African Swine Fever Virus 168Clinical Features and Epidemiology 169Pathogenesis and Pathology 170
Diagnosis 171Immunity, Prevention and Control 172
Members of the Family Iridoviridae 172Properties of Iridoviruses 172Ranaviruses 173Megalocytiviruses 175Lymphocystiviruses 175Other Iridoviruses of Fish 176Iridoviruses of Mollusks 176
Viruses in the families Asfarviridae and Iridoviridae are taxonomically and biologically distinct, but both fami-lies include large viruses with highly complex genomes of double-stranded DNA that are distantly related to one another, as well as to other large DNA viruses in the fam-ily Poxviridae and the order Herpesvirales (Figure 8.1). African swine fever virus in the family Asfarviridae is the cause of African swine fever, an important disease that remains a serious threat to swine industries throughout the world. The family Iridoviridae includes numerous viruses in several genera that have been isolated from poikilothermic animals, including fish, arthropods, mol-lusks, amphibians, and reptiles. Many iridovirus infections are subclinical or asymptomatic, but individual viruses are the cause of important and emerging diseases of fish and amphibians.
MeMbers of the faMily AsfArviridAe
ProPerties oF AsFArviruses
Classification
African swine fever virus is a large enveloped DNA virus that is the sole member of the genus Asfivirus within the family Asfarviridae (Asfar African swine fever and related viruses). African swine fever virus is the only known DNA arbovirus and is transmitted by soft ticks of the genus Ornithodoros. Virus strains are distinguished by their viru-lence to swine, which ranges from highly lethal to subclinical
disease. Strains can also be differentiated by their genetic sequences, and the virus-encoded p72 (also referred to as p73) gene can be used for genotyping the virus; however, the genomic diversity of the virus in nature remains to be
thoroughly characterized. The genome of African swine fever virus contains a unique complement of multigene families.
virion Properties
Asfarvirus virions are enveloped, approximately 200 nm in diameter, and possess a nucleocapsid core that is sur-rounded by internal lipid layers and a complex icosahedral capsid (Figure 8.2; Table 8.1). The capsid consists of a hex-agonal arrangement of structural units, each of which appears as a hexagonal prism with a central hole. The genome con-sists of a single molecule of linear double-stranded DNA, 170–190 kbp in size, depending on the virus strain. The DNA has covalently closed ends with inverted terminal repeats and hairpin loops, and includes approximately 150 open read-ing frames that are closely spaced and read from both DNA strands. More than 50 proteins are present in virions, including a number of enzymes and factors required for early messen-ger RNA (mRNA) transcription and processing.
African swine fever virus is thermolabile and sensitive to lipid solvents. However, the virus is very resistant to a wide range of pH (several hours at pH 4 or pH 13), and survives for months and even years in refrigerated meat.
virus replication
Primary isolates of African swine fever virus replicate in swine monocytes and macrophages. After adaptation, some isolates can replicate in certain mammalian cell lines. Replication occurs primarily in the cytoplasm, although the nucleus is needed for viral DNA synthesis and viral DNA is present in the nucleus soon after infection. Virus enters susceptible cells by receptor-mediated endocytosis, and cell binding and neutralization studies suggest that the viral p72 and p54 proteins are involved in virus attachment, and p30 in virus internalization. Like that of poxviruses, virion genomic DNA includes genes for all the machin-ery necessary for transcription and replication: after entry into the cytoplasm, virions are uncoated and their DNA is transcribed by a virion-associated, DNA-dependent RNA polymerase (transcriptase). DNA replication is similar to that of poxviruses: parental genomic DNA serves as the template for the first round of DNA replication, the prod-uct of which then serves as a template for the synthesis of large replicative complexes that are cleaved to produce mature virion DNA. Late in infection, African swine fever virus produces paracrystalline arrays of virions in the cyto-plasm. Infected cells form many microvillus-like projec-tions through which virions bud; however, acquisition of an envelope is not necessary for viral infectivity.
