2013. [Google Scholar] [CrossRef]
- Palacios, G.; Quan, P.L.; Jabado, O.J.; Conlan, S.; Hirschberg, D.L.; Liu, Y.; Zhai, J.; Renwick, N.; Hui, J.; Hegyi, H.; et al. Panmicrobial oligonucleotide array for diagnosis of infectious diseases. Emerg. Infect. Dis. 2007, 13, 73–81. [Google Scholar] [CrossRef]
- Ksiazek, T.G.; Rollin, P.E.; Williams, A.J.; Bressler, D.S.; Martin, M.L.; Swanepoel, R.; Burt, F.J.; Leman, P.A.; Khan, A.S.; Rowe, A.K.; et al. Clinical virology of Ebola hemorrhagic fever (EHF): Virus, virus antigen, and IgG and IgM antibody findings among EHF patients in Kikwit, Democratic Republic of the Congo, 1995. J. Infect. Dis. 1999, 179, S177–S187. [Google Scholar]
- Ksiazek, T.G.; West, C.P.; Rollin, P.E.; Jahrling, P.B.; Peters, C.J. ELISA for the detection of antibodies to Ebola viruses. J. Infect. Dis. 1999, 179, S192–S198. [Google Scholar] [CrossRef]
- Bausch, D.G.; Rollin, P.E.; Demby, A.H.; Coulibaly, M.; Kanu, J.; Conteh, A.S.; Wagoner, K.D.; McMullan, L.K.; Bowen, M.D.; Peters, C.J.; et al. Diagnosis and clinical virology of Lassa fever as evaluated by enzyme-linked immunosorbent assay, indirect fluorescent-antibody test, and virus isolation. J. Clin. Microbiol. 2000, 38, 2670–2607. [Google Scholar]
- Emmerich, P.; Thome-Bolduan, C.; Drosten, C.; Gunther, S.; Ban, E.; Sawinsky, I.; Schmitz, H. Reverse ELISA for IgG and IgM antibodies to detect Lassa virus infections in Africa. J. Clin. Virol. 2006, 37, 277–281. [Google Scholar] [CrossRef]
- Meyer, B.J.; de la Torre, J.C.; Southern, P.J. Arenaviruses: Genomic RNAs, transcription, and replication. Curr. Top. Microbiol. Immunol. 2002, 262, 139–157. [Google Scholar]
- Ter Meulen, J.; Koulemou, K.; Wittekindt, T.; Windisch, K.; Strigl, S.; Conde, S.; Schmitz, H. Detection of Lassa virus antinucleoprotein immunoglobulin G (IgG) and IgM antibodies by a simple recombinant immunoblot assay for field use. J. Clin. Microbiol. 1998, 36, 3143–3148. [Google Scholar]
- Takimoto, K.; Taharaguchi, M.; Morikawa, S.; Ike, F.; Yamada, Y.K. Detection of the antibody to lymphocytic choriomeningitis virus in sera of laboratory rodents infected with viruses of laboratory and newly isolated strains by ELISA using purified recombinant nucleoprotein. Exp. Anim. 2008, 57, 357–365. [Google Scholar] [CrossRef]
- Ure, A.E.; Ghiringhelli, P.D.; Possee, R.D.; Morikawa, S.; Romanowski, V. Argentine hemorrhagic fever diagnostic test based on recombinant Junin virus N protein. J. Med. Virol. 2008, 80, 2127–2133. [Google Scholar] [CrossRef]
- Nakauchi, M.; Fukushi, S.; Saijo, M.; Mizutani, T.; Ure, A.E.; Romanowski, V.; Kurane, I.; Morikawa, S. Characterization of monoclonal antibodies to Junin virus nucleocapsid protein and application to the diagnosis of hemorrhagic fever caused by South American arenaviruses. Clin. Vaccine Immunol. 2009, 16, 1132–1138. [Google Scholar] [CrossRef]
- Saijo, M.; Georges-Courbot, M.C.; Marianneau, P.; Romanowski, V.; Fukushi, S.; Mizutani, T.; Georges, A.J.; Kurata, T.; Kurane, I.; Morikawa, S. Development of recombinant nucleoprotein-based diagnostic systems for Lassa fever. Clin. Vaccine Immunol. 2007, 14, 1182–1189. [Google Scholar] [CrossRef]
