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Peter J. Walker, Rachel Breyta, Kim R. Blasdell, Charles H. Calisher, Ralf G. Dietzgen, Anthony R. Fooks, Juliana Fieitas-Astúa, Hideki Kondo, Gael Kurath, Ivan V. Kuzmin, Ben Longdon, David M. Stone, Robert B. Tesh, Noël Tordo, Nikos Vasilakis and Anna E. Whitfield
A summary of this ICTV Report chapter has been published as an ICTV Virus Taxonomy Profile article in the Journal of General Virology, and should be cited when referencing this online chapter as follows:
Walker, P.J., Blasdell, K.R., Calisher, C.H., Dietzgen, R.G., Kondo, H., Kurath, G., Longdon, B., Stone, D.M., Tesh, R.B., Tordo, N., Vasilakis, N., Whitfield, A.E., and ICTV Report Consortium. 2018, ICTV Virus Taxonomy Profile: Rhabdoviridae, Journal of General Virology, 99:447–448
The family Rhabdoviridae includes 20 genera and 144 species of viruses with negative-sense, single-stranded RNA genomes of approximately 10–16 kb. Virions are typically enveloped with bullet-shaped or bacilliform morphology but non-enveloped filamentous virions have also been reported. The genomes are usually (but not always) single RNA molecules with partially complementary termini. Almost all rhabdovirus genomes have 5 genes encoding the structural proteins (N, P, M, G and L); however, many rhabdovirus genomes encode other proteins in additional genes or in alternative open reading frames (ORFs) within the structural protein genes. The family is ecologically diverse with members infecting plants or animals including mammals, birds, reptiles or fish. Rhabdoviruses are also detected in invertebrates, including arthropods some of which may serve as unique hosts or may act as biological vectors for transmission to other animals or plants. Rhabdoviruses include important pathogens of humans, livestock, fish or agricultural crops.
Table 1.Rhabdoviridae. Characteristics of members of the family Rhabdoviridae.
vesicular stomatitis Indiana virus (AF473864), species Indiana vesiculovirus, genus Vesiculovirus
Bullet-shaped or bacilliform particle 100–430 nm in length and 45–100 nm in diameter comprised of a helical nucleocapsid surrounded by a matrix layer and a lipid envelope. Some rhabdoviruses have non-enveloped filamentous virions.
Negative-sense, single-stranded RNA of 10.8-16.1 kb (unsegmented or bi-segmented).
Ribonucleoprotein (RNP) complexes containing anti-genomic RNA are generated and serve as templates for synthesis of nascent RNP complexes containing genomic RNA.
From capped and polyadenylated mRNAs transcribed processively from each gene (3′ to 5′), sometimes containing multiple ORFs.
Vertebrates, arthropods and plants; many vertebrate and plant rhabdoviruses are arthropod-borne.
20 genera containing 143 species and one unassigned species (Moussa virus). Many rhabdoviruses remain unclassified.
Viruses assigned to each of the 20 genera form a monophyletic clade based on phylogenetic analysis of L sequences. They usually have similar genome architecture, including the number and locations of accessory genes, and have similarities in host range, modes of transmission and/or sites of replication in the cell.
Genus Lyssavirus. Lyssaviruses infect a wide range of mammals including humans in which they can cause fatal encephalitis (rabies). Natural transmission is via saliva, usually though a bite by an infected animal. The genome is relatively simple, containing the genes which encode five structural protein but feature a long 3′-untranslated region (ψ) in the G gene; additional proteins may be expressed from alternative initiation codons in the P gene.
Genus Novirhabdovirus. Novirhabdoviruses infect teleost fish of numerous species in which they can cause severe haemorrhagic disease. Transmission is waterborne and there is also evidence for egg-associated transmission. The genome features an additional gene (NV) that is located between the G gene and L gene. The NV protein appears to be involved in evasion of the host interferon response. Novirhabdoviruses are very distant phylogenetically from fish rhabdoviruses assigned to the genera Perhabdovirus and Sprivivirus (see below).
Genus Perhabdovirus. Perhabdoviruses infect a wide range of teleost fish. They are transmitted through infected water and can cause severe haemorrhagic disease. The genome is relatively simple, containing the five structural protein genes and short intergenic regions. Perhabdoviruses are phylogenetically related to but distinct from fish rhabdoviruses assigned to the genus Sprivivirus (see below).
Genus Sprivivirus. The viruses assigned to this genus infect a wide range of teleost fish. They are transmitted through infected water and can cause severe haemorrhagic disease. The genome of spriviviruses is relatively simple, containing the five structural protein genes and short intergenic regions. Spriviviruses are phylogenetically related to but distinct from fish rhabdoviruses assigned to the genus Perhabdovirus (see above).