AFriCAn swine Fever virus
African swine fever was considered a disease of only sub-Saharan Africa until 1957, when an outbreak occurred on the Iberian Peninsula. Sporadic outbreaks subsequently occurred in the 1970s in some Caribbean islands, including Cuba and the Dominican Republic, and the virus appeared
(A) (B)
(C)
(D)
Figure 8.2 Family Asfarviridae, genus Asfivirus, African swine fever virus. (A) Negatively stained virion showing the hexagonal outline of the capsid enclosed within the envelope. (B and C) Negatively stained dam-aged capsids showing the ordered arrangement of the very large number of capsomers (between 1892 and 2172 structural units) that make up the capsid. (D) Thin section of three virions showing multiple layers sur-rounding their cores. Bars: 100 nm. (Courtesy of J. L. Carrascosa.)
table 8.1 Properties of Asfarviruses and Iridoviruses
Asfarvirus virions are enveloped, approximately 200 nm in diameter, and contain a complex icosahedral capsid, approximately 180 nm in diameter
The genome of African swine fever virus is a single molecule of linear double-stranded DNA, approximately 170–190 kbp in size. It has covalently closed ends with inverted terminal repeats and hairpin loops, and encodes approximately 150 proteins, more than 50 of which are included in virions
Vertebrate iridovirus virions are similar in morphology to those of asfarviruses: the genome is a single molecule of linear double-stranded DNA, 140–200 kbp in size, that encodes up to 200 proteins. It is permuted circularly and has terminally redundant ends and methylated bases
The nucleus is involved in DNA replication; late functions and virion assembly occur in the cytoplasm
Chapter | 8 Asfarviridae and Iridoviridae 169
in France, Belgium, and other European countries in the 1980s. Since 2007, African swine fever virus has spread throughout portions of Georgia, Armenia, Azerbaijan, and Russia. The disease remains enzootic in sub-Saharan Africa and Sardinia. The presence of wild boar and extensive pig farming sustain enzootic African swine fever in Sardinia.
African swine fever virus infects domestic swine and other members of the family Suidae, including warthogs (Potamochoerus aethiopicus), bush pigs (P. porcus), and wild boar (Sus scrofa ferus). All efforts to infect other ani-mals have been unsuccessful. The virus may have origi-nated as a virus of ticks: in Africa, numerous isolates have been made from the soft tick Ornithodoros moubata col-lected in warthog burrows. When African swine fever virus was believed to be confined to sub-Saharan Africa, it was assumed that this was because of its natural cycle in argasid ticks and wild swine; however, the virus has spread on occasion beyond this traditional range and invaded por-tions of Europe, where the soft tick Ornithodoros erraticus can potentially serve as a vector.
Clinical Features and epidemiology
The acute or hyperacute form of African swine fever in susceptible swine is characterized by a severe, hemorrhagic disease with high mortality. After an incubation period of 5–15 days, swine develop fever (40.5–42°C), which per-sists for about 4 days. Starting 1–2 days after the onset of fever, there is inappetence, diarrhea, incoordination, and prostration. Swine may die at this stage without other clini-cal signs. In some swine there is dyspnea, vomiting, nasal and conjunctival discharge, reddening or cyanosis of the ears and snout, and hemorrhages from the nose and anus. Pregnant sows often abort. Mortality is often 100%, with domestic swine dying within 1–3 days after the onset of fever. In prior epizootic geographic extensions of African
swine fever virus infection, the disease was usually severe and fatal at first, but diminished quickly until cases were predominantly subclinical and persistent. Infected adult warthogs do not develop clinical disease.
Two distinct patterns of transmission occur: a sylvatic cycle in warthogs and ticks in Africa, and epizootic and enzootic cycles in domestic swine (Figure 8.3).
Sylvatic Cycle
In its original ecologic niche in southern and eastern Africa, African swine fever virus is maintained in a sylvatic cycle involving asymptomatic infection in wild pigs (warthogs and, to a lesser extent, bush pigs) and argasid ticks (soft ticks, genus Ornithodoros), which occur in the burrows used by these animals. Ticks are biological vectors of the virus. Most tick populations in southern and eastern Africa are infected, with infection rates as high as 25%. After feeding on viremic swine, the virus replicates in the gut of the tick and subse-quently infects its reproductive system, which leads to trans-ovarial and venereal transmission of the virus (primarily male to female tick). The virus is also transmitted between developmental stages of the tick (trans-stadial transmission), and is excreted in tick saliva, coxal fluid, and Malpighian excrement. Infected ticks may live for several years, and are capable of transmitting disease to swine at each blood meal.
Serologic studies indicate that many warthog popula-tions in southern and eastern Africa are infected. After primary infection, young warthogs develop viremia suffi-cient to infect at least some of the ticks feeding on them. Older warthogs are persistently infected, but are seldom viremic; it is therefore likely that the virus is maintained in a cycle involving young warthogs and ticks. The pri-mary source of virus in epizootics of African swine fever in southern and eastern Africa are infected ticks that are transported by live warthogs or their carcasses.