- Saijo, M.; Niikura, M.; Morikawa, S.; Ksiazek, T.G.; Meyer, R.F.; Peters, C.J.; Kurane, I. Enzyme-linked immunosorbent assays for detection of antibodies to Ebola and Marburg viruses using recombinant nucleoproteins. J. Clin. Microbiol. 2001, 39, 1–7. [Google Scholar] [CrossRef]
- Ikegami, T.; Niikura, M.; Saijo, M.; Miranda, M.E.; Calaor, A.B.; Hernandez, M.; Acosta, L.P.; Manalo, D.L.; Kurane, I.; Yoshikawa, Y.; et al. Antigen capture enzyme-linked immunosorbent assay for specific detection of Reston Ebola virus nucleoprotein. Clin. Diagn. Lab. Immunol. 2003, 10, 552–557. [Google Scholar]
- Saijo, M.; Tang, Q.; Shimayi, B.; Han, L.; Zhang, Y.; Asiguma, M.; Tianshu, D.; Maeda, A.; Kurane, I.; Morikawa, S. Antigen-capture enzyme-linked immunosorbent assay for the diagnosis of crimean-congo hemorrhagic fever using a novel monoclonal antibody. J. Med. Virol. 2005, 77, 83–88. [Google Scholar] [CrossRef]
- Saijo, M.; Qing, T.; Niikura, M.; Maeda, A.; Ikegami, T.; Prehaud, C.; Kurane, I.; Morikawa, S. Recombinant nucleoprotein-based enzyme-linked immunosorbent assay for detection of immunoglobulin G antibodies to Crimean-Congo hemorrhagic fever virus. J. Clin. Microbiol. 2002, 40, 1587–1591. [Google Scholar] [CrossRef]
- Matsuura, Y.; Possee, R.D.; Overton, H.A.; Bishop, D.H. Baculovirus expression vectors: The requirements for high level expression of proteins, including glycoproteins. J. Gen. Virol. 1987, 68, 1233–1250. [Google Scholar] [CrossRef]
- Tang, Q.; Saijo, M.; Zhang, Y.; Asiguma, M.; Tianshu, D.; Han, L.; Shimayi, B.; Maeda, A.; Kurane, I.; Morikawa, S. A patient with Crimean-Congo hemorrhagic fever serologically diagnosed by recombinant nucleoprotein-based antibody detection systems. Clin. Diagn. Lab. Immunol. 2003, 10, 489–491. [Google Scholar]
- Saijo, M. National Institute of Infectious Diseases, Tokyo, Japan. Unpublished work, 2012..
- van der Groen, G.; Piot, P.; Desmyter, J.; Colaert, J.; Muylle, L.; Tkachenko, E.A.; Ivanov, A.P.; Verhagen, R.; van Ypersele de Strihou, C. Seroepidemiology of Hantaan-related virus infections in Belgian populations. Lancet 1983, 2, 1493–1494. [Google Scholar]
- Morrill, J.C.; Soliman, A.K.; Imam, I.Z.; Botros, B.A.; Moussa, M.I.; Watts, D.M. Serological evidence of Crimean-Congo haemorrhagic fever viral infection among camels imported into Egypt. J. Trop. Med. Hyg. 1990, 93, 201–204. [Google Scholar]
- Emmerich, P.; Gunther, S.; Schmitz, H. Strain-specific antibody response to Lassa virus in the local population of west Africa. J. Clin. Virol. 2008, 42, 40–44. [Google Scholar] [CrossRef]
- Xia, H.; Li, P.; Yang, J.; Pan, L.; Zhao, J.; Wang, Z.; Li, Y.; Zhou, H.; Dong, Y.; Guo, S.; et al. Epidemiological survey of Crimean-Congo hemorrhagic fever virus in Yunnan, China, 2008. Int. J. Infect. Dis. 2011, 15, e459–e463. [Google Scholar] [CrossRef]
- Heinrich, N.; Saathoff, E.; Weller, N.; Clowes, P.; Kroidl, I.; Ntinginya, E.; Machibya, H.; Maboko, L.; Loscher, T.; Dobler, G.; et al. High seroprevalence of Rift Valley FEVER AND EVIDENCE FOR ENDEMIC circulation in Mbeya region, Tanzania, in a cross-sectional study. PLoS Negl. Trop. Dis. 2012, 6, e1557. [Google Scholar] [CrossRef]