Genus Tupavirus. Tupaviruses have been isolated from birds, insectivores and rodents, and there is evidence of infection in other vertebrates. The genome features a long alternative ORF in the P gene and an additional gene encoding a small hydrophobic protein between the M and G genes.
Genus Curiovirus. Curioviruses have been isolated from midges, sandflies and mosquitoes. Vertebrate hosts are largely unknown but there is evidence of infection of birds. The genome features one or more genes located between the M and G genes, and one or more genes located between the G and L genes, including a gene encoding a viroporin-like protein.
Genus Ephemerovirus. Viruses assigned to the genus have been isolated primarily from livestock, mosquitoes or midges. Some cause an acute febrile illness in bovines that is seldom fatal. The genome of ephemeroviruses features multiple genes between the G and L genes encoding accessory proteins including a non-structural class I transmembrane glycoprotein (GNS) and a viroporin (α1).
Genus Hapavirus. This genus comprises viruses that have been isolated from mosquitoes or midges and that infect birds and mammals. The genome of hapaviruses is large and complex, featuring multiple accessory genes between P and M genes, and between G and L genes, usually including a gene encoding a viroporin-like protein.
Genus Ledantevirus. Ledanteviruses infect mammals; many have been isolated from bats or rodents and some (or all) may be transmitted by arthropods. Some have been associated with disease in humans or livestock. The genome is relatively simple but some viruses feature an additional gene between the G and L genes encoding a small protein of unknown function.
Genus Sripuvirus. Viruses assigned to this genus have been isolated from either sandflies or lizards. The genome of sripuviruses features a small protein encoded in a consecutive ORF in the M gene and a small transmembrane protein encoded in an alternative ORF at the start of the G gene.
Genus Tibrovirus. Some tibroviruses infect cattle and water buffalo and are transmitted by midges; several other tibroviruses have been detected in humans but their role in human disease is currently unclear. The genome features two accessory genes between the M and G genes, and a gene encoding a viroporin-like protein between the G and L genes.
Genus Vesiculovirus. Vesiculoviruses infect a wide range of vertebrate hosts and are transmitted by insects; some may also be transmitted amongst vertebrates by direct contact. Several vesiculoviruses cause vesicular stomatitis in livestock and/or have been associated with influenza-like illness and occasional encephalitis in humans. The genome is relatively simple, containing the five structural protein genes and short intergenic regions, but may also include alternative ORFs in the P gene and use of alternative initiation codons in the M gene.
Genus Almendravirus. The viruses assigned to this genus were isolated from mosquitoes and appear to be poorly adapted (or not adapted) to replication in vertebrates. The genome of almendraviruses features an additional gene located between the G and L genes, encoding a small viroporin-like protein.
Genus Alphanemrhavirus. This genus comprises viruses that have been detected by high-throughput sequencing in parasitic nematodes (roundworms of the phylum Nematoda). The genome of alphanemrhaviruses is relatively simple, containing the five structural protein genes, but may include an additional small ORF in the M gene (Mx) overlapping the end of the M ORF. No alphanemrhaviruses have yet been isolated.
Genus Caligrhavirus. Caligrhaviruses have been detected in sea lice (crustaceans in the family Caligidae) in which they appear to cause active infections. The caligrhavirus genome is relatively simple, containing the five structural protein genes, but may include an additional gene (U1) between the G and L genes. No caligrhaviruses have yet been isolated but virions have been observed by electron microscopy.
Genus Sigmavirus. Sigmaviruses are transmitted vertically, each virus infecting a fly of a single species in the families Drosophilidae or Muscidae. Infection results in paralysis or death of flies upon exposure to carbon dioxide. The genome may feature an additional gene (X) located between the M and G genes, encoding a protein of unknown function.
Genus Cytorhabdovirus. Viruses assigned to this genus infect a wide range of plants and are transmitted by arthropod vectors (aphids, planthoppers and leafhoppers) in which they replicate. In plant cells, cytorhabdoviruses replicate in the cytoplasm. Cytorhabdoviruses have an unsegmented genome featuring an additional gene located between the P gene and M gene, encoding a movement protein; some may also encode a viroporin-like protein.
Genus Dichorhavirus. Dichorhaviruses infect plants and are transmitted by Brevipalpus mites. They cause localised lesions on leaves, stems, and fruits of economically important plants such as citrus, coffee and orchids. The genome of dichorhaviruses is bi-segmented: RNA1 contains the N, P, M and G genes, and an additional gene located between the P gene and M gene encoding a putative movement protein; RNA2 contains the L gene. Virions formed in plant cells may lack envelopes.
Genus Nucleorhabdovirus. Nucleorhabdoviruses infect a wide range of plants and are transmitted by arthropod vectors (aphids, planthoppers, leafhoppers) in which they replicate. Nucleorhabdoviruses replicate in the nucleus of infected plant cells. Nucleorhabdoviruses cluster phylogenetically with the bi-segmented dichorhaviruses. They feature an additional gene between the P gene and M gene encoding a movement protein.
