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equine encephalomyelitis (Eastern)

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Datasheet

equine encephalomyelitis (Eastern)

Summary

  • Last modified
  • 29 January 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • equine encephalomyelitis (Eastern)
  • Overview
  • Eastern equine encephalitis is caused by a member of the genus Alphavirus of the family Togaviridae. The disease was first recorded in the USA, possibly as early as 1831 in Massachusetts, and was first i...

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Identity

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Preferred Scientific Name

  • equine encephalomyelitis (Eastern)

International Common Names

  • English: arboviral encephalitis in horses; eastern equine encephalitis; eastern equine encephalitis virus-associated haemorrhagic enterocolitis; eastern equine encephalomyelitis; eastern equine encephalomyelitis, eee, in horses and cattle; eastern equine encephalomyelitis, eee, in pigs; epizootic equine encephalomyelitis; sleeping sickness; swine viral hepatitis

English acronym

  • EEE

Overview

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Eastern equine encephalitis is caused by a member of the genus Alphavirus of the family Togaviridae. The disease was first recorded in the USA, possibly as early as 1831 in Massachusetts, and was first isolated after the outbreaks in Delaware, Maryland, and Virginia in 1933 and 1934 (Hanson, 1957; Ten Broeck, 1938). Epidemics of eastern equine encephalitis are recorded at periodic intervals (Calisher, 1994). It is mainly restricted to North, Central and South America where the virus cycles between birds and ornithophilic mosquitoes. Eastern equine encephalitis was first recognized in horses on the eastern coast of the US (Giltner and Shahan, 1933; TenBroeck and Merrill, 1933; Elvinger and Baldwin, 1999). It has also been diagnosed and reported in cattle, dogs, goats, swine and a variety of wild mammalian species (Pursell et al., 1972, 1976, 1983; Bigler et al., 1976; McGee et al., 1992). Wild and domestic birds are susceptible to the virus (Scott and Weaver, 1989), although generally do not suffer disease, apart from pheasants and sparrows, which develop encephalitis and may die.

The alphavirus causes flu-like disease in horses and humans from mid-summer to late autumn when infection builds up in the bird populations to a level which results in spillover to other species of mosquito which feed on both birds and mammals on the margins of swampy areas. Many infections in horses and humans are inapparent, resulting in viraemia lasting up to a week, followed by long-lasting immunity (Thomson, 1994). During 1996-1997 there were 19 cases (5 fatal) of eastern equine encephalitis in humans in 8 states of the USA. Enzootic arboviral activity was reported in 18 states including 274 cases of arboviral encephalitis in horses and several cases of epizootics or sporadic clinical cases of haemorrhagic enterocolitis associated with eastern equine encephalitis virus (Anon, 1998).

Naturally occurring eastern equine encephalitis in swine was first reported in 1972 (Pursell et al., 1972) in southern Georgia when 160 of 200 pigs in a herd died in an outbreak. Exposure of domestic and feral swine to the agent had been described earlier in serological surveys in Georgia, Massachusetts and Wisconsin (Karstad and Hanson, 1958, 1959; Feemster et al., 1958; Elvinger and Baldwin, 1999). The virus has also been implicated as a primary cause of swine viral hepatitis (Marcato and Perillo, 2000).

This disease is on the list of diseases notifiable to the World Organisation for Animal Health (OIE).

Host Animals

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Animal nameContextLife stageSystem
Bos indicus (zebu)Domesticated host
Bos taurus (cattle)Domesticated host
Capra hircus (goats)Domesticated host
Coturnix japonica (Japanese quail)Wild host
Dromaius novaehollandiaeDomesticated host, Wild host
Equus caballus (horses)Domesticated host
Gallus gallus domesticus (chickens)Domesticated host
Meleagris gallopavo (turkey)Domesticated host
Sus scrofa (pigs)Domesticated host

Hosts/Species Affected

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Eastern equine encephalitis is specifically a disease of horses, but has a wide host range, including humans, rodents, reptiles, amphibians, monkeys, dogs, cats, foxes, skunks, cattle, pigs, birds and mosquitoes. Clearly, the disease requires appropriate climatic conditions for mosquitoes to breed to maintain its life cycle. Clinical disease is more common in young animals with eastern equine encephalitis frequently being fatal in horses. Between November 1993 and August 1995, 66 free-ranging Florida black bears (Ursus americanus) were tested for antibodies to eastern equine encephalitis and 7 (11%) were found positive (Dunbar et al., 1998). Antibodies to eastern equine encephalitis were found in neotropical bats in Guatemala in 1983-1994 (Ubico and McLean, 1995).