- Saijo, M.; Niikura, M.; Morikawa, S.; Kurane, I. Immunofluorescence method for detection of Ebola virus immunoglobulin g, using HeLa cells which express recombinant nucleoprotein. J. Clin. Microbiol. 2001, 39, 776–778. [Google Scholar] [CrossRef]
- Ikegami, T.; Saijo, M.; Niikura, M.; Miranda, M.E.; Calaor, A.B.; Hernandez, M.; Manalo, D.L.; Kurane, I.; Yoshikawa, Y.; Morikawa, S. Development of an immunofluorescence method for the detection of antibodies to Ebola virus subtype Reston by the use of recombinant nucleoprotein-expressing HeLa cells. Microbiol. Immunol. 2002, 46, 633–638. [Google Scholar]
- Sayama, Y.; Demetria, C.; Saito, M.; Azul, R.R.; Taniguchi, S.; Fukushi, S.; Yoshikawa, T.; Iizuka, I.; Mizutani, T.; Kurane, I.; et al. A seroepidemiologic study of Reston ebolavirus in swine in the Philippines. BMC Vet. Res. 2012, 8, 82. [Google Scholar] [CrossRef]
- Taniguchi, S.; Watanabe, S.; Masangkay, J.S.; Omatsu, T.; Ikegami, T.; Alviola, P.; Ueda, N.; Iha, K.; Fujii, H.; Ishii, Y.; et al. Reston Ebolavirus antibodies in bats, the Philippines. Emerg. Infect. Dis. 2011, 17, 1559–1560. [Google Scholar]
- Saijo, M.; Tang, Q.; Shimayi, B.; Han, L.; Zhang, Y.; Asiguma, M.; Tianshu, D.; Maeda, A.; Kurane, I.; Morikawa, S. Recombinant nucleoprotein-based serological diagnosis of Crimean-Congo hemorrhagic fever virus infections. J. Med. Virol. 2005, 75, 295–299. [Google Scholar] [CrossRef]
- Morrison, H.G.; Goldsmith, C.S.; Regnery, H.L.; Auperin, D.D. Simultaneous expression of the Lassa virus N and GPC genes from a single recombinant vaccinia virus. Virus Res. 1991, 18, 231–241. [Google Scholar] [CrossRef]
- Iha, K.; Nakauchi, M.; Taniguchi, S.; Fukushi, S.; Mizutani, T.; Ogata, M.; Saijo, M.; Romanowski, V.; Enria, D.A.; Kyuwa, S.; et al. Establishment of serological diagnosis of Argentine hemorrhagic fever using recombinant antigens. National Institute of Infectious Diseases, Tokyo, Japan. To be submitted for publication, 2012..
- Saijo, M.; Georges-Courbot, M.C.; Fukushi, S.; Mizutani, T.; Philippe, M.; Georges, A.J.; Kurane, I.; Morikawa, S. Marburgvirus nucleoprotein-capture enzyme-linked immunosorbent assay using monoclonal antibodies to recombinant nucleoprotein: Detection of authentic Marburgvirus. Jpn. J. Infect. Dis. 2006, 59, 323–325. [Google Scholar]
- Velumani, S.; Du, Q.; Fenner, B.J.; Prabakaran, M.; Wee, L.C.; Nuo, L.Y.; Kwang, J. Development of an antigen-capture ELISA for detection of H7 subtype avian influenza from experimentally infected chickens. J. Virol. Meth. 2008, 147, 219–225. [Google Scholar] [CrossRef]
- Ji, Y.; Guo, W.; Zhao, L.; Li, H.; Lu, G.; Wang, Z.; Wang, G.; Liu, C.; Xiang, W. Development of an antigen-capture ELISA for the detection of equine influenza virus nucleoprotein. J. Virol. Meth. 2011, 175, 120–124. [Google Scholar] [CrossRef]
- Jansen van Vuren, P.; Potgieter, A.C.; Paweska, J.T.; van Dijk, A.A. 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. Meth. 2007, 140, 106–114. [Google Scholar] [CrossRef]
- Fukushi, S.; Nakauchi, M.; Mizutani, T.; Saijo, M.; Kurane, I.; Morikawa, S. Antigen-capture ELISA for the detection of Rift Valley fever virus nucleoprotein using new monoclonal antibodies. J. Virol. Meth. 2012, 180, 68–74. [Google Scholar] [CrossRef]