Genus Varicosavirus. Varicosaviruses occur naturally in two families of plants (Compositae and Solanaceae) and are transmitted in soil and zoospores of a chytrid fungus, Olpidium brassicae. The genome is bi-segmented: RNA1 contains an ORF encoding a small protein followed by the L gene; RNA2 contains 5 ORFs including the coat protein gene. Virions observed in plant cells are non-enveloped rods resembling intracellular nucleocapsids of other rhabdoviruses.
Enveloped virions have been reported to be in the range of 100–460 nm in length and 45–100 nm in diameter (Hummeler et al., 1967, Hummeler and Koprowski 1969, Nakai and Howatson 1968, Knudson 1973, Francki 1973) (Figure 1.Rhabdoviridae). The longer forms may represent virions fused end-to-end. Defective-interfering (DI) virus particles are proportionally shorter (Huang et al., 1966). Viruses infecting vertebrates are typically bullet-shaped or cone-shaped; however, some rhabdoviruses infecting animals and most plant rhabdoviruses appear bacilliform when fixed prior to staining (Vasilakis et al., 2013, Kurz et al., 1986). In unfixed preparations, they may appear bullet-shaped or pleomorphic. The outer surface of virions (except for the quasi-planar end of bullet-shaped viruses) is covered with projections (peplomers) which are 5–10 nm long and about 3 nm in diameter (Hummeler et al., 1967). They consist of trimers of the viral envelope glycoprotein (G). A honeycomb pattern of peplomers is observed on the surface of some viruses. Internally, the nucleocapsid (30–70 nm in diameter) has helical symmetry and appears to have cross-striations (spacing 4.5–5 nm) in negatively-stained and thin-sectioned virions (Nakai and Howatson 1968, Cartwright et al., 1972, Simpson and Hauser 1966). The nucleocapsid consists of a ribonucleoprotein (RNP) complex comprising the genomic RNA and tightly bound nucleoprotein (N) together with an RNA-dependent RNA polymerase (L) and polymerase-associated phosphoprotein (P). The RNP complex is active for transcription and replication: the N-RNA template is processed by L, which contains various enzymatic activities, and its cofactor P (Emerson and Wagner 1972, Emerson and Yu 1975). In the cytoplasm, the RNP complex is uncoiled and filamentous, about 700 nm in length and 20 nm in diameter (Sokol et al., 1969). In the virion, the lipid envelope containing G interacts with the coiled RNP complex via the matrix protein (M). Filamentous virions reported for some plant rhabdoviruses appear to lack a viral envelope (Dietzgen et al., 2014).
Figure 1.Rhabdoviridae. (A) Negative-contrast electron micrograph of vesicular stomatitis Indiana virus particles. The bar represents 100 nm (Courtesy of P. Perrin). (B) Negative-contrast electron micrograph of RABV defective-interfering (DI) particles. (Courtesy of P. Perrin). (C) Schematic illustration of a rhabdovirus virion and ribonucleocapsid structure. Unravelling of the RNP is illustrative only to show more clearly its association with the L and P proteins (Courtesy of P. Le Mercier).
Reported virion Mr ranges from 0.3–1.0 x 109 and the S20w is in the range 550–1045 S (plant rhabdoviruses usually have larger S20w values) (Neurath et al., 1966, Jackson and Christie 1977). Virion buoyant density is 1.19–1.20 g cm−3 in CsCl and 1.16–1.19 g cm−3 in sucrose (Jackson and Christie 1977, McCombs et al., 1966, Warrington 1965, Sokol et al., 1968). Virus infectivity is rapidly inactivated at 56 °C, or following UV-, gamma- or X-irradiation, or exposure to formalin or to lipid solvents such as detergents (Olitsky and Long 1928, Shechmeister et al., 1962).
Virions typically contain a single molecule of linear, negative-sense single-stranded RNA (Mr 3.4 x 106 to 5.4 x106; approximately 10–16 kb); rhabdoviruses with segmented genomes also occur with each RNA segment encapsidated independently (Kormelink et al., 2011). The RNA typically represents about 1–3% of virion weight (Knudson 1973, McSharry and Wagner 1971, Thomas et al., 1985). The RNA has a 3′-terminal free hydroxyl group and a 5′-triphosphate and is not polyadenylated (Moyer et al., 1975, Ehrenfeld and Summers 1972). The ends have inverted complementary sequences encoding transcription and replication initiation signals (Keene et al., 1979, Li and Pattnaik 1997, Whelan and Wertz 1999). Defective-interfering RNAs, usually substantially shorter than full-length RNA (less than half length), may be identified in RNA recovered from virus populations (Brown et al., 1967). They are usually negative-sense; however, hairpin RNA forms are also found. Defective-interfering RNAs replicate only in the presence of homologous and, occasionally, certain heterologous helper rhabdoviruses which provide the functional genes (Perrault 1981, Perrault and Semler 1979). Full-length positive-sense RNA, which is an intermediate during the replication process, may constitute a significant proportion of a viral RNA population (Soria et al., 1974). Like the full-length negative-sense RNA genome, the anti-genome is tightly bound to N and does not occur as naked RNA.