Evidence for human and equine infection is plentiful (Moore, 1991; Krenick, 1991; Crans, 1991, 1992; Hlady, 1993; Ross and Kaneene, 1995; Fowler, 1997; Silva et al., 1999; Romano-Lieber and Iversson, 2000).

Many species of mosquitoes have also been implicated in the life cycle of disease transmission, including Culiseta melanura, Aedes vexans, Aedes canadensis, Aedes sollicitans, Aedes taeniorhynchus, Aedes triseriauts, Coquillettidiea perturbans, Anopheles bradleyi, Anopheles quadrimaculatus, Anopheles puncitipennis, Culex pipiens, Culex salinarius, Culex restuans, Culex territans and Psorophora columbiae (Crans, 1992; Reinert, 1995). In Michigan in 1991, 55 equids (horses, ponies and a zebra) were affected by eastern equine encephalitis. In a retrospective case control study of the outbreak, annual vaccination against eastern equine encephalitis and the use of insect repellent methods decreased the risk of eastern equine encephalitis. Woodlands and swamps on the farm were shown to increase the risk, providing a breeding ground for mosquitoes (Ross and Kaneene, 1995.).

Avian infection in wild and domestic species is very common, and birds are believed to be the basis for overwintering when conditions are too harsh for mosquitoes to survive. Serum neutralizing antibodies were found to eastern equine encephalitis virus in 4 of 121 (3%) wild bird sera in South Carolina in 1995 (Durden et al., 1997), suggesting some wild birds may be reservoir hosts for encephalitis viruses. Encephalitis was found in 0.48% of 12,423 samples of avian sera or viscera representing 304 species from 193 genera and 40 families collected in 198 localities in the Amazonian states of Acre, Amazonas, Mato Grosso and Pará, Brazil (Dégallier et al., 1992). In 1980-83 in New Jersey, reservoirs of eastern equine encephalitis were found in blue jay (Cyanocitta cristata), wood thrush (Hylocichla mustelina), tufted titmouse (Parus bicolor), Carolina chickadee (Parus carolinensis), ovenbird (Seiurus aurocapillus), northern cardinal (Cardinalis cardinalis) and hairy woodpecker (Picoides villosus) (Rupp and Hajek, 1994.).

The disease also affects common avian species. Eastern equine encephalitis viraemias are more intense and longer lasting in starlings than in American robins (Turdus migratorius) and other birds (Agelaius phoeniceus, Quiscalus quiscala, Melospiza georgiana, Melospiza melodia, Molothrus ater, Cardinalis cardinalus, Zenaida macroura and Columba livia). Starlings frequently die as their viraemia begins to wane; other birds generally survive. Various Aedes (A. aegypti, A. albopictus, A. triseriatus) as well as Culiseta melanura mosquitoes can acquire eastern equine encephalitis viral infection from infected starlings under laboratory conditions. Starlings are not originally native where eastern equine encephalitis is enzootic, but a starling can infect about 3 times as many mosquitoes as can a robin (Komar et al., 1999).

High mortality has been reported in domestic fowl, pheasants, chukars, emus and quail, most often caused by eastern equine encephalitis in the eastern coastal States of the USA. After introduction by mosquitoes the horizontal transmission within flocks is primarily by feather picking and cannibalism (OIE, 2000), or in turkeys, pheasants and ostriches by the faecal-oral route (Brown et al., 1993). Haemagglutination inhibition and neutralizing antibodies to eastern equine encephalitis were detected in emus imported to Palm Beach Florida, November 1993 to January 1995. Emus imported from California, Louisiana and Texas had evidence of naturally acquired antibodies to eastern equine encephalitis. This observation underscores the potential threat of arboviral introduction by infected vertebrates and emphasizes the importance of instituting quarantine procedures to regulate the transport of hosts that may be infected with arboviral agents (Day and Stark, 1998). Ratities are not generally regarded as amplifiers of eastern equine encephalitis viruses in endemic areas and are considered accidental hosts (Veazey et al., 1994).

However, glossy ibises (Plegadis falcinellus) and snowy egrets (Egretta thula) have been shown to be involved in the maintenance of the epizootiology of eastern equine encephalitis (McLean et al, 1994 and 1995) as under experimental inoculation, 93% of the birds became viraemic and all developed neutralizing antibodies. Aedes albopictus became infected with eastern equine encephalitis virus after feeding on viraemic snowy egrets and demonstrated a low threshold of infection.