- Ksiazek, T.G.; Rollin, P.E.; Jahrling, P.B.; Johnson, E.; Dalgard, D.W.; Peters, C.J. Enzyme immunosorbent assay for Ebola virus antigens in tissues of infected primates. J. Clin. Microbiol. 1992, 30, 947–950. [Google Scholar]
- Logan, T.M.; Linthicum, K.J.; Moulton, J.R.; Ksiazek, T.G. Antigen-capture enzyme-linked immunosorbent assay for detection and quantification of Crimean-Congo hemorrhagic fever virus in the tick, Hyalomma truncatum. J. Virol. Meth. 1993, 42, 33–44. [Google Scholar] [CrossRef]
- Mills, J.N.; Ellis, B.A.; McKee, K.T., Jr.; Ksiazek, T.G.; Oro, J.G.; Maiztegui, J.I.; Calderon, G.E.; Peters, C.J.; Childs, J.E. Junin virus activity in rodents from endemic and nonendemic loci in central Argentina. Am. J. Trop. Med. Hyg. 1991, 44, 589–597. [Google Scholar]
- Niklasson, B.; Peters, C.J.; Grandien, M.; Wood, O. Detection of human immunoglobulins G and M antibodies to Rift Valley fever virus by enzyme-linked immunosorbent assay. J. Clin. Microbiol. 1984, 19, 225–229. [Google Scholar]
- Roehrig, J.T.; Hombach, J.; Barrett, A.D. Guidelines for plaque-reduction neutralization testing of human antibodies to dengue viruses. Viral Immunol. 2008, 21, 123–132. [Google Scholar] [CrossRef]
- Hassantoufighi, A.; Zhang, H.; Sandbulte, M.; Gao, J.; Manischewitz, J.; King, L.; Golding, H.; Straight, T.M.; Eichelberger, M.C. A practical influenza neutralization assay to simultaneously quantify hemagglutinin and neuraminidase-inhibiting antibody responses. Vaccine 2010, 28, 790–797. [Google Scholar] [CrossRef]
- Holzmann, H. Diagnosis of tick-borne encephalitis. Vaccine 2003, 21, S36–S40. [Google Scholar] [CrossRef]
- Daniels, P.; Ksiazek, T.; Eaton, B.T. Laboratory diagnosis of Nipah and Hendra virus infections. Microbes Infect. 2001, 3, 289–295. [Google Scholar] [CrossRef]
- Tomori, O.; Johnson, K.M.; Kiley, M.P.; Elliott, L.H. Standardization of a plaque assay for Lassa virus. J. Med. Virol. 1987, 22, 77–89. [Google Scholar] [CrossRef]
- Alche, L.E.; Coto, C.E. Differentiation of Junin virus and antigenic variants isolated in vivo by kinetic neutralization assays. J. Gen. Virol. 1988, 69, 2123–2127. [Google Scholar] [CrossRef]
- Fukushi, S.; Mizutani, T.; Saijo, M.; Kurane, I.; Taguchi, F.; Tashiro, M.; Morikawa, S. Evaluation of a novel vesicular stomatitis virus pseudotype-based assay for detection of neutralizing antibody responses to SARS-CoV. J. Med. Virol. 2006, 78, 1509–1512. [Google Scholar] [CrossRef]
- Jahrling, P.B. Protection of Lassa virus-infected guinea pigs with Lassa-immune plasma of guinea pig, primate, and human origin. J. Med. Virol. 1983, 12, 93–102. [Google Scholar] [CrossRef]
- Takada, A.; Ebihara, H.; Jones, S.; Feldmann, H.; Kawaoka, Y. Protective efficacy of neutralizing antibodies against Ebola virus infection. Vaccine 2007, 25, 993–999. [Google Scholar] [CrossRef]
- McKee, K.T., Jr.; Oro, J.G.; Kuehne, A.I.; Spisso, J.A.; Mahlandt, B.G. Candid No. 1 Argentine hemorrhagic fever vaccine protects against lethal Junin virus challenge in rhesus macaques. Intervirology 1992, 34, 154–163. [Google Scholar] [CrossRef]