Adult warthogs
Tick to-tick-transmission
Juvenile warthogs
Persistent infection of ticks
Sylvaticcycle
Domesticcycle
• no viremia• virus in various lymphoid tissues
• significant viremia
• trans-stadial• transovarial• sexual
Figure 8.3 Patterns of transmis-sion of African swine fever virus. [From E. R. Tulman, G. A. Delhon, B. K. Ku, D. L. Rock. African swine fever virus. Curr. Top. Microbiol. Immunol. 328, 43–87 (2009), with permission.]
PArt | ii Veterinary and Zoonotic Viruses170
Domestic Cycle
Primary outbreaks of African swine fever in domes-tic swine in Africa probably result from the bite of an infected tick, although tissues of acutely infected warth-ogs, if eaten by domestic swine, can also cause infection. Introduction of the virus into a previously non-infected country may result in transmission amongst swine, as well as infection of indigenous ticks. Several species of soft tick found in association with domestic and feral swine in the western hemisphere have been shown in experimental studies to be capable of biological transmis-sion of the virus, although there is no evidence that they became infected during the epizootics in the Caribbean islands and South America.
Once the virus has been introduced into domestic swine, either by the bite of infected ticks or through infected meat, infected animals constitute the most important source of virus for susceptible swine. High titers of virus are present in nasopharyngeal excretions during onset of clinical signs, and virus is also present in other excretions, includ-ing high amounts in feces during acute disease. Disease spreads rapidly by contact and within buildings by aerosol. Mechanical spread by people, vehicles, and fomites is pos-sible because of the stability of the virus in swine blood, feces, and tissues.
The international spread of African swine fever virus has been linked to feeding scraps of uncooked meat from
infected swine. When the virus appeared in Portugal in 1957 and in Brazil in 1978, it was first reported in the vicinity of international airports, among swine fed on food scraps. Virus spread to the Caribbean and Mediterranean islands in 1978 may have arisen from the unloading of infected food scraps from ships. The source of the virus responsible for the outbreak in Georgia in 2007 is uncertain, although food waste from ships in Black Sea ports is suspected.
Pathogenesis and Pathology
African swine fever virus infection of domestic swine results in leukopenia, lymphopenia, thrombocytopenia, and apopto-sis of both lymphocytes and mononuclear phagocytic cells. The ability of African swine fever virus to efficiently induce cytopathology in macrophages is a critical factor in viral virulence. In infected macrophages, the virus effectively inhibits the expression of pro-inflammatory cytokines such as tissue necrosis factor (TNF), type 1 interferon (IFN), and interleukin-8, but induces expression of transform-ing growth factor . In contrast, increased expression of TNF has been also reported after African swine fever virus infection in vitro and in vivo. Importantly, African swine fever virus strains with different virulence phenotypes differ in their ability to induce (or inhibit) expression of pro-inflammatory cytokines or IFN-related genes early in infection of macrophages (Figure 8.4). Inhibition of
ASFV
Macrophage
Mitochondria
B318LS273R
ER
dUTPase
CD69/NKG2 CD2
TNK
E
MGF360/530
ApoptosiselF2a
PP1a8DR8CR
4CLIAP
UBCv1
Caspase 3
UKNL
TK
9GL
5HLBcl-2
ALR/ERV1
?
?
??
5EL IκB
NFκBCaN
Cellulartranscription
SMCp?
NFAT
Cytokines
(–)TNF-αINF-αIL-8
(+)TNF-αTGF-β
IL-4IL-10
p36
?
Macrophage
Figure 8.4 African swine fever virus (ASFV) – macrophage interactions in the swine host. ASFV contains several genes (white boxes) that interact or potentially interact with cellular regulatory pathways in macrophages, the primary target cells infected by ASFV. A viral homologue of IB (5EL) inhibits both NFB and calcineurin (CaN)/NFAT transcrip-tional pathways. The SMCp DNA-binding domain protein is a possible substrate for viral ubiquitin con-jugating enzyme (UBCv1), and viral Bcl-2 and IAP homologues (5HL and 4CL, respectively) exhibit anti-apoptotic properties. ASFV infection affects host immune responses through induction of apoptosis in uninfected lymphocytes, through modulation of cytokine expression, and potentially through 8CR and 8DR, which are virally encoded homologues of immune cell proteins such as CD2 and CD69/NKG2. Efficient virus assembly and viral production in mac-rophages requires or may utilize viral genes similar to cellular ALR/ERV1 (9GL), nucleotide metabo-lism enzymes (dUTPase and thymidine kinase, TK), SUMO-1-specific protease (S273R) and trans- geranylgeranyl-diphosphate synthase (B318L). ASFV genes that affect viral virulence in domestic swine include NL, UK, and members of the MGF360 and MGF530 multigene families. [From E. R. Tulman, D. L. Rock. Novel virulence and host range genes of African swine fever virus. Curr. Opin. Microbiol. 4, 456–461 (2001), with permission.]
