The rhabdoviruses: Biodiversity, phylogenetics, and evolution

The rhabdoviruses: Biodiversity, phylogenetics, and evolution

Infection, Genetics and Evolution 9 (2009) 541–553 Contents lists available at ScienceDirect Infection, Genetics and Evolution journal homepage: www...

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Infection, Genetics and Evolution 9 (2009) 541–553

Contents lists available at ScienceDirect

Infection, Genetics and Evolution journal homepage: www.elsevier.com/locate/meegid

The rhabdoviruses: Biodiversity, phylogenetics, and evolution§ I.V. Kuzmin a,*, I.S. Novella b, R.G. Dietzgen c, A. Padhi d, C.E. Rupprecht a a

National Center for Zoonotic, Vectorborne and Enteric Diseases, Centers for Disease Control and Prevention, 1600 Clifton Rd, Atlanta, GA 30333, USA Department of Medical Microbiology and Immunology, University of Toledo College of Medicine, 3055 Arlington Ave., Toledo, OH 43614, USA Queensland Department of Primary Industries and Fisheries, Emerging Technologies, PO Box 6097, St. Lucia, Qld 4067, Australia d Center for Infectious Disease Dynamics, 208 Mueller Laboratory, Department of Biology, The Pennsylvania State University, University Park, PA 16802, USA b c

A R T I C L E I N F O

A B S T R A C T

Article history: Received 14 October 2008 Received in revised form 9 February 2009 Accepted 13 February 2009 Available online 24 February 2009

Rhabdoviruses (family Rhabdoviridae) include a diversity of important pathogens of animals and plants. They share morphology and genome organization. The understanding of rhabdovirus phylogeny, ecology and evolution has progressed greatly during the last 30 years, due to enhanced surveillance and improved methodologies of molecular characterization. Along with six established genera, several phylogenetic groups at different levels were described within the Rhabdoviridae. However, comparative relationships between viral phylogeny and taxonomy remains incomplete, with multiple representatives awaiting further genetic characterization. The same is true for rhabdovirus evolution. To date, rather simplistic molecular clock models only partially describe the evolutionary dynamics of postulated viral lineages. Ongoing progress in viral evolutionary and ecological investigations will provide the platform for future studies of this diverse family. Published by Elsevier B.V.

Keywords: Rhabdovirus Phylogeny Evolution Lyssavirus Ephemerovirus Novirhabdovirus Vesiculovirus Nucleorhabdovirus Cytorhabdovirus

1. Introduction Rhabdoviruses (family Rhabdoviridae) are enveloped negativestrand RNA viruses that belong to the order Mononegavirales, which also includes the families Bornaviridae, Filoviridae and Paramyxoviridae. Rhabdoviruses are bullet or cone-shaped (from vertebrates and invertebrates) or bacilliform (from plants). These morphological characteristics have served as the basis for primary classification purposes compared to other families (Wagner, 1987). Further studies, performed for different rhabdoviruses, demonstrated their antigenic relatedness (Shope et al., 1970; Shope, 1982; Calisher et al., 1989). In addition, phylogenetic comparisons of available rhabdovirus genome fragments suggested a monophyletic origin (Bourhy et al., 2005; Tordo et al., 2005). The family

§ Disclaimer: Use of trade names and commercial sources are for identification only and do not imply endorsement by the U.S. Department of Health and Human Services. The findings and conclusions in this report are those of the authors and do not necessarily represent the views of the funding agency. * Corresponding author at: Centers for Disease Control and Prevention, 1600 Clifton Rd., Bldg 17, MS G-33, Atlanta, GA, USA. Tel.: +1 404 639 1050; fax: +1 404 639 1564. E-mail address: [email protected] (I.V. Kuzmin).

1567-1348/$ – see front matter . Published by Elsevier B.V. doi:10.1016/j.meegid.2009.02.005

includes pathogens of many animals and plants, including humans, livestock, fish and crops, with significant public health, veterinary and agricultural impact. Many are transmitted by arthropods, whereas lyssaviruses are transmitted directly between mammals by bite, and fish rhabdoviruses are waterborne. Some plant rhabdoviruses are also transmissible mechanically, by direct contact or ‘‘rub’’ inoculation. To date, 6 genera (Vesiculovirus, Lyssavirus, Ephemerovirus, Novirhabdovirus, Cytorhabdovirus and Nucleorhabdovirus) and more than 130 unassigned viruses are recognized within the Rhabdoviridae (Tordo et al., 2005; Jackson et al., 2005). In addition, the Vesiculovirus and the Ephemerovirus genera, together with currently non-classified viruses from the tentatively named Hart Park group, Le Dantec group, Almpiwar group and Tibrogargan group, were considered as members of phylogenetic ‘supergroup’ Dimarhabdovirus (‘dipteran-mammal associated rhabdovirus’) (Bourhy et al., 2005). Other non-classified members of this ‘supergroup’ have been recognized, based on phylogenetic relationships as well, such as Oita virus (OITAV), Mount Elgon bat virus (MEBV), Kern canyon virus (KCV), Kolongo virus (KOLV), Sanjimba virus (SJAV), Rochambeau virus (RBUV), and Tupaia virus (TUPV) (Iwasaki et al., 2004; Springfeld et al., 2005; Kuzmin et al., 2006). The terminology for this ‘supergroup’ should be

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re-evaluated, as it includes viruses isolated from birds, such as KOLV and SJAV, and fish vesiculoviruses. Moreover, Almpiwar virus (ALMV) and Charleville virus (CHVV) were isolated from reptiles. However, for consistency, in the present paper we will call them ‘‘dimarhabdoviruses’’. Historical antigenic comparisons suggested number of other viruses within these groups, as well as the existence of other antigenically related groups of rhabdoviruses, such as Bahia Grande group, Sawgrass group, and Timbo group (Tordo et al., 2005). However, as phylogenetic information on these viruses is lacking, we omit them in the present review. Sigma virus (SIGMAV), which does not show significant phylogenetic relatedness to other rhabdoviruses and has unique specific association with Drosophila melanogaster, is not covered by our review as well. Rhabdovirus virions are 100–430 nm long and 45–100 nm in diameter and composed of two structural units: an internal helical ribonucleoprotein complex (RNP), and a lipid envelope which is derived from the host cell membranes during budding. The RNP is comprised of the tightly associated RNA genome and nucleoprotein (N). The heavily phosphorylated phosphoprotein (P) and RNAdependent RNA-polymerase (L) are also bound to the RNP. The exact position of the matrix protein (M) remains controversial, and may be either contained in the central channel of RNP or embedded into the inner layer of the virion membrane. Knobbed glycoprotein (G) spikes, serving for binding of the virions to host cell receptors, protrude through the virion membrane. Rhabdoviruses contain a single molecule of linear, negativesense ssRNA. The RNA has a 30 -terminal free OH group and a 50 triphosphate. The termini have inverted complementary sequences. All rhabdoviruses described to date ultimately have five structural genes in the order 30 -N-P-M-G-L-50 . The corresponding cistrons are flanked by conserved start and stop transcription signals, and for certain viruses, additional genes are interposed. For example, vesiculoviruses encompass three ORFs in the P gene that code for P, C0 and C proteins (Spiropoulou and Nichol, 1993). The vesiculovirus M gene (at least in the vesicular stomatitis Indiana virus) may also encode two additional products, M2 and M3, that are generated from translation starting at either of two alternative downstream ATGs in the same frame as the M protein and result in proteins that lack the first 32 or 50 amino acids (Jayakar and Whitt, 2002). A more complex genome organization, including a second non-structural G gene (Gns), as well as a1/a2/a3, b and g ORFs between the G and L genes was reported in the ephemeroviruses (Wang et al., 1994; McWilliam et al., 1997). The novirhabdoviruses have a NV gene between the G and L genes (Morzunov et al., 1995). An additional transcription unit, SH, was detected between the M and G genes of the TUPV (Springfeld et al., 2005). Putative cell-to-cell movement protein genes are located between the P and M genes of the cytorhabdoviruses and nucleorhabdoviruses, and genes of unknown function between the G and L genes of some plant rhabdoviruses (Jackson et al., 2005). An additional gene was also detected between the P and M genes of the SIGMAV genome (Lande`s-Devauchelle et al., 1995). Even more complex genome organization was reported recently for Wongabel virus (WONV), where three additional genes (U1, U2 and U3) were detected between the P and M genes, and two ORFs (U4 and U5) overlapping with the N and G genes (Gubala et al., 2008). The significance of the non-structural genes is largely unknown. The N protein encapsidates the entire RNA and tightly packages it into an RNase-resistant core, which serves as a template for both replication and transcription. The comparisons of the N protein from different rhabdoviruses demonstrated that the protein fold is highly conserved, despite limited identity in amino acid sequence. The RNA binding cavity is the highest conserved feature in the N protein, even if different amino acids are used for encapsidation (Luo et al., 2007). The N protein, prior to association with viral