Virions generally have five structural proteins (designated N, P, M, G and L; see Table 2.Rhabdoviridae for a summary of their locations, masses and functions). The structural proteins represent 65–75% of dry weight of the virion (Knudson 1973, McSharry and Wagner 1971, Thomas et al., 1985). The function(s) of each of these proteins have been determined largely from studies of the model rhabdoviruses, vesicular stomatitis Indiana virus (VSIV) and/or rabies virus (RABV); the same functions are typically assumed to apply to other rhabdoviruses, although this is not often confirmed experimentally. Most rhabdoviruses also encode multiple additional (accessory) proteins but few of the encoded proteins have been characterised. Ephemeroviruses express a class 1a viroporin (α1) and proteins with viroporin-like structures occur commonly in animal rhabdoviruses (Joubert et al., 2014, Walker et al., 2015) and plant cytorhabdoviruses. Ephemeroviruses and some hapaviruses also express large non-structural class I transmembrane glycoproteins (GNS) that are related to the envelope glycoprotein (G) and appear to have arisen by gene duplication (Walker et al., 1992, Wang and Walker 1993, Gubala et al., 2010). Novirhabdoviruses infecting fish express a non-structural protein (NV) that appears to be required for efficient replication and plays a role in evading the host innate anti-viral response (Kurath and Leong 1985, Biacchesi 2011). Plant-adapted viruses have one or more additional non-structural proteins, one of which has been shown to facilitate virus movement between plant cells (Jackson et al., 2005b). Vesiculovirus express two small proteins (C and C′) from an alternative ORF in the P gene (Spiropoulou and Nichol 1993, Peluso et al., 1996); in lyssaviruses, variant forms of P are expressed from alternative initiation codons in the same frame and are involved in modulating the interferon response (Moseley et al., 2007, Chenik et al., 1995).
For certain rhabdoviruses, other nomenclature has previously been used for P (NS, M1 or M2) and M (M1 or M2). The large number and diversity of accessory proteins encoded in rhabdovirus genomes has presented challenges for nomenclature. Some well described accessory proteins have established names that are in common use. However, as the amino acid sequences of most accessory proteins are not highly conserved and their functions are largely unknown, a universal system of nomenclature based on genome location rather than structural or functional homology has been proposed (Walker et al., 2015). According to this system: i) each additional transcriptional unit (other than N, P, M, G and L) is designated U (unknown) followed by a number in the order they appear in the genome in positive polarity (i.e., U1, U2, U3, etc); ii) the first ORF within each transcriptional unit is assigned the same designation as the transcriptional unit; and iii) each subsequent ORF (alternative, overlapping or consecutive) within any transcriptional unit is designated with a letter (i.e., U1x, U1y, U1z). Alternative ORFs are defined as those which occur within the frame of a longer ORF; overlapping ORFs are alternative ORFs that extend beyond the frame of the primary ORF; and consecutive ORFs are those which do not overlap but follow consecutively within the same transcriptional unit. The VSIV C and C′ proteins (55 and 65 amino acids, respectively) are the smallest rhabdovirus proteins known to be expressed in infected cells (Spiropoulou and Nichol 1993, Peluso et al., 1996) and so ORFs ≥ 180 nucleotides may be considered as potentially significant, depending on their location in the transcriptional unit, the Kozak context of the initiation codon and their conservation in multiple virus isolates or related rhabdoviruses (Walker et al., 2015).
Table 2.Rhabdoviridae. Location and functions of rhabdovirus structural proteins.
Location, mass and function
A component of the viral nucleocapsid (ca. 220–240 kDa) responsible for most of the functions required for transcription and replication: RdRP, mRNA 5′-capping, 3′-poly(A) synthesis and protein kinase activities. Observed masses by SDS-PAGE are 150–240 kDa.
Associates into trimers to form the virus surface peplomers (monomer ca. 65–90 kDa). Binds to host cell receptor(s), induces virus endocytosis then mediates fusion of viral and endosomal membranes. G is variously N-glycosylated and palmitoylated; it lacks O-linked glycans and may have hemagglutinin activity. Induces and binds virus-neutralizing antibodies and elicits cell-mediated immune responses. In some cases, G is involved in tropism and pathogenicity.