Chapter | 8 Asfarviridae and Iridoviridae 171
inflammation is mediated at least in part by the viral gene A238L, which encodes a protein that is similar to an inhibi-tor of the cellular transcription factor, nuclear factor B (NFB). This viral protein has been shown to inhibit acti-vation of NFB and thus downregulate the expression of all of the antiviral cytokines that are controlled by NFB. Mechanistically, the A238L protein acts as an analog of the immunosuppressive drug cyclosporin A, which represents a novel viral immune evasion strategy. Furthermore, this pro-tein may be central to expression of fatal hemorrhagic disease in domestic pigs but mild, persistent infection in its natu-ral host, the African warthog. Additional proteins encoded by African swine fever virus also modulate host immune responses; these include 8DR (pEP402R), a viral homolog of cellular CD2 involved in T lymphocyte activation and medi-ation of hemadsorption by cells infected with African swine fever virus.
If infection is acquired via the respiratory tract, the virus replicates first in the pharyngeal tonsils and lymph nodes draining the nasal mucosa, before being dis-seminated rapidly throughout the body via a primary viremia in which virions are associated with both eryth-rocytes and leukocytes. A generalized infection follows, with very high virus titers (up to 109 infectious doses per ml of blood or per gram of tissue), and all secre-tions and excretions contain large amounts of infectious virus.
Swine that survive the acute infection may appear healthy or chronically diseased, but both groups may remain persistently infected. Indeed, swine may become persist-ently infected without ever showing clinical signs. The dura-tion of the persistent infection is not known, but low levels of virus have been detected in tissues more than a year after exposure.
In acutely fatal cases in domestic swine, gross lesions are most prominent in the lymphoid and vascular systems (Figure 8.5). Hemorrhages occur widely, and the visceral lymph nodes may resemble blood clots. There is marked petechiation of all serous surfaces, lymph nodes, epicardium and endocardium, renal cortex, and bladder, and edema and congestion of the colon and lungs. The spleen is often large and friable, and there are petechial hemorrhages in the cortex of the kidney. The chronic disease is characterized by cuta-neous ulcers, pneumonia, pericarditis, pleuritis, and arthritis.
Diagnosis
The clinical signs of African swine fever are similar to those of several diseases, including bacterial septicemias such as erysipelas and acute salmonellosis, but the major diagnostic problem is in distinguishing it from classical swine fever (hog cholera). Any febrile disease in swine associated with disseminated hemorrhage (hemorrhagic diathesis) and high mortality should raise suspicion of African swine fever. Diagnosis of chronic infections is problematic as the clinical signs and lesions in affected pigs are highly variable.
Laboratory confirmation is essential, and samples of blood, spleen, kidney, visceral lymph nodes, and tonsils, in particular, should be collected for virus isolation, detection of antigen, or polymerase chain reaction (PCR) for detec-tion of the p72 gene. Virus isolation is done in swine bone marrow or peripheral blood leukocyte cultures, in which hemadsorption can be demonstrated and a cytopathic effect is manifest within a few days after inoculation. After initial isolation, the virus can be adapted to grow in various cell lines, such as Vero cells. Antigen detection is achieved by immunofluorescence staining of tissue smears or frozen
(A) (B)
Figure 8.5 Lesions of acute African swine fever. (A) Subcutaneous hemorrhages in the ear. (B) Splenomegaly. (Courtesy of R. Harutynan, State Veterinary and Epizootic Diagnostic Center, Yerevan, Armenia and W. Laegried, University of Illinois.)
PArt | ii Veterinary and Zoonotic Viruses172
sections, by immunodiffusion using tissue suspensions as the source of antigen, and by enzyme immunoassay.
immunity, Prevention and Control
Both humoral and cellular (including virus-specific CD8 lymphocyte) components contribute to the protective immune response of swine to African swine fever virus. Antibody responses to African swine fever virus have been shown to protect pigs from lethal challenge; however, neu-tralizing antibodies to virion proteins p30, p54, and p72 are not sufficient to confer antibody-mediated protection.