genome, is associated with the P protein, which acts as a chaperone to maintain the N protein in a soluble form (Jayakar et al., 2004). The L protein performs two activities (transcription and replication), which are probably performed by different polymerase complexes, consisting of either L-(P)3, which is the transcriptase, or N-P-L which is the replicase (Gupta et al., 2003). The L protein is tightly associated with the RNP core. Assembly of the viral polymerase into the virion is essential since negative-sense RNA viral genomes cannot be used as mRNA, and host cells do not have appropriate enzymes to catalyze transcription. The polymerase protein of rhabdoviruses and other mononegavirales is organized into six well-conserved blocks (Poch et al., 1990). The G protein of rhabdoviruses is a type I glycoprotein, with a large ectodomain, a transmembrane domain, and a cytoplasmic domain. Ectodomain trimers protrude from the surface of the viral envelope. This trimer is considered the functional unit for both assembly into virions and virus entry into vertebrate and invertebrate host cells (Jayakar et al., 2004). The M protein plays multiple roles in rhabdovirus assembly, including condensation of RNP cores and formation of cone- or bullet-shaped virion particles, as well as participating in the budding of virions from cell membranes (Jayakar et al., 2004). The M protein also mediates inhibition of gene expression in host cells (Kopecky and Lyles, 2003). Except for plant rhabdoviruses that generally penetrate the cell through mechanical damage provoked by insect vectors, rhabdovirus adsorption is mediated by the G protein attachment to cell surface receptors. Penetration of the cell is by endocytosis. After endocytosis, the pH decreases within the endosome, leading to fusion between the endosomal and viral membranes. This liberates the RNP into the cytoplasm. Once the RNP is liberated, the genome RNA is repetitively transcribed by the virion transcriptase (N protein removal is not required, and the RNP serves as template for transcription and replication). The translation is ensured by cellular machinery. In general, all processes of viral transcription, translation and replication take place in the cytoplasm. The glycoprotein is delivered to the cytoplasmic membrane, whereas other viral proteins are expressed in the cytosol by free polyribosomes. At the final stage, viral transcription and replication are inhibited, the RNP becomes intensively condensed and subsequently delivered to the cell membrane, and virions are ready for budding. During budding, the virions acquire the glycoprotein and lipid envelope provided by the cell membrane for selfassembly. Nucleorhabdoviruses replicate in large inclusions or ‘‘viroplasms’’ in the cell nucleus (Jackson et al., 2005). Morphogenesis of such viruses occurs at the inner nuclear envelope, and enveloped virus particles accumulate in perinuclear spaces. The objective of this communication is to discuss recent insights into the biodiversity, molecular phylogeny, and proposed evolution of Rhabdoviridae. 2. Vesiculoviruses Members of the vesicular stomatitis virus (VSV) group are dimarhabdoviruses that include two serotypes of VSV, the Indiana serotype and the New Jersey serotype (Vesicular stomatitis New Jersey virus, VSNJV), as well as Piry virus (PIRYV), Isfahan virus (ISFV), Carajas virus (CJSV), Maraba virus (MARAV) and Chandipura virus (CHPV). Within the Indiana serotype, there are three subtypes: Vesicular stomatitis Indiana virus (‘‘classical’’ Indiana 1; VSIV), Cocal virus (Indiana 2; COCV) and Alagoas virus (Indiana 3; VSAV). These viruses were originally classified based on serology, but all available analyses based on gene sequences supports the initial grouping. The phylogeny of these viruses show two distinct clades, one with VSIV, VSNJV, VSAV and COCV, and another that includes PIRYV, ISFV and CHPV. Sequence information for CJSV and

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MARAV is limited, and therefore their phylogeny is unclear. All vesiculoviruses are arthropod-borne, except PIRYV, which to this date has never been isolated from insects. Insect infection can result in transovarial transmission, at least in VSIV and VSNJV (Tesh et al., 1972, 1987). The members of this group have different geographical distributions, host preferences and various associations with disease. VSIV and VSNJV are endemic between southern North America and northern South America, and they cause annual outbreaks that usually correlate with the change from the rainy to dry seasons (Rodriguez, 2002). Both viruses can be isolated from many insect species, but sand flies (Lutzomiya spp.) and black flies (Simulium spp.) are considered the predominant vectors. These viruses infect livestock, including cows, pigs and horses, as well as a variety of wild mammals. Vesicular stomatitis of livestock has considerable economic impact. Human infections are possible, but are usually restricted to individuals working with sick livestock, or laboratory personnel who do not follow proper handling procedures. COCV and VSAV are South American viruses that can be isolated from sand flies and cause sporadic outbreaks of vesicular stomatitis in cattle (Pauszek et al., 2008). COCV was reported as cause of the disease in Brazil and Argentina. VSAV causes epizootics in Brazil, but it has also been isolated from sand flies and seropositive (but otherwise healthy) livestock in Colombia (Tesh et al., 1987). PIRYV is the third member of the group found in South America, specifically in Brazil, and caused a febrile disease in humans (Marriott, 2005). The distribution of CHPV includes several areas of the Indian subcontinent. In addition, it has been identified in Senegal (Marriott, 2005). CHPV is pathogenic to humans, and infection can have different outcomes, ranging from subclinical cases to fatal encephalitis (Arankalle et al., 2005). The virus has been isolated from sand flies and there is serological evidence of infection of domestic animals (Basak et al., 2007). Also from serological analyses, the presence of ISFV has been detected in India, Iran, Turkmenistan and other Asian countries (Marriott, 2005). Several fish rhabdoviruses demonstrate phylogenetic relatedness to vesiculoviruses, have the same genome organization (Hoffmann et al., 2005), and are included into the genus as tentative species. These include spring viremia carp virus (SVCV), pike fry rhabdovirus (PFRV), Eel virus American (EVA) and ulcerative disease rhabdovirus (UDRV) (Tordo et al., 2005). Several other fish viruses have been tentatively assigned into the vesiculovirus genus, but their exact relationships are somewhat inconclusive, as gene sequences for most of them are unavailable to date. For example, Eel virus Europe X (EVEX) and EVA are placed in this group, based on serological evidence only (Hoffmann et al., 2005). Trout rhabdovirus (TRV 903/87) and sea trout rhabdovirus (STRV 28/97) are closely related to each other, and show high gene sequence identity to vesiculoviruses, even if placed on the phylogenetic tree ancestrally to other genus representatives (Fig. 1). However, based on the L and M gene sequences, phylogenetic position of these viruses is uncertain, suggesting their relatedness to other rhabdoviruses, such as novirhabdoviruses and ephemeroviruses (Johansson et al., 2001; Hoffmann et al., 2005). Partial sequencing of the L gene of Scophthalmus maximus rhabdovirus (SMRV) shows relatively high identity with sequences of vesiculoviruses, and this relationship is supported by serological cross-reactivity (Zhang et al., 2007). Similarly, relatedness to vesiculoviruses was demonstrated for Siniperca chuatsi rhabdovirus (SCRV), the complete genome sequence of which was recovered recently (Tao et al., 2008). Lastly, Perinet virus (PERV), isolated from mosquitoes in Madagascar (Clerc et al., 1982), demonstrated significant relatedness to vesiculoviruses based on the available limited L gene sequence (Bourhy et al., 2005; Fig. 1B).