Major component of the viral nucleocapsid (ca. 47–62 kDa). It associates with full-length negative- and positive-sense genomic RNAs, or defective-interfering RNAs, but not mRNAs. N is an active element of the template, presenting the bases to the polymerase. Newly synthesised N probably modulates the balance between genome transcription and replication by influencing the recognition of the transcription signals. N elicits cell-mediated immune responses and humoral antibodies. In plant nucleorhabdoviruses, N translocates to a sub-nuclear compartment when co-expressed with the cognate P.
A cofactor of the viral polymerase (ca. 20–30 kDa). It is variously phosphorylated and generally migrates by SDS-PAGE
as a protein of about 40–50 kDa; nucleorhabdovirus P migrates faster. P is essential for at least two fundamental functions: (i) it mediates the physical link and the correct positioning of L on the N-RNA template; and (ii) it acts as a chaperone during the synthesis of N, by forming N-P complexes that prevent N from self-aggregation and binding to cellular RNA. During the genome replication process, N is then transferred from these N-P complexes to the nascent viral RNA to ensure its specific encapsidation into new RNPs. P elicits cell-mediated immune responses. In several rhabdoviruses P also plays a fundamental role in evading the host innate anti-viral response.
A basic protein that is an inner component of the virion (ca. 20–30 kDa). It is believed to regulate genome RNA transcription. M binds to nucleocapsids and the cytoplasmic domain of G, thereby facilitating the process of budding. It is sometimes phosphorylated or palmitoylated. M is found in the nucleus and inhibits host cell transcription. It also mediates other pathological effects (cell rounding for VSIV, apoptosis for RABV, intracellular accumulation of the inner nuclear membrane for potato yellow dwarf virus (PYDV).
Virions are composed of about 15–25% lipid, with their composition reflecting that of the host cell membrane where virions bud (Knudson 1973, McSharry and Wagner 1971, Thomas et al., 1985). Generally, phospholipids represent about 55–60%, and sterols and glycolipids about 35–40% of the total lipids. G may have covalently associated fatty acids proximal to the lipid envelope (Schmidt and Schlesinger 1979, Gaudin et al., 1991).
Virions are composed of about 3% carbohydrate by weight (Knudson 1973, McSharry and Wagner 1971, Thomas et al., 1985). The carbohydrates are present as N-linked glycan chains on G and as glycolipids. Ephemerovirus GNS is also N-glycosylated (Walker et al., 1991). In mammalian cells, the oligosaccharide chains are generally of the complex type; in insect cells they are of non-complex types (Jarvis and Finn 1995). The number and location of N-glycosylation sites varies for G of different rhabdoviruses.
Rhabdovirus genomes contain at least five ORFs in the negative-sense genome in the order 3′-N-P-M-G-L-5′ (Kuzmin et al., 2009, Walker et al., 2011). The genes are flanked by conserved transcription initiation and termination/polyadenylation signals, about 10 nt in length. For many rhabdoviruses, additional genes are interposed between the structural protein genes and alternative, overlapping or consecutive ORFs may occur within the structural protein genes or in the additional genes. Some rhabdovirus genomes are segmented. Consequently, genomes of viruses assigned to different genera may vary greatly in length and organisation (Figure 2.Rhabdoviridae).
Figure 2.Rhabdoviridae. Schematic representation of rhabdovirus genome organizations exemplifying variations in the number and location of accessory genes. A typical member of each genus is represented. Each arrow indicates the position of a long open reading frame (ORF). Other alternative ORFs occur in some genes; only ORFs (≥180 nt) that appear likely to be expressed are shown.
Most understanding of rhabdovirus replication and transcription has been obtained from studies of vesiculoviruses and lyssaviruses (Banerjee 1987, Finke and Conzelmann 2005, Banerjee and Barik 1992). Genes are transcribed sequentially (from 3′ to 5′ from the template virus RNA and in decreasing molar abundance) as 5′-capped, 3′-polyadenylated, monocistronic mRNAs (Moyer et al., 1975, Ehrenfeld and Summers 1972, Abraham et al., 1975, Abraham and Banerjee 1976, Ehrenfeld 1974, Banerjee and Rhodes 1973, Naito and Ishihama 1976) (Figure 3.Rhabdoviridae). A short uncapped, non-polyadenylated and untranslated leader RNA, corresponding to the complement of the 3′-terminus of the viral RNA (i.e., preceding the N mRNA), is also transcribed (Colonno and Banerjee 1976, Colonno and Banerjee 1978). Unlike mRNAs, leader RNA has a 5′-triphosphate terminus (Figure 3.Rhabdoviridae). Leader RNA of some viruses has been identified in the nucleus of infected cells. The mRNAs generally have common 5′-terminal sequences corresponding to the cap structure fused to the first nucleotides copied from the transcription initiation signal. The mRNAs also each contain a 3′-poly(A) tail which is produced by the viral transcriptase upon copying in a reiterative mode at uridine residues present in each transcription termination signal (Naito and Ishihama 1976). Very long 3′-untranslated regions (up to 750 nt) occur in some mRNAs (e.g., lyssaviruses, ephemeroviruses and hapaviruses) (Walker et al., 2015). Intergenic sequences are generally short but may be up to about 100 nt in length. In some cases, the transcription initiation signal of one gene overlaps the 3′-end of the preceding gene.