- Kretzschmar, E.; Buonocore, L.; Schnell, M.J.; Rose, J.K. High-efficiency incorporation of functional influenza virus glycoproteins into recombinant vesicular stomatitis viruses. J. Virol. 1997, 71, 5982–5989. [Google Scholar]
- Schnell, M.J.; Buonocore, L.; Kretzschmar, E.; Johnson, E.; Rose, J.K. Foreign glycoproteins expressed from recombinant vesicular stomatitis viruses are incorporated efficiently into virus particles. Proc. Natl. Acad. Sci. U. S. A. 1996, 93, 11359–11365. [Google Scholar]
- Garbutt, M.; Liebscher, R.; Wahl-Jensen, V.; Jones, S.; Moller, P.; Wagner, R.; Volchkov, V.; Klenk, H.D.; Feldmann, H.; Stroher, U. Properties of replication-competent vesicular stomatitis virus vectors expressing glycoproteins of filoviruses and arenaviruses. J. Virol. 2004, 78, 5458–5465. [Google Scholar]
- Fukushi, S.; Mizutani, T.; Sakai, K.; Saijo, M.; Taguchi, F.; Yokoyama, M.; Kurane, I.; Morikawa, S. Amino acid substitutions in the s2 region enhance severe acute respiratory syndrome coronavirus infectivity in rat angiotensin-converting enzyme 2-expressing cells. J. Virol. 2007, 81, 10831–10834. [Google Scholar] [CrossRef]
- Watanabe, R.; Matsuyama, S.; Shirato, K.; Maejima, M.; Fukushi, S.; Morikawa, S.; Taguchi, F. Entry from the cell surface of severe acute respiratory syndrome coronavirus with cleaved S protein as revealed by pseudotype virus bearing cleaved S protein. J. Virol. 2008, 82, 11985–11991. [Google Scholar] [CrossRef]
- Wang, Y.; Keck, Z.Y.; Foung, S.K. Neutralizing antibody response to hepatitis C virus. Viruses 2011, 3, 2127–2145. [Google Scholar] [CrossRef]
- Lin, H.X.; Feng, Y.; Tu, X.; Zhao, X.; Hsieh, C.H.; Griffin, L.; Junop, M.; Zhang, C. Characterization of the spike protein of human coronavirus NL63 in receptor binding and pseudotype virus entry. Virus Res. 2011, 160, 283–293. [Google Scholar] [CrossRef]
- Shah, P.P.; Wang, T.; Kaletsky, R.L.; Myers, M.C.; Purvis, J.E.; Jing, H.; Huryn, D.M.; Greenbaum, D.C.; Smith, A.B., 3rd; Bates, P.; et al. A small-molecule oxocarbazate inhibitor of human cathepsin L blocks severe acute respiratory syndrome and ebola pseudotype virus infection into human embryonic kidney 293T cells. Mol. Pharmacol. 2010, 78, 319–324. [Google Scholar] [CrossRef]
- Kolokoltsov, A.A.; Wang, E.; Colpitts, T.M.; Weaver, S.C.; Davey, R.A. Pseudotyped viruses permit rapid detection of neutralizing antibodies in human and equine serum against Venezuelan equine encephalitis virus. Am. J. Trop. Med. Hyg. 2006, 75, 702–709. [Google Scholar]
- Basu, A.; Li, B.; Mills, D.M.; Panchal, R.G.; Cardinale, S.C.; Butler, M.M.; Peet, N.P.; Majgier-Baranowska, H.; Williams, J.D.; Patel, I.; et al. Identification of a small-molecule entry inhibitor for filoviruses. J. Virol. 2011, 85, 3106–3119. [Google Scholar]
- Dylla, D.E.; Xie, L.; Michele, D.E.; Kunz, S.; McCray, P.B., Jr. Altering alpha-dystroglycan receptor affinity of LCMV pseudotyped lentivirus yields unique cell and tissue tropism. Genet. Vaccines Ther. 2011, 9, 8. [Google Scholar] [CrossRef]
- Matsuno, K.; Kishida, N.; Usami, K.; Igarashi, M.; Yoshida, R.; Nakayama, E.; Shimojima, M.; Feldmann, H.; Irimura, T.; Kawaoka, Y.; Takada, A. Different potential of C-type lectin-mediated entry between Marburg virus strains. J. Virol. 2010, 84, 5140–5147. [Google Scholar]