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Nichol (1988) have performed extensive phylogenetic analysis of VSNJV and VSIV natural isolates. The degree of diversity in enzootic zones (e.g. Central America) is much more substantial than the degree of diversity in epizootic zones (Northern Mexico and Southern US), demonstrating co-circulation of multiple strains within the former area of distribution. Furthermore, variation followed geographical, not temporal clines. Strains isolated from distant areas during the same year diverged more than strains isolated from the same area years apart (Nichol, 1988; Bisel et al., 1990; Bisel and Nichol, 1990; Rodriguez et al., 2000). These observations indicate co-circulation over extensive periods of time. Additional analyses showed a stepwise distribution of isolates that correlated with a south-to-north gradient. The putative origin of vesiculoviruses is placed in southern Central America (Nichol et al., 1993). Subsequently, Rodriguez et al. (1996) found that endemic VSNJV from Central America grouped within ecological zones. Of the two clades identified, clade A was found in lowlands of Costa Rica and Panama, while clade B was found in the highlands of Costa Rica and Nicaragua. Divergence between the two clades may be driven by replication in different vectors: sand flies in lowlands and black flies in the highlands. Extensive spatio-temporal studies of VSNJV in epizootic areas indicated that both periodic reintroductions and endemic cycles contribute to the development of outbreaks (Rainwater-Lovett et al., 2007). Isolation of closely related VSIV strains from livestock over several years suggested the possibility of viral persistence in epizootic areas (Rodriguez et al., 2000). 3. Ephemeroviruses The genus Ephemerovirus currently contains 3 species, including Adelaide river virus (ARV), Bovine ephemeral fever virus (BEFV) and Berrimah virus (BRMV), and three tentative species: Kimberley virus (KIMV), Malakal virus (MALV) and Puchong virus (PUCV) (Tordo et al., 2005). Phylogenetic placement of KIMV into the genus was confirmed by gene sequencing (Bourhy et al., 2005), whereas MALV and PUCV were added to the genus based on serology only. BEFV is the most broadly distributed member of the genus, isolated in the tropics and subtropics of Australia, Africa, Asia and the Middle East. Strains isolated from these distant territories are closely related to each other antigenically and phylogenetically (Walker, 2005). Besides cattle, water buffalo are susceptible to BEFV, but it is unclear if subclinical infection of this species is possible (St.George et al., 1977). The role of wildlife as a reservoir is uncertain at present. In Kenya, antibodies to BEFV were detected in African buffalo, water buck, wildebeest and hartebeest (Davies and Walker, 1974). Red deer in Australia demonstrated a significant seroprevalence to BEFV (St.George, 1988). In contrast, ARV has been isolated to date only in Australia (Gard et al., 1984) but serologic findings suggested that ARV or closely related viruses may circulate in China and Indonesia (Bai et al., 1993; Daniels et al., 1995). BRMV was isolated only once from an apparently healthy steer in Australia in 1981 (Gard et al., 1983), while KIMV was repeatedly isolated from mosquitoes and apparently healthy cattle in Australia. MALV and PUCV have been isolated from mosquitoes in Sudan and Malaysia, respectively (Walker, 2005). In addition, the unclassified Kotonkan virus (KOTV) and Obodhiang virus (OBOV), each known by a single isolate from dipterans in Nigeria and Sudan, respectively, were shown to belong to ephemeroviruses based on phylogeny (Kuzmin et al., 2006). Members of different ephemerovirus species exhibit low to no cross-neutralization, but they cross-react significantly in the complement fixation and immunofluorescence tests. In addition, very low levels of antigenic cross-reactivity were demonstrated between ephemeroviruses and certain lyssaviruses. For that reason, KOTV, OBOV and RBUV were initially mistakenly suggested as lyssavirus

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representatives (Shope, 1982; Calisher et al., 1989; Tordo et al., 2005), which has not been corroborated by sequence data (Kuzmin et al., 2006). Ephemeroviruses are distributed widely in the Old World tropics and subtropics. Dipterans are the only insects from which ephemeroviruses have been isolated. Windborne spread across the tropics has been suggested for viruses of this group, and cattle translocation between continents has been suggested to explain their broad distribution. The apparent emergence and re-emergence of the disease over 125 years is likely due to the expansive growth of the cattle industry and improved surveillance (Walker, 2005). Genetic similarity between the Australian ephemeroviruses ARV and BEFV is less than the similarity between ARV and African OBOV. The presence in this cluster of the more distantly related African viruses KOTV and RBUV suggests that intercontinental translocations of ephemeroviruses have occurred several times. 4. Other dimarhabdoviruses Unclassified dimarhabdoviruses include the Hart Park group, [Parry Creek virus (PCRV), WONV, and Ngaingan virus (NGAV), all

isolated in Australia, and Flanders virus (FLAV) isolated in North America]; the Almpiwar group [ALMV, CHVV, and Humpty Doo virus (HDOOV), all isolated in Australia]; the Tibrogargan group, represented by a single available sequence of Tibrogargan virus (TIBV) from Australia; and the Le Dantec group [Le Dantec virus (LDV) from Senegal and Fukuoka virus (FUKV) from Japan]. These groups have been confirmed by Bourhy et al. (2005) via comparisons of limited L gene sequences, but more number was proposed based on antigenic cross-reactivity (Calisher et al., 1989; Tordo et al., 2005). The Oak-Vale virus (OVRV) did not demonstrate significant phylogenetic relatedness to any dimarhabdovirus group based on limited L gene sequence (Fig. 1B). Another phylogenetic analysis of N gene sequences (Fig. 1A) demonstrated that Kern Canyon virus (KCV) is monophyletic with Oita virus (OITAV) and Mont Elgon bat virus (MEBV) (Kuzmin et al., 2006). Since Calisher et al. (1989) demonstrated that KCV is antigenically related to FUKV, there is a probability that FUKV and LDV are also members of this phylogenetic group, but so far their sequence data represent different genome regions, and cannot be compared directly. Two viruses isolated in central Africa, Kolongo (KOLV) and Sanjimba (SJAV), together with TUPV, constitute another mono-

Fig. 1. Phylogenetic tree of Rhabdoviridae based on available partial sequences. A: nucleoprotein gene (alignment of 1027 nucleotides); B: polymerase gene (alignment of 460 nucleotides). For virus abbreviations see Table 1. The trees were generated by the neighbor-joining method, for 1000 bootstrap replicates. Significant bootstrap support (over 70) is shown for key nodes.

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Fig. 1. (Continued ).

phyletic clade within the dimarhabdoviruses (Kuzmin et al., 2006). Both KOLV and SJAV were isolated from birds, so from the standpoint of mobility there is no reason to suggest that their distribution is restricted to the central Africa only. Furthermore, TUPV was isolated from a tree shrew (Tupaia belangeri), imported from Thailand and has been shown to replicate only in tree shrew cells (Springfeld et al., 2005). These mammals are indigenous to tropical Asia, but not to Africa. Taken together, these data suggest that representatives of this group could be quite numerous and circulate intercontinentally among various hosts. 5. Lyssaviruses Members of the genus Lyssavirus constitute a single monophyletic clade, distinct from all other rhabdoviruses. Lyssavirus taxonomy is complex due to a combination of historical precedents based on serological methods, current knowledge based on phylogenetic analyses, and unique pathobiology of these viruses. The genus includes seven species that correspond to seven genotypes (Bourhy et al., 1993; Tordo et al., 1993, 2005). Rabies virus (RABV; genotype 1) is distributed most broadly in carnivores in the Old World and in both carnivores and bats in the New World. This lyssavirus is responsible for most human rabies cases, and is the best studied. Lagos bat virus