Figure 3.Rhabdoviridae. Genome organization, transcription and replication of vesicular stomatitis Indiana virus. Top: genome structure Middle: process of consecutive transcription of leader RNA and messenger RNAs. The role of N (green circles), P (red blob) and L (grey oval) is indicated. Bottom: replication of the negative-sense genome (light green N) via a positive-sense anti-genome intermediate (dark green N). The switch from transcription to replication is regulated by N. The genome and anti-genome strands are not generated in equimolar amounts.
Non-canonical mechanisms of translation from alternative, overlapping of consecutive ORFs appear to occur commonly in viruses assigned to some genera. Although not yet demonstrated experimentally, the likely mechanisms include: i) leaky ribosomal scanning; ii) a stop-start mechanism involving overlapping or consecutive termination and initiation codons and a ‘termination upstream ribosome-binding site’ (TURBS); and iii) ribosomal frame shifts featuring a ‘slippery’ sequence followed by a predicted pseudoknot structure (Walker et al., 2015). In the case of some rhabdoviruses, polycistronic mRNAs result from the read-through of the transcription termination signal, allowing transcription extension across the adjacent 5′-gene. However, in most cases, this appears to be due to corruption of the transcription termination signal during adaptation to growth in cell culture.
Except for plant rhabdoviruses, which generally penetrate plant cells through mechanical damage caused by insect vectors, rhabdovirus adsorption is mediated by G attachment to cell surface receptors, and penetration of the cell occurs by endocytosis via coated pits (Regan and Whittaker 2013, Albertini et al., 2012). Various candidate receptors have been postulated for RABV (nicotinic acetylcholine receptor AChR, neural cell adhesion molecule NCAM, low affinity nerve growth factor receptor p75NTR), VSIV (phosphatidyl serine), viral hemorrhagic septicemia virus (VHSV) (fibronectin), and others (Lafon 2005, Schlegel et al., 1983, Bearzotti et al., 1999). In addition, carbohydrate moieties, phospholipids and gangliosides may play a complementary role for virus binding (Coil and Miller 2004, Superti et al., 1986, Superti et al., 1984). After penetration by endocytosis, low pH within the endosome triggers fusion between endosomal and viral membranes, liberating the RNP complex into the cytoplasm. The pH-induced fusion depends on conformational changes of the glycoprotein, a process that is reversible upon raising the pH (Roche et al., 2008, Roche and Gaudin 2004). Once the nucleocapsid is released into the cytoplasm, the RNA genome is repetitively transcribed (primary transcription) by the virion transcriptase (Banerjee 1987). N removal does not occur since the transcriptase only recognizes the RNA-N protein complex as template (Arnheiter et al., 1985). The capped and polyadenylated mRNAs are generally translated in cytoplasmic polysomes, except for the G mRNA which is translated on membrane-bound polysomes (Both et al., 1975, Grubman et al., 1975, Morrison and Lodish 1975). Transcription occurs in the presence of protein synthesis inhibitors, indicating that it does not depend on de novo host protein synthesis (Villarreal and Holland 1974, Marcus et al., 1971). Following translation, RNA replication occurs in the cytoplasm (full-length positive-sense and then full-length negative-sense RNA synthesis).
Nucleorhabdoviruses and dichorhaviruses replicate in viroplasms in the cell nucleus (van Beek et al., 1985, Redinbaugh et al., 2002, Jackson et al., 2005a, Kondo et al., 2013, Kitajima et al., 2001). Replication again occurs on the RNA-N protein complex and requires the newly synthesised N, P and L species to concomitantly encapsidate the nascent RNA into a nucleocapsid structure. Apart from freshly translated N, P and L, replication may require host factors. Vesiculoviruses can replicate in enucleated cells, indicating that newly synthesised host gene products are not required (Follett et al., 1974, Wiktor and Koprowski 1974). However, as for some other negative-sense RNA viruses, trafficking of rabies virus proteins to and from the nucleus appears to play an important role in pathogenesis and modulating the host immune response to infection (Audsley et al., 2016, Wiltzer et al., 2012).
It has been proposed that the concomitant binding of N to the nascent positive- or negative-sense viral RNA species may promote replication rather than transcription, by favoring read-through of transcription termination signals (Arnheiter et al., 1985, Blumberg et al., 1981). Replication leads to the synthesis of a full-length positive-sense anti-genome RNA. This, in turn, serves as a replicative intermediate for the synthesis of negative-sense genome RNA for the progeny virions. Following replication, further rounds of transcription (secondary transcription), translation and replication ensue. A typical feature of negative-sense RNA viruses (shared by all members of the order Mononegavirales) is that the RNA genome (or anti-genome) is never “naked” in the cell but is always encapsidated by the nucleoprotein. This RNA-N complex is the true template recognised by the viral polymerase (transcriptase or replicase) (Emerson and Wagner 1972, Moyer et al., 1991).