The prevention and control of African swine fever can be complicated by several factors, including the lack of an effective vaccine, the transmission of virus in fresh meat and some cured pork products, the existence of persistent infection in some swine, diagnostic confusion with agents that cause similar disease syndromes such as classical swine fever (hog cholera), and (in some parts of the world) the participation of soft ticks in virus transmission. The presence of the virus in ticks and warthogs in many coun-tries of sub-Saharan Africa makes it difficult, if not impos-sible, to break the sylvatic cycle of the virus. However, domestic swine can be reared in Africa if the management system avoids feeding uncooked waste food scraps and prevents the access of ticks and contact with warthogs, usually by double fencing with a wire mesh perimeter fence extending beneath the ground.
Elsewhere in the world, countries that are free of African swine fever maintain their virus-free status by pro-hibiting the importation of live swine and swine products from infected countries, and by monitoring the destruction of all waste food scraps from ships and aircraft involved in international routings.
If disease does occur in a previously non-infected country, control depends first on early recognition and rapid laboratory diagnosis. The virulent forms of African swine fever cause such dramatic mortality that episodes are brought quickly to the attention of veterinary authori-ties, but the disease caused by less virulent strains that has occurred outside Africa in the past can cause confusion with other diseases and therefore may not be recognized until the virus is well established in the swine population.
Once African swine fever is confirmed in a country that has hitherto been free of disease, prompt action is required to control and then eradicate the infection. All non-African countries that have become infected have elected to attempt eradication. The strategy for eradication involves slaugh-ter of infected swine and swine in contact with them, and disposal of carcasses, preferably by burning. Movement of swine between farms is controlled, and feeding of waste food prohibited. Where soft ticks are known to occur, infested buildings are sprayed with acaricides. Re-stocking of farms is allowed only if sentinel swine do not become
infected. Elimination has been widely successful using this approach, except in Sardinia.
MeMbers of the faMily iridoviridAeThe family Iridoviridae is large and complex; viruses within this family infect arthropods, fish, amphibians, and reptiles. Iridoviruses in the genera Ranavirus, Megalocytivirus, and Lymphocystivirus are the cause of a range of disor-ders in fish, including systemic lethal diseases (genera Ranavirus, Megalocytivirus) and tumor-like skin lesions (Lymphocystivirus). Ranaviruses are considered as a poten-tial cause of the global decline in amphibian populations. Viruses in all three genera are capable of long-term persist-ence in their fish or amphibian hosts, following recovery from acute or inapparent infections.
ProPerties oF iriDoviruses
Members of the Iridoviridae are generally 120–200 nm diameter, but are sometimes even larger DNA viruses, with virions that are similar morphologically to those of the Asfarviridae. Virions exhibit icosahedral symmetry, with a virus core and outer capsid that are separated by an internal lipid membrane (Figure 8.6). Up to 36 proteins are contained in the virions. A viral envelope is present on virions that bud from infected cells, but is not necessary for infectivity. The genomes of iridoviruses consist of a single linear double-stranded DNA molecule that ranges from 140 to 200 kbp in size, and individual viruses encode between approximately 100 and 200 proteins. Termini are different from those of African swine fever virus, being circularly permuted and ter-minally redundant. The family is segregated into two groups on the basis of levels of genomic methylation: a methyltrans-ferase present in the iridoviruses of fish facilitates methyla-tion of up to 20% of cytosine residues in the genomic DNA, similar to that in bacterial genomic DNA.
The family Iridoviridae includes five genera, specifically Iridovirus, Chloriridovirus, Ranavirus, Megalocytivirus, and Lymphocystivirus (Table 8.2). Viruses in this family are of emerging significance, as several are important pathogens in commercial fish production and others cause mortality in captive and wild amphibians. Iridoviruses also cause dis-ease among reptiles, including chelonians (turtles and tor-toises), snakes, and lizards. Interestingly, although viruses in the genera Iridovirus and Chloriridovirus are considered to be viruses of arthropods, these viruses recently have been identified in several species of lizards and scorpions. The genetic similarity of viruses isolated from insects and rep-tiles suggests they may be transmitted to the lizards from their insect prey.