(LBV; genotype 2) was identified in pteropid bats across Africa. Occasionally LBV has been isolated from cats, dogs and a mongoose. Further investigations may facilitate subdivision of LBV into two or more independent genotypes (Markotter et al., 2008). Mokola virus (MOKV; genotype 3) was first isolated from shrews in Nigeria (Shope et al., 1970). Thereafter, MOKV was detected in shrews from Nigeria and Cameroon; humans in Nigeria; cats in Zimbabwe, Ethiopia and South Africa; a dog in Zimbabwe; and a rodent from the Central Africa Republic (Nel et al., 2000). Duvenhage virus (DUVV; genotype 4) was isolated from insectivorous bats and humans who died after bat bites in South Africa, Zimbabwe and Kenya (Meredith et al., 1971; King and Crick, 1988; Van Thiel et al., 2008). European bat lyssavirus, type 1 (EBLV-1; genotype 5) circulates in Europe among insectivorous bats (predominantly Eptesicus serotinus), and European bat lyssavirus, type 2 (EBLV-2; genotype 6) circulates in insectivorous bats (predominantly of Myotis genus) in northern and western Europe (Amengual et al., 1997; Davis et al., 2005). Australian bat lyssavirus (ABLV; genotype 7) was described in Australia, where two separate lineages circulate in pteropid and insectivorous bats (Fraser et al., 1996; Gould et al., 1998, 2002; Guyatt et al., 2003). Four new lyssaviruses were described from Eurasia: Aravan virus (ARAV), Khujand virus (KHUV), Irkut virus (IRKV) and West

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Table 1 Alphabetic list of viruses referred in the present article. Virus name

Abbreviation

Genus

Host range (source); vector

Continent of origin/distribution

Adelaide river virus Alagoas virus Almpiwar virus Aravan virus Australian bat lyssavirus Berrimah virus Bovine ephemeral fever virus Carajas virus Chandipura virus Charleville virus Cocal virus Duvenhage virus Eel virus American Eel virus B12 Eel virus C26 Eel virus Europe X European bat lyssavirus, type 1 European bat lyssavirus, type 2 Flanders virus Fukuoka virus Hirame rhabdovirus Humpty Doo virus Infectious hematopoietic necrosis virus Irkut virus Isfahan virus Kern canyon virus Khujand virus Kimberley virus Kolongo virus Kotonkan virus Lagos bat virus Le Dantec virus Lettuce necrotic yellows virus Lettuce yellow mottle virus Maize fine streak virus Malakal virus Maraba virus Mokola virus Mount Elgon bat virus Ngaingan virus Northern cereal mosaic virus Oak-Vale virus Obodhiang virus Oita virus Parry Creek virus Perinet virus Pike fry rhabdovirus Piry virus Puchong virus Rabies virus Rice yellow stunt virus Rochambeau virus Sanjimba virus Scophthalmus maximus rhabdovirus Sea trout rhabdovirus Sigma virus Siniperca chuatsi rhabdovirus Snakehead rhabdovirus Sonchus yellow net virus Spring viremia of carp virus Strawberry crinkle virus Taro vein chlorosis virus Tibrogargan virus Tupaia virus Ulcerative disease rhabdovirus Vesicular stomatitis Indiana virus Vesicular stomatitis New Jersey virus Viral hemorrhagic septicemia virus West Caucasian bat virus Wongabel virus

ARV VSAV ALMV ARAV ABLV BRMV BEFV CJSV CHPV CHVV COCV DUVV EVA EEV-B12 EEV-C26 EVEX EBLV-1 EBLV-2 FLAV FUKV HIRRV HDOOV IHNV IRKV ISFV KCV KHUV KIMV KOLV KOTV LBV LDV LNYV LYMoV MFSV MALV MARAV MOKV MEBV NGAV NCMV OVRV OBOV OITAV PCRV PERV PFRV PIRYV PUCV RABV RYSV RBUV SJAV SMRV STRV SIGMAV SCRV SHRV SYNV SVCV SCV TaVCV TIBV TUPV UDRV VSIV VSNJV VHSV WCBV WONV

Ephemerovirus Vesiculovirus Unassigned Lyssavirus Lyssavirus Ephemerovirus Ephemerovirus Vesiculovirus Vesiculorirus Unassigned Vesiculovirus Lyssavirus Vesiculovirusa Novirhabdovirus Novirhabdovirus Vesiculovirusa Lyssavirus Lyssavirus Unassigned Unassigned Novirhabdovirus Unassigned Novirhabdovirus Lyssavirus Vesiculovirus Unassigned Lyssavirus Ephemerovirus Unassigned Unassignedb Lyssavirus Unassigned Cytorhabdovirus Unassignedc Unassignedd Ephemerovirus Vesiculovirus Lyssavirus Unassigned Unassigned Cytorhabdovirus Unassigned Unassignedb Unassigned Unassigned Unassignede Vesiculovirus Vesiculovirus Ephemerovirus Lyssavirus Nucleorhabdovirus Unassignedf Unassigned Unassignedg Unassigned Unassigned Unassignedh Novirhabdovirus Nucleorhabdovirus Vesiculovirus Cytorhabdovirus Unassignedi Unassigned Unassigned Vesiculovirus Vesiculovirus Vesiculovirus Novirhabdovirus Lyssavirus Unassigned

Livestock; dipterans Livestock; sand flies Lizard Ablepharus boutonii virgatus Bat Myotis blythii Pteropid and insectivorous bats Livestock Livestock; dipterans Livestock; sand flies Livestock; sand flies Mosquitoes; lizard Gehyra australis Livestock; sand flies Insectivorous bats Fish Fish Fish Fish Insectivorous bats Insectivorous bats Mosquitoes; birds Mosquitoes Fish Gnat Lasiohelea spp Fish Bat Murina leucogaster Sand flies Bat Myotis yumanensis Bat Myotis mystacinus Livestock; dipterans Bird Euplecies afra Midges Pteropid bats Human Compositae; aphids Lettuce; vector unknown Graminae; leafhoppers Mosquitoes Sand flies Shrews, cats, a dog Bat Rhinolophus hildebrandtii Midges Graminae; planthoppers Mosquitoes Mosquitoes Bat Rhinolophus cornutus Mosquitoes Mosquitoes Fish Sand flies Mosquitoes Carnivores and bats Rice; leafhoppers Mosquitoes Bird Acrocephalus schoenobaenus Fish Fish Drosophila melanogaster Fish Fish Compositae; aphids Fish Strawberry; aphids Taro; vector unknown Midges Tree shrew Fish Livestock; sand flies and other dipterans Livestock; sand flies and other dipterans Fish Bat Miniopterus schreibersii Midges

Australia South America Australia Asia Australia Australia Tropics and subtropics of Old World South America Asia, Africa Australia South America Africa North America, Japan Europe Europe Worldwide Europe Europe North America Japan Japan Australia Worldwide Asia Asia North America Asia Australia Africa Africa Africa Africa Australia, New Zealand Europe North America Africa South America Africa Africa Australia Asia Australia Africa Japan Australia Madagascar Europe South America Asia Worldwide Asia Africa Africa Asia Europe Worldwide Asia Asia North America Worldwide Worldwide Pacific Islands Australia Asia Asia North and South America North and South America Worldwide Europe Australia

a b c d e

Tentative species, based on serological relationship only (Hoffmann et al., 2005). Ephemerovirus according to recent phylogenetic comparisons (Bourhy et al., 2005; Kuzmin et al., 2006). Cytorhabdovirus according to recent phylogenetic comparisons (Heim et al., 2008). Nucleorhabdovirus according to recent phylogenetic comparisons (Redinbaugh et al., 2002). Vesiculovirus according to recent phylogenetic comparisons (Bourhy et al., 2005).