Post-translational trafficking and modification of G involves translocation across the endoplasmic reticulum membrane, removal of the amino-proximal signal sequence and step-wise glycosylation in compartments of the Golgi apparatus (Rothman and Lodish 1977, Zilberstein et al., 1980). Depending on the cell, G may move to the plasma membrane, particularly to the basolateral surfaces of polarised cells (Fuller et al., 1984, Pfeiffer et al., 1985).
Viral nucleocapsid structures are assembled in association with M and lipid envelopes containing viral G to form virions (Mebatsion et al., 1999). The site of formation of particles depends on the virus and host cell. For vesiculoviruses, lyssaviruses, ephemeroviruses and novirhabdoviruses, nucleocapsids are synthesised in the cytoplasm and virus particles bud from the plasma membrane in most, but not all cells. Some lyssaviruses produce particles that bud predominantly from intracytoplasmic membranes and in some cases prominent virus-specific cytoplasmic inclusion bodies containing N are observed in infected cells (RABV inclusion bodies are called Negri bodies) (Manghani et al., 1986, Matsumoto 1962, Matsumoto et al., 1974, Lahaye et al., 2009). Cytorhabdovirus virions bud from intracytoplasmic membranes associated with viroplasms; none have been observed to bud from plasma membranes (Jackson et al., 2005a). Nucleorhabdovirus and dichorhavirus virions bud from the inner nuclear membrane and accumulate in the perinuclear space (van Beek et al., 1985, Redinbaugh et al., 2002, Jackson et al., 2005a).
Depending on the virus and host cell type, rhabdovirus infections may inhibit cellular protein synthesis and cause apoptosis by mechanisms that are mediated by M (Koyama 1995, Kopecky et al., 2001, Weck and Wagner 1979, Jackson and Rossiter 1997, Larrous et al., 2010, Faria et al., 2005). Complementation between viral mutants of related viruses may occur (e.g., between vesiculoviruses), but not between viruses assigned to different genera (Pringle et al., 1971, Repik et al., 1976). Complementation has also been reported to occur by re-utilisation of the structural components of UV-irradiated virus (VSIV). Inter-molecular genetic recombination between different virus isolates is very rare, but intra-molecular recombination may occur during the formation of defective-interfering RNAs. Phenotypic mixing occurs between some animal rhabdoviruses and other enveloped animal viruses (e.g., paramyxoviruses, orthomyxoviruses, retroviruses, herpesviruses).
G induces virus-neutralising antibodies which define viruses as serotypes and can provide protective immunity. Antigenic cross-reactions in complement-fixation or indirect immunofluorescence tests occur primarily between rhabdoviruses within a genus and involve antigenic determinants located on the N protein. Cross-reactions in indirect immunofluorescence tests have also been detected between some animal rhabdoviruses that are now assigned to different genera (Calisher et al., 1989, Tesh et al., 1983).
Rhabdoviruses are ecologically diverse with members infecting plants or animals including mammals, birds, reptiles or fish (Kuzmin et al., 2009). Some of the vertebrate rhabdoviruses have a wide experimental host range; rhabdoviruses infecting plants usually have a narrow host range among higher plants. Rhabdoviruses are also detected in invertebrates, including many arthropods, some of which may serve as biological vectors for transmission to animals or plants. A diverse range of vertebrate and invertebrate cell lines are susceptible to vertebrate rhabdoviruses in vitro.
Rhabdoviruses are not usually transmitted vertically in vertebrates or plants, but transovarial transmission has been documented in insects. Sigmaviruses were recognised first as a congenital infection in fruit flies. Vector transmission may involve mosquitoes, sandflies, midges, aphids, leafhoppers or planthoppers. Some viruses are transmitted mechanically in sap or from the body fluids of infected hosts. Mechanical transmission of viruses infecting vertebrates may be by contact, aerosol, bite, or venereal. Fish rhabdoviruses can be transmitted by exposure to infected water.
Twenty genera have been established to date. Viruses assigned to a genus form a monophyletic clade in well supported Maximum Likelihood trees using full-length L sequences. Use of L for taxonomic purposes is justified by the presence of broadly conserved domains and the rarity of genetic recombination. Demarcation of genera is based upon considerations of significant differences in genome architecture, antigenicity and ecological properties (such as host range, pathobiology and transmission patterns).
Rhabdoviridae: from rhabdos (Greek) meaning rod, referring to virion morphology.