- Radoshitzky, S.R.; Kuhn, J.H.; Spiropoulou, C.F.; Albarino, C.G.; Nguyen, D.P.; Salazar-Bravo, J.; Dorfman, T.; Lee, A.S.; Wang, E.; Ross, S.R.; et al. Receptor determinants of zoonotic transmission of New World hemorrhagic fever arenaviruses. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 2664–2669. [Google Scholar]
- Vela, E.M.; Zhang, L.; Colpitts, T.M.; Davey, R.A.; Aronson, J.F. Arenavirus entry occurs through a cholesterol-dependent, non-caveolar, clathrin-mediated endocytic mechanism. Virology 2007, 369, 1–11. [Google Scholar] [CrossRef]
- Ogino, M.; Ebihara, H.; Lee, B.H.; Araki, K.; Lundkvist, A.; Kawaoka, Y.; Yoshimatsu, K.; Arikawa, J. Use of vesicular stomatitis virus pseudotypes bearing hantaan or seoul virus envelope proteins in a rapid and safe neutralization test. Clin. Diagn. Lab. Immunol. 2003, 10, 154–160. [Google Scholar]
- Fukushi, S.; Mizutani, T.; Saijo, M.; Matsuyama, S.; Miyajima, N.; Taguchi, F.; Itamura, S.; Kurane, I.; Morikawa, S. Vesicular stomatitis virus pseudotyped with severe acute respiratory syndrome coronavirus spike protein. J. Gen. Virol. 2005, 86, 2269–2274. [Google Scholar] [CrossRef]
- Moore, M.J.; Dorfman, T.; Li, W.; Wong, S.K.; Li, Y.; Kuhn, J.H.; Coderre, J.; Vasilieva, N.; Han, Z.; Greenough, T.C.; et al. Retroviruses pseudotyped with the severe acute respiratory syndrome coronavirus spike protein efficiently infect cells expressing angiotensin-converting enzyme 2. J. Virol. 2004, 78, 10628–10635. [Google Scholar]
- Nie, Y.; Wang, P.; Shi, X.; Wang, G.; Chen, J.; Zheng, A.; Wang, W.; Wang, Z.; Qu, X.; Luo, M.; et al. Highly infectious SARS-CoV pseudotyped virus reveals the cell tropism and its correlation with receptor expression. Biochem. Biophys. Res. Commun. 2004, 321, 994–1000. [Google Scholar] [CrossRef]
- Fukushi, S.; Watanabe, R.; Taguchi, F. Pseudotyped vesicular stomatitis virus for analysis of virus entry mediated by SARS coronavirus spike proteins. Meth. Mol. Biol. 2008, 454, 331–338. [Google Scholar]
- Kaku, Y.; Noguchi, A.; Marsh, G.A.; Barr, J.A.; Okutani, A.; Hotta, K.; Bazartseren, B.; Fukushi, S.; Broder, C.C.; Yamada, A.; et al. Second generation of pseudotype-based serum neutralization assay for Nipah virus antibodies: Sensitive and high-throughput analysis utilizing secreted alkaline phosphatase. J. Virol. Meth. 2012, 179, 226–232. [Google Scholar] [CrossRef]
- Coulibaly-N'Golo, D.; Allali, B.; Kouassi, S.K.; Fichet-Calvet, E.; Becker-Ziaja, B.; Rieger, T.; Olschlager, S.; Dosso, H.; Denys, C.; Ter Meulen, J.; et al. Novel arenavirus sequences in Hylomyscus sp. and Mus (Nannomys) setulosus from Cote d'Ivoire: Implications for evolution of arenaviruses in Africa. PLoS One 2011, 6, e20893. [Google Scholar]
- Irwin, N.R.; Bayerlova, M.; Missa, O.; Martinkova, N. Complex patterns of host switching in New World arenaviruses. Mol. Ecol. 2012, 21, 4137–4150. [Google Scholar] [CrossRef]
© 2012 by the authors; licensee MDPI, Basel, Switzerland. This article is an open-access article distributed under the terms and conditions of the Creative Commons Attribution license (http://creativecommons.org/licenses/by/3.0/).