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Caucasian bat virus (WCBV) (Kuzmin et al., 2003, 2005). Only one isolate of each of these viruses is available to date. They are included into the ICTV list as tentative lyssavirus species. One other tentative species, Rochambeau virus (RBUV), is currently listed within the genus, but was shown recently to have no significant phylogenetic relatedness to lyssaviruses (Kuzmin et al., 2006). In general, the criteria for demarcation of lyssavirus species are challenging. Many characteristics, including genetic, antigenic and ecological properties should be considered. Genetic distances between established lyssavirus species are shorter than between species within other rhabdovirus genera. Placement of lyssaviruses in comparative genetic scale with other rhabdoviruses may support a taxonomy that all belong to a single species, potentially named ‘‘rabies virus’’. All lyssaviruses have similar pathobiology and cause acute progressive encephalitis (rabies) in mammals and, in contrast to other rhabdoviruses, demonstrate significant serologic cross-reactivity. As one simplified approach, the operational definition of genotype is broadly used for lyssavirus classification. The threshold of 80–82% identity for nucleotide sequences of the entire nucleoprotein genes was suggested for separation of lyssavirus genotypes (Kissi et al., 1995; Kuzmin et al., 2005). Complete genome sequences, which have been published in an increasing number during most recent years, may provide better tools for phylogenetic classification (Delmas et al., 2008). Early taxonomy of the Rhabdoviridae was based on virion morphology and antigenic cross-reactivity. Indeed, many as yet unclassified rhabdoviruses exhibited cross-reactivity with lyssaviruses (Calisher et al., 1989). When the terms ‘‘rabies-related viruses’’ and the ‘‘rabies serogroup’’ were introduced (Shope et al., 1970), these included RABV, MOKV and LBV. Shortly thereafter, new members of the ‘‘serogroup’’ were described, such as DUVV (including at that time EBLV-1), KOTV and OBOV (Meredith et al., 1971; Kemp et al., 1973; Tignor et al., 1977; Shope, 1982). The latter two viruses, isolated from dipterans, demonstrated a limited cross-reactivity with MOKV, but not with other members of the ‘‘serogroup’’. Additionally, MOKV can replicate in insect cell lines (Buckley, 1975), although no insectderived isolates have been obtained from nature. As was speculated, the proposed evolutionary pathway for the rabiesrelated viruses included OBOV and KOTV as progenitors, through MOKV, to other highly neurotropic mammalian viruses, such as LBV, DUVV and RABV (Shope, 1982). Africa was suggested to be the continent of primary lyssavirus origination. Later, other rhabdoviruses, isolated from both arthropods and vertebrates predominantly in Africa, were described as serologically related to those listed above (Calisher et al., 1989). However, when genes of OBOV, KOTV and some other rhabdoviruses were sequenced, they were not phylogenetically related to lyssaviruses (Kuzmin et al., 2006). As a global concern, two phylogroups were delineated within the genus, based on phylogenetic relatedness, serologic crossreactivity and peripheral pathogenicity in mice (Badrane et al., 2001). Phylogroup 1 includes RABV, DUVV, EBLV-1, EBLV-2, ABLV, ARAV, KHUV and IRKV. Phylogroup 2 includes LBV and MOKV. As was additionally demonstrated, WCBV is not a member of either phylogroups 1 or 2 because of its ancestral phylogenetic position, equally low identity to both phylogroups, and absence of crossneutralization with all other lyssaviruses (Hanlon et al., 2005; Kuzmin et al., 2005).

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Within the phylogroup 1, DUVV, EBLVs, ARAV, KHUV and IRKV are monophyletic, and represent a cluster of Old World bat lyssaviruses (of African and Eurasian origin). Their relatedness is evident for each gene compared (Kuzmin et al., 2005), although it is not seen in the tree based on limited L gene sequences (Fig. 1B). As was proposed, DUVV (or a progenitor virus) had been introduced from Africa to Europe, where it evolved into EBLV (Shope, 1982; Schneider and Cox, 1994; Amengual et al., 1997; Serra-Cobo et al., 2002). However, to date there are no significant evidences to support this hypothesis. Other viruses of this group (EBLVs, ARAV, KHUV and IRKV) have been identified in Eurasia. Moreover, the most divergent member of the genus, WCBV, was isolated in Eurasia as well. The principal host of WCBV is unknown. It was isolated once from Miniopterus schreibersi, a broadly distributed colonial insectivorous bat. Several species of Miniopterus bats are common in the tropics and subtropics across the Old World. Recently, significant levels of seroprevalence to WCBV were detected in Miniopterus bats from Kenya (Kuzmin et al., 2008c). It appears that the Lyssavirus genus evolved in bats. The only species that had not been identified in bats is MOKV. However, the ecology of MOKV has not been sufficiently studied and it is not possible to determine a principal host range or the circulation properties of this virus at present. The evolutionary history within the genus is unclear. For example, all non-rabies lyssaviruses described to date are found in the Old World only. RABV isolation from Old World bats was never confirmed (Kuzmin et al., 2006). In contrast, in the New World, only RABV is presented in both carnivores and bats. According to the hypothesis of Badrane and Tordo (2001), inferred from phylogenetic reconstructions, there were at least two major switches in lyssavirus history, both from bats to the terrestrial mammals. The first switch occurred on the inter-species level, and the second one occurred within RABV, when it switched from bats to carnivores. However, a major question remains, in which mammal group and where did these switches occur? Why are nonRABV lyssaviruses (except MOKV, which cannot be considered as the direct predecessor of RABV) not found in terrestrial reservoirs, and why is RABV not found in the Old World bats? In the phylogenetic tree of RABV representatives, the North American RABV lineages, circulating among raccoons and skunks, appear to be more related to bat RABVs rather than to other terrestrial lineages of RABV, which are represented in the New World by the ‘‘cosmopolitan’’ RABV variant, believed to have originated in Europe, and widely disseminated as a consequence of colonial activity during 16th to 19th centuries (Smith and Seidel, 1993; Badrane and Tordo, 2001), and an Arctic RABV, distributed circumpolarly. Thus, if we consider the phylogenetic tree from geographical and historical perspectives, rather than from the viewpoint of host adaptation, an alternative hypothesis would be that the raccoon and skunk RABVs, together with bat RABVs, appear to be indigenous New World lineages that could have one common ancestor. This scenario does not require that the raccoon and skunk RABVs originated directly from bat RABVs, as the result of one host switch. Although, cases of host switches of bat RABVs to terrestrial mammals, with further circulation in their populations, are well documented (Leslie et al., 2006). 6. Novirhabdoviruses Members of the genus Novirhabdovirus include four fish pathogenic viruses: Infectious hematopoietic necrosis virus (IHNV),

f Currently listed as tentative species of the Lyssavirus genus (Tordo et al., 2005), but recent analysis demonstrated no phylogenetic relatedness to lyssaviruses (Kuzmin et al., 2006). g Vesiculovirus according to recent phylogenetic comparisons (Zhang et al., 2007). h Vesiculovirus according to recent phylogenetic comparisons (Tao et al., 2008). i Nucleorhabdovirus according to recent phylogenetic comparisons (Revill et al., 2005).