Phylogenetic relationships across the family have been established from Maximum Likelihood trees generated from conserved regions of phylogenetically informative sequence in L (Figure 4.Rhabdoviridae). These can be identified by aligning full-length L sequences and eliminating ambiguously aligned regions using the Gblocks algorithm (http://molevol.cmima.csic.es/castresana/Gblocks_server.html). Phylogenetic relationships between viruses assigned to more closely related genera and within genera can also be established using other structural protein genes, notably N and G.
Figure 4.Rhabdoviridae. A Maximum Likelihood phylogenetic tree inferred from a MUSCLE alignment of the full-length L sequences of 134 rhabdoviruses assigned to 20 genera and one rhabdovirus (Moussa virus) representing an unassigned rhabdovirus species. Full-length L sequences are not currently available for other rhabdoviruses assigned to species and so they have not been included in data set. Ambiguously aligned amino acid residues were pruned using Gblocks (Castresana 2000) with 442 positions remaining in the final dataset. The evolutionary history was inferred by using the WAG + frequency model of amino acid substitution (Whelan and Goldman 2001) with sub-tree pruning and re-grafting (SPR) branch-swapping. The initial tree for the heuristic search was obtained automatically by applying the neighbour-joining algorithm to a matrix of pairwise distances estimated using a JTT model, and then selecting the topology with superior log likelihood value. The tree with the highest log likelihood (-43388.77) is shown. Asterisks (*) indicate well-supported nodes in the tree (bootstrap proportion ≥ 75%) following 1000 iterations. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site, the scale bar indicating a value of 0.5. Evolutionary analyses were conducted in MEGA7 (Kumar et al., 2016). This phylogenetic tree and corresponding sequence alignment are available to download from the Resources page.
Many general characteristics of rhabdovirus genome organisation, replication and transcription are shared with other members of the order Mononegavirales.
Table 3.Rhabdoviridae. Unclassified rhabdoviruses (additional unclassified rhabdoviruses that are probable members of existing genera are listed under individual genus descriptions).
American dog tick rhabdovirus 1
American dog tick rhabdovirus 2
Apis rhabdovirus 1
Apis rhabdovirus 2
Bahia Grande virus
BeAn 157575 virus
Beihai dimarhabdovirus 1
Beihai barnacle virus 7
blacklegged tick rhabdovirus 1
blue crab virus
Bole tick virus 2
Caledonia dog whelk rhabdo-like virus 2
Culex rhabdo-like virus
Culex tritaeniorhynchus rhabdovirus
DakArk 7292 virus
Diachasminorpha longicaudata rhabdovirus
Dillard’s Draw virus
Drosophila busckii sigmavirus
Drosophila sturtevanti rhabdovirus 1
Drosophila subobscura rhabdovirus
eel virus B12
eel virus C26
fox fecal rhabdovirus
Grenada mosquito rhabdovirus 1
Harrison Dam virus
Huangpi tick virus 3
Hubei dimarhabdovirus virus 2
Hubei dimarhabdovirus virus 3
Hubei dimarhabdovirus virus 4
Hubei lepidoptera virus 2
Hubei myriapoda virus 7
Hubei rhabdo-like virus 1
Hubei rhabdo-like virus 2
Hubei rhabdo-like virus 9
Humpty Doo virus
hybrid snakehead virus
Jingshan fly virus 2
Lone Star tick rhabdovirus
Long Island tick rhabdovirus
Lye Green virus
Muir Springs virus
murine feces-associated rhabdovirus
Nayun tick rhabdovirus
New Minto virus
Norway mononegavirus 1
Norway mononegavirus 1-like virus
North Creek virus
Oak vale rhabdovirus
Pararge aegeria rhabdovirus
Reed Ranch virus
Rhipicephalus associated rhabdo-like virus
Rhode Island virus
Rio Grande cichlid virus
Santa Barbara virus
Sanxia water strider virus 5
Shayang ascaridia galli virus 2
Shayang fly virus 3
Shuangao bedbug virus 2
Shuangao insect virus 6
soybean cyst nematode-associated northern cereal mosaic virus
Spodoptera frugiperda rhabdovirus
Tacheng tick virus 3
Tacheng tick virus 7
Taishun tick virus
Tetrastichus brontispae RNA virus 1
ulcerative disease rhabdovirus
Walkabout Creek virus
Wenling crustacean virus 10
Wenling crustacean virus 11
Wenling dimarhabdovirus 1
Wenling dimarhabdovirus 8
Wenling dimarhabdovirus 9
Wenling dimarhabdovirus 10
Wuhan ant virus
Wuhan fly virus 3
Wuhan house fly virus 2
Wuhan insect virus 7
Wuhan louse fly virus 11
Wuhan mosquito virus 9
Wuhan pillworm virus 2
Wuhan tick virus 1
Xinjiang tick rhabdovirus
Xinzhou dimarhabdovirus virus 1
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