Most information concerning the iridovirus replica-tion cycle is derived from studies of frog virus 3, the type
Chapter | 8 Asfarviridae and Iridoviridae 173
species for the genus Ranavirus (Figure 8.7). The iri-doviruses of vertebrates grow in a wide variety of cells of piscine, amphibian, avian, and mammalian origin at temperatures between 12 and 32°C. Their replication is similar to that of African swine fever virus; however, the viruses do not encode an RNA polymerase, but instead use cellular RNA polymerase II, which their structural proteins modify to favor viral mRNA synthesis. Like African swine fever virus, there is a limited round of initial replication in the nucleus, followed by extensive cytoplasmic replication. Late in infection, vertebrate iri-doviruses produce paracrystalline arrays of virions in the cytoplasm. Infected cells form many microvillus-like pro-jections through which virions bud; however, acquisition of an envelope is not necessary for viral infectivity, and infectious, naked virus particles are released after lysis of infected cells.
rAnAviruses
Since the initial detection of frog virus 3 in the 1960s, an increasing number of related viruses have been associ-ated with diseases in amphibians and fish in their fresh-water environments. Frog virus 3 was initially isolated from leopard frogs in the eastern United States, during an
Lymphocystivirus Lymphocystis disease virus 1 (LCDV-1) and LCDV-2
Unclassified White sturgeon iridovirus
PArt | ii Veterinary and Zoonotic Viruses174
investigation into causes of naturally occurring renal carci-nomas that was later traced to infection with an oncogenic alloherpesvirus, ranid herpesvirus 1. Although ranavi-ruses such as frog virus 3 were initially considered to be relatively benign, by the mid-1980s it was increasingly apparent that ranaviruses were associated with severe and widespread disease epizootics amongst wild amphibian populations in North America, Europe, and Asia. Infected tadpoles, which are most susceptible, and frogs may exhibit localized cutaneous hemorrhage and/or ulceration or severe systemic disease with edema, hemorrhage, and necrosis in numerous organs. Subclinical infections occur in apparently normal wild and captive populations of frogs, in which ranavirus is detected in kidney tissues, including macrophages, which serve as a site for virus persistence.
Ambystoma tigrinum virus is a ranavirus that causes mortality in both larval and adult salamanders in west-ern North America from late summer to early autumn. Mortality that can exceed 90% occurs within 7–14 days of exposure to virus in the water or by direct contact with diseased salamanders. Diseased animals may exhibit any combination of necrosis and hemorrhage within the spleen, liver, kidney, and gastrointestinal tract, sloughing of the skin, development of skin polyps, and discharge of inflam-matory exudate from the vent. Environmental temperature plays an important role in the pathogenesis of infection,
as most salamanders infected at 26°C survive, whereas at 18°C most die of the infection. The role of vertical trans-mission from infected adults to eggs is unknown, and the virus does not appear to have an alternate reservoir host.
Ranavirus-associated diseases of fish were first reported in Australia in 1986, initially amongst lake populations of redfin perch that had a systemic disease characterized by extensive necrosis of the liver, pancreas, and hemopoietic cells of the kidney and spleen. This disease, termed “epizootic hemopoietic necrosis,” was later identified in farmed popu-lations of rainbow trout in the same water systems as the affected redfin perch. The causative virus was transmit-ted experimentally to seven additional fish species found in Australia. Fingerling and juvenile fish are commonly affected; however, when epizootic hematopoietic necro-sis virus is newly introduced, adults are also susceptible. In general, the ranaviruses of fish can be readily detected by isolation from internal organs (kidney, spleen, liver) on a range of cell lines, usually of fish origin, which are incubated at 20–25°C. The virus can be distinguished from other ranaviruses by DNA-based diagnostic procedures.
Additional ranaviruses have recently been detected dur-ing disease episodes among cultured freshwater populations of silurid and ictalurid catfish in Europe and Atlantic cod fry in a hatchery in Denmark. Largemouth bass virus (syn. Santee–Cooper virus) is a ranavirus that has been associated
Enveloped virion
DNAcore
Uncoating via receptor-mediated endocytosis
Second stage viralDNA replication.