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Viral hemorrhagic septicemia virus (VHSV), Hirame rhabdovirus (HIRRV), Snakehead rhabdovirus (SHRV); and two tentative species, Eel virus B12 (EEV-B12), and Eel virus C26 (EEV-C26) (Tordo et al., 2005). These viruses have been isolated from several freshwater and marine fish species endemic to North America, Europe, and Asia (Skall et al., 2005; Hoffmann et al., 2005). Both IHNV and VHSV cause severe economic losses to the salmonid farming industries in North America and Europe (Wolf, 1988). IHNV was first discovered in the Pacific Northwest of the United States in 1953 (Rucker et al., 1953), and has subsequently spread into Japan (Kimura and Awakura, 1977; Sano et al., 1977; Yamazaki and Motonishi, 1992), South Korea (Park et al., 1993) and many European countries (Baudin-Laurencin, 1987; Bovo et al., 1987; Enzmann et al., 1992, 2005; Fichtner et al., 2000; Vardic´ et al., 2007; Rudakova et al., 2007). IHNV is endemic to Western North America, and dispersal of the virus outside North America has occurred by inadvertent transport of infected eggs and juvenile fish (Kimura and Awakura, 1977; Sano et al., 1977; Hill, 1992). Within North America, dispersal of IHNV is thought to have involved the historical use of unpasteurized salmon viscera in feed for salmon hatcheries, and possibly the historically common practice of salmon transplantations (Wolf, 1988; Roppel, 1982; Burgner, 1991; Watson et al., 1954). VHSV was first discovered in Western Europe (Wolf, 1988). So far, VHSV has been isolated from over 60 fish species from both marine and freshwater habitats representing North America, Asia, and Europe (Skall et al., 2005). VHSV is endemic to numerous marine species in both the Atlantic and Pacific Oceans of the northern hemisphere, and could have been introduced into freshwater habitats by marine fish species (e.g. herring, sprat, sand eel) that are used as fresh feed for commercial farming in some countries (Skall et al., 2005). HIRRV was first isolated in Japan from Japanese flounder (Kimura et al., 1986) and subsequently was also reported in Korea (Oh and Choi, 1998; Kim et al., 2005). It has recently been reported to infect several other fish species endemic to Japan (Kimura et al., 1986). SHRV was first isolated from snakehead fish in Thailand, and has not been reported outside Southeast Asia to date (Kasornchandra et al., 1992). The G gene showed relatively high genetic diversity compared to other genes, and therefore has been widely used to infer intraspecific phylogeography of respective viral species. Nichol et al. (1995) first reported the epizootiology of the IHNV in North America based on the full length G- and NV-genes. These authors reported that IHNV revealed a strong spatial structure, but with a high degree of genetic homogeneity. In a broad survey, based on partial G-gene sequence data, Kurath et al. (2003) have also reported similar observations. Despite the low genetic diversity, Kurath et al. (2003) reported the existence of three distinct genogroups of IHNV designated as upper (U), middle (M), and lower (L). The M genogroup, which comprise isolates from an Idaho trout-farming region, showed three-to-four fold greater diversity than other genogroups (Kurath et al., 2003; Troyer and Kurath, 2003), indicating a rapid evolution of this group. In contrast, genogroup U, which comprises isolates from Alaska, British Columbia, the Washington coast, and the Columbia River basin origin, has a slower rate of evolution (Kurath et al., 2003). Although the reason for such a discrepancy is unclear, Kurath et al. (2003) postulated that the ocean migration ranges of host populations, as well as anthropogenic effects associated with aquaculture, could explain such discrepancies in diversity and rates of evolution among different genogroups of IHNV. The U genogroup may represent IHNV genotypes with the highest fitness in free-ranging salmonids (Kurath et al., 2003). Recent phylogenetic analyses have revealed the existence of two distinct subgroups (L1 and LII) within the L genogroup (Kelley et al., 2007). While subgroup L1 have restricted distribution within coastal rivers of southern Oregon and

northern California, the subgroup LII is mainly restricted to inland valley watersheds of California such as Sacramento River, San Joaquin River, and their tributaries. Both subgroups were also reported to have contrasting divergence pattern. Similar to U genogroup, the LI has also shown to have slower evolutionary rate, whereas like M genogroup, the LII groups showed rapid evolutionary rate (Kelley et al., 2007). These discrepancies in evolutionary rates could be associated with the habitats. At a global scale, while the European isolates were more closely related to the North American genogroup M, the Japanese and Korean isolates formed a unique cluster with older Japanese isolates more closely related to the North American genogroup U (Padhi and Verghese, 2008; Kim et al., 2007; Nishizawa et al., 2006). A detailed molecular evolutionary analysis is required to understand the evolutionary pattern of these IHNV isolates representing three continents. Purifying selection has been the major driving force in the evolution of a vast majority of codons of IHNV glycoprotein, but approximately 2.8% of the codons have evolved under positive selection (Padhi and Verghese, 2008; LaPatra et al., 2008). Some of these positively selected sites can be mapped to the regions of antigenic determinants of IHNV that have been previously reported (Huang et al., 1994, 1996). The G, N, and NV genes phylogenies (Snow et al., 1999, 2004; Einer-Jensen et al., 2004, 2005) revealed the existence of multiple genogroups of VHSV. The four major genogroups of VHSV include: (I) European freshwater VHSV isolates, a group of marine isolates from the Baltic Sea, North Sea, Skagerrak and Kattegat and the English channel; (II) a group of marine isolates from the Baltic sea; (III) Isolates from the North sea, Skagerrak and Kattegat; and (IV) North American isolates. All Japanese isolates, except one, fall into the North American genogroup, whereas the remaining isolates fall into the European group and are considered to have been introduced from outside Japan (Nishizawa et al., 2002). Like IHNV, VHSV has a strong spatial structure, rather than clustering based on the year of isolation or host species (Snow et al., 1999; EinerJensen et al., 2004). Based on the molecular clock analyses, the time to the most recent common ancestor of the European freshwater isolates is estimated to be 50 years ago, the timing which is consistent with the initial reports in the 1950s on the clinical observations of VHSV in Danish freshwater rainbow trout farms (Jensen, 1965). The VHSV freshwater isolates appear to be evolving 2.5 times faster than the marine isolates (Einer-Jensen et al., 2004), suggesting a relatively recent adaptation of these viruses to freshwater habitats. The successful recent viral adaptation in new hosts is one of the possible explanations for such higher evolutionary rates of VHSV in freshwater fish (Moya et al., 2000). Alternative explanations for the increased substitution rates in freshwater VHSV is the intensive aquaculture practices, as well as the higher water temperature in culture ponds, which could cause an increase in virus replication rates (Einer-Jensen et al., 2004). A similar pattern has also been observed for IHNV in North America, where the evolutionary rate was found to be three to four times higher in regions with intensive aquaculture, as compared with other regions (Troyer and Kurath, 2003). Despite the identification and molecular characterization of HIRRV (Oh and Choi, 1998; Kasornchandra et al., 1992; Johnson et al., 1999; Kim et al., 2005) and SHRV (Nishizawa et al., 1991, 1995), little is known about the intraspecific genealogy of these two viruses. Future studies should focus on understanding the distribution of genetic diversity of HIRRV and SHRV across the geographical range of their hosts. 7. Plant rhabdoviruses Members of two genera of the Rhabdoviridae infect plants and are transmitted via arthropod vectors, such as leafhoppers,

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planthoppers and aphids (Jackson et al., 2005). The cytorhabdoviruses and nucleorhabdoviruses are primarily distinguished based on their sites of virion maturation, in the cytoplasm and the nucleus, respectively. This classification is supported by available genome sequence data. However, as of yet, more than 75 putative members have not been assigned to a genus, because their replication sites remain to be clearly identified and/or genome sequence data are not available. There are currently eight recognized species in the genus Cytorhabdovirus. Lettuce necrotic yellows virus (LNYV) is the type species. Its distribution is restricted to Australia and New Zealand, but appears to be similar to the recently reported lettuce yellow mottle virus (LYMoV), isolated from lettuce in France (Heim et al., 2008). There are seven recognized species in the genus Nucleorhabdovirus. Sonchus yellow net virus (SYNV) is the best-studied member of the genus (type species Potato yellow dwarf virus). Species in both genera are primarily differentiated by plant host range and insect vector specificity. Species and strain demarcation within plant rhabdovirus genera has been aided by several methods. Serology, nucleic acid hybridization, and PCR have been used for some time to verify common species that infect different hosts. Increasingly, genomic sequencing and live cell imaging of intracellular protein localization are providing valuable additional information (Jackson et al., 2005; Goodin et al., 2007). The genomes of four cytorhabdoviruses and five nucleorhabdoviruses have been completely sequenced. Phylogenies of plant rhabdoviruses based on the nucleoprotein gene or the conserved polymerase module of the L gene suggest a monophyletic origin, and place plant-infecting rhabdoviruses into two distinct clades, corresponding to the two genera Cytorhabdovirus and Nucleorhabdovirus, respectively (Jackson et al., 2005). The LNYV clusters with the other sequenced cytorhabdoviruses, with an apparent close evolutionary link to LYMoV and to Strawberry crinkle virus (SCV). The nucleorhabdovirus species fall into two sister clades, members of which appear to be evolutionarily closely related (Jackson et al., 2005; Ghosh et al., 2008). Comparative analyses of sequences within the N and L genes indicate considerable genetic variability between field isolates of LNYV, SCV and taro vein chlorosis virus (TaVCV). Phylogenetic analysis of the complete N gene of 12 LNYV field isolates collected from different host plants and across Australia between 1985 and 2003 revealed two distinct subgroups. Nucleotide sequences within each subgroup were more than 96% identical. The N gene sequences between subgroups differed by about 20% at the nucleotide level, but by less than 4% at the amino acid level (Callaghan and Dietzgen, 2005). This indicates a high level of synonymous versus non-synonymous nucleotide substitutions in the third codon position of the N gene coding sequence. Isolates belonging to either subgroup appear to have co-existed in space and time across Australia, but no subgroup I isolates have been found for 15 years. It remains to be determined if such isolates still occur today, or if they have been replaced by subgroup II isolates (Dietzgen et al., 2007). Evidence for two subgroups among eight European isolates of SCV comes from phylogenetic analysis of partial L gene sequences (Klerks et al., 2004). Nucleotide sequences within each subgroup were 98% identical, but 11% different between subgroups. The groupings appeared to be independent of symptom severity or geographic origin of the isolates. High levels of nucleotide sequence diversity were also found when 1 kb sequences of the N and L genes of TaVCV isolates from six Pacific Island countries were compared (Revill et al., 2005). Nucleotide sequences of the L gene differed by up to 27.4% among 20 isolates; maximum variability within the N gene was 19.3% among 15 isolates. Deduced amino acid sequences for the L and N gene fragments differed by up to 11.3% and 6.3%, respectively. The much-reduced variability of the N gene amino acid sequences

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versus nucleotide sequences for plant rhabdoviruses in both genera indicates evolutionary constraints on variability to maintain primary N protein function. 8. Evolution of rhabdoviruses Rhabdoviruses are distributed worldwide, with no definitive geographical origin. Most are transmitted by arthropods. Given this proximate relationship, one may speculate that initially rhabdoviruses (or their progenitors) were hosted by arthropods (Hogenhout et al., 2003). Subsequently, various groups of rhabdoviruses would have adapted to various plant and vertebrate hosts, with or without arthropod vectors. Unfortunately, this hypothesis is not readily supported from the phylogenetic tree of available rhabdovirus gene sequences. Moreover, for many viral representatives, the sequence data do not exist as of yet. Hence, evolutionary pathways of particular genera are currently impossible to recover. For example, there is no evidence for a virus that constitutes a link between plant and animal rhabdoviruses, or between lyssaviruses and any other genera of rhabdoviruses. Furthermore, estimation of divergence times for rhabdoviruses is highly problematic. Although it is possible to estimate substitution rates for some individual lineages, and for a limited period of time, using molecular clock models (Hughes et al., 2005; Davis et al., 2006, 2007; David et al., 2007), application of these rates and the overall molecular clock approach to longer trees is confounded by substitution saturation and selection pressures that may have occurred during adaptation of progenitor viruses to new species. Therefore, the true age estimations and applicability of molecular clocks for rhabdovirus genome sequences are highly questionable (Kuzmin et al., 2006). For example, the N and G genes of Arctic rabies viruses circulating during the 1950s were 98–99% identical to those of viruses, belonging to the same lineage, but circulating in 2000–2005. At the same time, other lineages of Arctic rabies viruses demonstrated greater diversity even if sampled contemporaneously (Kuzmin et al., 2008a). Similarly, the N and G genes of the LBV isolate, obtained in Kenya in 2007, demonstrated over 99% identity to those of another isolate of LBV, obtained in Senegal in 1985 (Kuzmin et al., 2008b). Furthermore, another LBV lineage, circulating in southern Africa, demonstrates greater sequence identity between isolates obtained in 1980s and 2000s than between isolates obtained during 2003–2006 in the same area (Markotter et al., 2008). Similarly, analysis of partial P sequences (including the hypervariable region) in VSV, showed only 2 nt substitutions among 27 isolates obtained over a 7-year time span on Ossabaw Island (Nichol et al., 1993), but as many as 44 nt substitutions in samples isolated the same year, from different ecological zones, in Costa Rica (Rodriguez et al., 1996). Inferred major forces of rhabdovirus evolution include point mutations and purifying selection. There is no strong supportive evidence of homologous recombination in the family, or within the whole order Mononegavirales, and no suggestions for positive selection towards host adaptation have been reported (Badrane and Tordo, 2001; Hughes et al., 2005; Davis et al., 2005, 2006; Holmes et al., 2002; Guyatt et al., 2003). However, such identification of positive selection depends on the ratios of the non-synonymous to synonymous mutations (dN/dS) and the assumption that non-synonymous mutations will more often have a selective effect than synonymous mutations. Sequence analyses of VSV strains adapting to cell culture have shown that the dN/dS ratios are close to 1, even under conditions where fitness increases steadily (Novella et al., 2004a,b). In addition, synonymous parallel mutations were as common as non-synonymous parallel mutations, suggesting that the dN/dS values alone may be misleading when applied to the identification of the selective forces driving the evolution of rhabdoviruses.

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Rhabdoviruses, as other RNA viruses, are characterized by a high mutation rate during replication (Drake and Holland, 1999) due to the lack of proofreading mechanisms in RNA polymerases (Steinhauer et al., 1992). These high mutation rates confer the potential for rapid evolution, which can be observed readily under laboratory conditions (Clarke et al., 1993; Novella et al., 1995, 1999; Za´rate and Novella, 2004). However, compared to other RNA viruses, such as influenza virus, the genomes of natural populations of rhabdoviruses evolve much more slowly. Several factors may contribute to limiting this relative speed of evolution. Long-term co-evolution and successful adaptation of viruses to their hosts may create bottlenecks or place the viral populations at the top of a fitness peak, and thus limit the potential to generate additional beneficial variation (Jenkins et al., 2002; Bourhy et al., 2005; Za´rate and Novella, 2004; Kuzmin et al., 2006; Presloid et al., 2008). Because of the very limited genome size of rhabdoviruses, individual genomic regions may be responsible for multiple functions. Therefore, a single mutation may have multiple antagonistic effects (Woelk and Holmes, 2002). One obvious example is the P ORF, which also codes for the C and C0 in alternative reading frames. In addition, some coding genomic regions may be involved in specific viral functions, independent of the proteins they encode. Such functions have not been studied in rhabdoviruses, but there are well-characterized examples in other RNA viruses, such as the presence of cis-acting replication signals within the ORFs of rhinovirus (Yang et al., 2002), and encapsidation signals in the phage MS2 (Helgstrand et al., 2002), as well as other cases (for additional examples covering diverse RNA virus species from different families, see Novella, 2003). In addition, antagonistic effects that limit positive selection may arise because of RNA and protein secondary structure (Simmonds and Smith, 1999). Another factor that may lead to an underestimation of positive selection is the relative lack of correlation between fitness increase and accumulated mutations. In vitro adaptation of VSV has resulted in the striking observation that strains that showed a several-fold fitness increase had no changes in the consensus sequence (Novella and Ebendick-Corpus, 2004). Thus, genomic stasis does not necessarily correspond to adaptive stasis. This lack of correlation is due to two factors. First, beneficial variation is usually present within any given population, provided there are enough numbers of particles in the infecting founder sample (Novella et al., 2007; Dutta et al., 2008). This preexisting variation will drive an initial phase of adaptation, but genomes with beneficial mutations may have an overall effect on the fitness of the population before they become dominant or, at least, frequent enough to produce a peak in the chromatogram when the consensus sequence is determined. In this case, beneficial mutations can eventually be identified at later time points (Novella and Ebendick-Corpus, 2004). In a second scenario, it is possible to generate multiple mutations with similar beneficial effects at different loci, that will increase in frequency and replace the parental wild type, but because none will outcompete others (at least for a limited period of time), the consensus will not change, even though the overall fitness varies. Another factor that limits genomic evolution is complementation. Complementation can be defined as the sharing of resources among coinfecting strains, depending on the level of coinfection. Complementation promotes the maintenance of variation and effectively weakens selection because it allows the extended survival of deleterious mutants, and delays the fixation of beneficial mutants (Wilke and Novella, 2003; Novella et al., 2004a,b; Wilke et al., 2004). Complementation is expected to be high among rhabdoviruses that replicate in arthropods, because they typically establish persistent infections that last for life (Tesh et al., 1972) and such conditions promote coinfection.