Concatemer formation,DNA methylation
First stage viralDNA replicationsynthesis ofgenome-size DNAto twice genome-size DNA
with substantial seasonal loss of wild adult largemouth bass in lakes in the United States. The virus affects a variety of internal tissues, including the swim bladder, which becomes reddened and enlarged and contains a yellow exudate. Involvement of the swim bladder results in moribund fish that float to the surface, which is often the first indication of disease in wild fish. In experimental studies, the virus caused only low-grade mortality in largemouth bass, which suggests that the epizootic mortality that occurs during disease out-breaks among wild fish is probably due to additional con-tributing factors. As in amphibians, ranaviruses can often be isolated from asymptomatic fish, a feature that contributes to the unintentional dispersal of virus with the international trade of live amphibians and fish. Transmission of ranavi-ruses between amphibian and fish has been demonstrated in both natural and experimental settings.
Ranaviruses are increasingly recognized as the cause of disease among wild and captive reptiles. Ranavirus infec-tions have been identified among chelonians (turtles and tortoises), lizards, and snakes on several continents, some-times in association with disease syndromes similar to those encountered in ranavirus-infected amphibians.
MegAloCytiviruses
The emerging and significant impact of megalocytiviruses on the commercial production of both food and ornamen-tal fish has become increasingly apparent since their initial detection in 1990 among cultured populations of red sea bream in Japan. Over 30 species of marine and freshwater fish from Japan, the South China Sea, and several Southeast Asian countries are now documented as potential hosts of megalocytiviruses. The viruses all share significant homol-ogy, with 97% or greater identity at the deduced amino acid level for the major capsid protein. The entire genome sequence has been determined for at least three megalocyti-viruses, namely infectious spleen and kidney necrosis virus, rock bream iridovirus, and orange spotted grouper irido- virus. Mortality of up to 100% has been described during epizootics in captive fish populations, and after experimental infection. Signs exhibited by diseased fish include lethargy, severe anemia, and branchial hemorrhages. At necropsy, the spleen may be greatly enlarged. On microscopic evaluation, numerous large, basophilic, “cytomegalic” cells that have a subendothelial location are typically present in internal organs such as spleen, kidney, intestine, eye, pancreas, liver, heart, gill, brain, and intestine; these characteristic cells are reflected in the genus name for these viruses. The enlarged cells contain abundant numbers of developing and complete virions. In contrast to ranaviruses, the megalocytiviruses are often difficult to isolate in cell culture, and thus diag-nosis has traditionally been reliant on histologic evaluation followed by confirmation with electron microscopy. DNA-based diagnostic methods such as PCR are now routinely
used to detect and distinguish megalocytiviruses in captive and wild fish populations.
Control methods include the use of pathogen-free fish, improved sanitation on fish farms and husbandry practices that minimize stress (lower fish densities, good water qual-ity, etc.). A formalin-killed virus vaccine administered by injection has proven efficacious in the control of the red sea bream iridovirus in Japan. The megalocytiviruses are horizontally transmitted among fish in the water, and there is no evidence to date for vertical transmission from adults to progeny. The broad host range and detection of megalo-cytiviruses in many ornamental fish species shipped from enzootic areas create a major concern for the control of this important group of fish pathogens.
lyMPhoCystiviruses
Lymphocystis is a benign and self-limiting disease described in a broad range of freshwater and marine fish species. The condition is caused by a group of iridoviruses that infect and then transform fibroblasts of the skin and gills and internal connective tissues, resulting in remark-able hypertrophy of the affected cells (Figure 8.8). These cells, termed “lymphocysts,” appear as raised pearl-like lesions and can be observed readily with the naked eye. Infections occur in over 125 species and 34 families of fish from warm, temperate, and cold, and marine or fresh- water environments. Lymphocysts, which may reach 100,000 times the normal cell size, are a result of virus-mediated arrest of cell division but not cell growth, which leads to the formation of megalocytes. Lymphocysts pos-sess a distinct hyaline-like capsule, an enlarged nucleus, and bizarre and segmented cytoplasmic inclusions that contain developing virions. The characteristic histologic appearance of lymphocysts is pathognomonic for lym-phocystis disease, although electron microscopy is often used to confirm the presence of typical iridovirus virions.