Preexisting variation may be a confounding factor when phylogeny is used to understand the epidemiology of rhabdoviruses. For example, VSV strains adapted to persistent infection of sand fly cells in culture are maladapted for mammalian BHK-21 cells. Recovery in the latter for only ten passages results in the loss of all mutations accumulated during the adaptation to persistence (including insertions) and the acquisition of a new set of mutations. The most parsimonious explanation for these sequence changes is that the mammalian-adapted mutants were present at a low frequency in the persistence-adapted populations, as confirmed experimentally (Novella et al., 2007). From a biological standpoint, the mammalian-adapted strains originated from infections with the persistence-adapted strains. However, in a phylogenetic tree, the two sequences would belong to very different clades, and appear as unrelated epidemiologically. In general, the maximum diversity in the Rhabdoviridae is seen among the insect-borne plant viruses (Cytorhabdovirus and Nucleorhabdovirus genera), followed by the Novirhabdovirus, isolated from fish and other aquatic animals (including invertebrates), the dimarhabdoviruses (with the possibility to replicate in vertebrate and invertebrate hosts), and finally by the lyssaviruses, which replicate exclusively in mammals (Kuzmin et al., 2006). This observation is opposite to the general suggestion that vector-borne RNA viruses are more constrained than non-vector-borne (Woelk and Holmes, 2002; Jenkins et al., 2002). Rhabdoviral infection, as infections caused by other arboviruses, may have at least two results. As suggested earlier, insect infection is typically persistent, lasting for the life of the insect, and can result in transovarial transmission from infected females to their offspring. Persistent infections are characterized by an initial phase of rapid replication and substantial viral production (acute phase), followed by a decrease in the extent of viral replication to levels that are orders of magnitude lower, but continue for life (persistent phase). A hallmark of these persistent infections is little or no cell killing. In contrast, mammalian infection with these viruses is typically acute and cytolytic. Virus production is massive and occurs rapidly, but ceases with the death of the cells. Experimental studies of VSV in vitro demonstrated that alternate acute replication in insect and mammalian cells did not reduce the rate at which mutations appeared or limit fitness increases (Novella et al., 1999). Alternation between persistent insect infection and mammalian acute infection did not limit the rates of mutation accumulation (Za´rate and Novella, 2004; Presloid et al., 2008). By comparison, Holmes et al. (2002) proposed that the constrains in RABV genes are probably caused by the necessity to be able to replicate in different types of cells within a host, such as neurons, myocytes, fibroblasts and acinar cells of the salivary glands. However, the difference of cell types within mammalian species (and predominantly within the particular representatives of the Carnivora and Chiroptera, which serve as predominant hosts for lyssaviruses) is not as great as difference of cell types between invertebrate and vertebrate hosts, or between plants and insects. Furthermore, with one exception, all studies where fitness analyses were performed on arboviruses adapting to changing environments in cell culture have shown a general lack of limitations in the ability of viruses to adapt to the various environments under investigation (Novella et al., 1999; Weaver et al., 1999; Turner and Elena, 2000; Greene et al., 2005). The one exception is the study of VSV alternation between persistent insect infection and acute mammalian infection, but in this case the alternating viruses evolve as rapidly (and accumulate as many mutations) as the fastest-evolving controls (e.g. the persistent viruses) (Za´rate and Novella, 2004; Presloid et al., 2008). Some authors suggested that low diversity may be a consequence of the young age of lyssaviruses (Bourhy et al., 2005). If so, this would imply that the predominance of purifying selection and strong constrains are due to very rapid (and successful)

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adaptation to mammalian transmission and constraints placed upon lyssaviruses by their unique pathobiology. Unfortunately, to date, there is no evidence reflecting any intermediate steps along this pathobiological adaptation. Fitness is not the only parameter that determines the evolutionary success of a virus. Selection favors high beneficial mutation rates (high adaptability) (Novella, 2004) and low deleterious mutation rates (high robustness) (Forster et al., 2006). Thus, mutations that do not have a fitness effect but decrease robustness or adaptability will not be tolerated when selection prevails and may result in additional limitations to the number of mutations that can be fixed. 9. Concluding remarks With a relatively simple morphology and genome, rhabdoviruses have successfully adapted to a great variety of plant and animal hosts. An improved understanding of their phylogeny and evolution has occurred gradually during the last 30 years. This progression began with concomitant enhanced surveillance and improved methodology for virus detection and characterization. Rapid automatic gene sequencing has facilitated phylogenetic comparisons of viral genomes. However, as of yet, the knowledge on rhabdovirus phylogeny remains incomplete, with multiple described rhabdoviruses awaiting further genetic characterization. Additional surveillance is needed as well, particularly in the poorly investigated tropical areas of the Old and New World, where the family has probably originated. Additional rhabdovirus representatives are certain to be identified. Current understanding of the mechanisms and history of viral evolution are based on operational observations, experimental studies, and extensive modeling efforts. These approaches evolve and improve almost daily, and the presence of multiple contradictive theories suggests that a more comprehensive appreciation of viral evolution is quite far from ideal at this point. Nevertheless, the accumulated data reviewed in this article demonstrate the progress in investigations of rhabdovirus evolution, and serve as a platform for future research. References Amengual, B., Whitby, J.E., King, A., Serra Cobo, J., Bourhy, H., 1997. Evolution of European bat lyssaviruses. J. Gen. Virol. 78, 2319–2328. Arankalle, V.A., Prabhakar, S.S., Hanumaih, W.A.M., Dattatraya, P.S., Chandra, M.A., 2005. G, N and P gene-based analysis of Chandipura viruses. Ind. Emerg. Infect. Dis. 11, 123–126. Badrane, H., Tordo, N., 2001. Host switching in Lyssavirus history from the Chiroptera to the Carnivora orders. J. Virol. 75, 8096–8104. Badrane, H., Bahloul, C., Perrin, P., Tordo, N., 2001. Evidence of two lyssavirus phylogroups with distinct pathogenecity and immunogenecity. J. Virol. 75, 3268–3276. Bai, W., Yan, J., Zhang, Z., JiaNG, c., Lin, Y., 1993. Studies on vaccine against ephemeral fever. In: St.George, T.D., Uren, M.F., Young, P.L., Hoffman, D. (Eds.), Bovine Ephemeral Fever and Related Rhabdoviruses. ACIAR, Canberra, pp. 111–114. Basak, S., Mondal, A., Polley, S., Mukhopadhyay, S., Chattopadhyyay, D., 2007. Reviewing Chandipura: a vesiculovirus in human epidemics. Biosci. Rep. 27, 275–298. Baudin-Laurencin, F., 1987. IHN in France. Bull. Eur. Assoc. Fish Pathol. 7, 104. Bisel, P.A., Nichol, S.T., 1990. Polymerase errors accumulating during natural evolution of the glycoprotein gene of vesicular stomatitis virus Indiana serotype isolates. J. Virol. 64, 4873–4883. Bisel, P.A., Rowe, J.E., Fitch, W.M., Nichol, S.T., 1990. Phosphoprotein and nucleocapsid protein evolution of vesicular stomatitis virus New Jersey. J. Virol. 64, 2498–2504. Bourhy, H., Cowley, J.A., Larrous, F., Holmes, E.C., Walker, P.G., 2005. Phylogenetic relationships among rhabdoviruses inferred using the L polymerase gene. J. Gen. Virol. 86, 2849–2858. Bourhy, H., Kissi, B., Tordo, N., 1993. Molecular diversity of the Lyssavirus genus. Virology 194, 70–81. Bovo, G., Giorgetti, G., Jørgensen, P.E.V., Olesen, N.J., 1987. Infectious hematopoietic necrosis: first detection in Italy. Bull. Eur. Assoc. Fish Pathol. 7, 124.

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