Lymphocystis disease virus 1 is associated with infec-tions in two marine fish species, flounder and plaice, whereas lymphocystis disease virus 2 occurs in a third marine fish, dab. There are many additional and related viruses associated with lymphocystis that occur in other fish species in marine and freshwater habitats, but these have not been fully characterized. Infections with lym-phocystis disease virus are seldom fatal and most often fish recover by sloughing external lymphocysts. The most important impact of the virus is the loss of commercial value as a result of cosmetic effects that occur in cultured or wild-caught fish sold as food. In addition to the cos-metic effects with ornamental fish, heavy infections in the oral region may inhibit feeding, and the effects of the viral infections may result in entry points for secondary patho-gens. Transmission from fish to fish is probably via contact with virus released from ruptured lymphocysts that spreads
PArt | ii Veterinary and Zoonotic Viruses176
the virus among crowded fish populations. Separation and quarantine of infected fish until lymphocysts resolve are the means to reduce infections in captive fish populations. Genetic analyses of Japanese flounder with lymphocystis suggest a genetic basis for susceptibility to the virus, a find-ing that may eventually aid in selective breeding to reduce the prevalence of disease.
other iriDoviruses oF Fish
Erythrocytic necrosis virus is a group of unassigned irido-viruses that share morphologic features with members of the family Iridoviridae. Virions are present in the cyto-plasm of immature hemopoietic cells or mature erythro-cytes of a range of marine fish (e.g., herring and cod) and some species of salmonid fish found in the North Pacific and North Atlantic oceans. Heavy infections result in sig-nificant anemia and losses among both wild and farmed populations of fish. Infected erythrocytes contain a distinct circular cytoplasmic inclusion(s) as seen in stained blood smears. Viral infections in erythrocytes are confirmed by the presence of iridovirus-like virions in the cytoplasm by electron microscopy. Viruses associated with erythrocytic necrosis have also been observed in reptiles and amphib-ians. The viruses found in fish have not been isolated, pre-sumably because of the absence of suitable cell lines of hemopoietic origin. Fish erythrocytic necrosis virus can be transmitted experimentally by intraperitoneal injections with infected erythrocytes. Many features of the disease remain uncharacterized.
The white sturgeon iridovirus, a currently unassigned virus in the family Iridoviridae, was first recognized as the cause of epizootic mortality of farmed juvenile stur-geon in the 1980s in California. Infection with this virus results in destruction of the epithelium of the skin and gills,
compromising both respiration and osmotic balance. White sturgeon iridovirus disease is considered the most prob-lematic viral disease of white sturgeon cultured for meat or caviar. The virus has been identified in wild and cap-tive populations of white sturgeon throughout the Pacific Northwest of North America. It has also been moved beyond its original range through the export of live white stur-geon. Infections are detected by histologic examination that reveals the presence of characteristic enlarged amphophilic to basophilic-staining cells in the epithelium, often associ-ated with necrosis of surrounding cells. Virions can be iden-tified by electron microscopic examination of enlarged cells. More recently, specific PCR tests have been developed to assist in confirming infections with the white sturgeon iri-dovirus. Virus transmission occurs in contaminated water, and there is strong evidence of vertical transmission of the virus with gametes from infected adult fish. Separation of year classes of sturgeon and segregation of infected lots of juvenile fish are the principal control methods. Infections and significant losses of several different species of juvenile sturgeon with viruses related to the white sturgeon irido- virus have now been reported in wild and captive populations of shovelnose, pallid, and lake sturgeon in the United States, and Italian and Russian sturgeon in Europe.
iriDoviruses oF Mollusks
Iridovirus or iridovirus-like agents associated with mor-tality of larval and adult oysters have been described in both Europe and North America. Catastrophic losses of the Portuguese oyster cultured along the Atlantic coast of France during the early 1970s were attributed to iridovirus infection that caused severe necrosis of the gill epithe-lium, or that infected hemocytes. A subsequent outbreak of the hemocytic disease occurred among Pacific oysters in
(A) (B)
Figure 8.8 (A) Lymphocystis in a walleye. (B) Histological appearance of lymphocystis, depicting cellular hypertrophy. (Courtesy of P. Brower, Cornell University and R. Hedrick, University of California.)
Chapter | 8 Asfarviridae and Iridoviridae 177
France in 1977, suggesting that this introduced oyster spe-cies was the potential source of the virus that infected the resident oyster populations. Oyster velar virus disease was first described in the late 1970s as the cause of mortality—that approached 100%—among larval stages of the Pacific oyster in hatcheries in the state of Washington. The target tissue of this virus is the velum, a ciliated structure respon-sible for locomotion and feeding of the larvae. Infection
results in the formation of blisters and sloughing of the ciliated epithelium and then death. Virions in infected cells share morphologic properties with those in affected adult oysters in France, although they are slightly smaller in size (228 nm diameter). Control measures for iridovirus infec-tions in mollusks rely upon early detection and destruction of infected groups, followed by vigorous disinfection, par-ticularly in hatchery settings.