In 1991-92, 1247 serum samples were collected from domestic pigs in GA USA and 376 serum samples were collected from feral pigs on Ossabaw Island, GA, and were tested for neutralizing antibodies to eastern equine encephalitis. 16 (1.7%) of 931 samples from domestic pigs in 1991 had serum neutralization titres of 1:4 or 1:8 and 16 (5.1%) of 316 samples were positive with similar titres in 1992. Evidence of eastern equine encephalitis virus exposure in pigs was found in 12 of the 28 counties in southern Georgia tested. 62 (16.5%) of 276 feral pig samples were positive for eastern equine encephalitis antibodies. Results indicated domestic and feral pigs are exposed to eastern equine encephalitis virus in GA. It is suggested that eastern equine encephalitis infection is under-diagnosed in pigs even though it causes significant financial losses in affected herds (Elvinger et al., 1996).

Systems Affected

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nervous system diseases of pigs

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

AfghanistanNo information availableOIE, 2009
ArmeniaDisease not reportedOIE, 2009
AzerbaijanDisease never reportedOIE, 2009
BahrainDisease never reportedOIE, 2009
BangladeshDisease never reportedOIE, 2009
BhutanDisease never reportedOIE, 2009
CambodiaNo information availableOIE, 2009
ChinaDisease never reportedOIE, 2009
-Hong KongNo information availableOIE, 2009
IndiaDisease never reportedNULLAlstad and Pearson, 1995; OIE, 2009
IndonesiaDisease never reportedOIE, 2009
IranDisease never reportedOIE, 2009
IraqDisease never reportedOIE, 2009
IsraelDisease never reportedOIE, 2009
JapanDisease never reportedOIE, 2009
JordanDisease never reportedOIE, 2009
KazakhstanDisease never reportedOIE, 2009
Korea, Republic ofDisease never reportedOIE, 2009
KuwaitDisease not reportedOIE, 2009
KyrgyzstanDisease never reportedOIE, 2009
LaosDisease never reportedOIE, 2009
LebanonDisease never reportedOIE, 2009
MalaysiaDisease never reportedOIE, 2009
MongoliaNo information availableOIE, 2009
MyanmarDisease never reportedOIE, 2009
NepalNo information availableOIE, 2009
OmanDisease not reportedOIE, 2009
PakistanDisease not reportedOIE, 2009
PhilippinesDisease never reportedOIE, 2009
QatarDisease never reportedOIE, 2009
Saudi ArabiaNo information availableOIE, 2009
SingaporeDisease never reportedOIE, 2009
Sri LankaDisease never reportedOIE, 2009
SyriaNo information availableOIE, 2009
TajikistanDisease not reportedOIE, 2009
ThailandDisease never reportedOIE, 2009
TurkeyNo information availableOIE, 2009
United Arab EmiratesDisease never reportedOIE, 2009
VietnamNo information availableOIE, 2009
YemenNo information availableOIE, 2009

Africa

AlgeriaNo information availableOIE, 2009
AngolaNo information availableOIE, 2009
BeninNo information availableOIE, 2009
BotswanaDisease never reportedOIE, 2009
Burkina FasoNo information availableOIE, 2009
ChadNo information availableOIE, 2009
CongoNo information availableOIE, 2009
DjiboutiDisease never reportedOIE, 2009
EgyptNo information availableOIE, 2009
EritreaNo information availableOIE, 2009
EthiopiaNo information availableOIE, 2009
GabonNo information availableOIE, 2009
GambiaNo information availableOIE, 2009
GhanaNo information availableOIE, 2009
GuineaNo information availableOIE, 2009
Guinea-BissauNo information availableOIE, 2009
KenyaDisease never reportedOIE, 2009
LesothoDisease never reportedOIE, 2009
MadagascarDisease never reportedOIE, 2009
MalawiDisease never reportedOIE, 2009
MaliNo information availableOIE, 2009
MauritiusDisease never reportedOIE, 2009
MoroccoDisease never reportedOIE, 2009
MozambiqueDisease not reportedOIE, 2009
NamibiaDisease never reportedOIE, 2009
NigeriaDisease never reportedOIE, 2009
RwandaNo information availableOIE, 2009
SenegalNo information availableOIE, 2009
South AfricaDisease never reportedOIE, 2009
SudanDisease never reportedOIE, 2009
SwazilandDisease not reportedOIE, 2009
TanzaniaNo information availableOIE, 2009
TogoNo information availableOIE, 2009
TunisiaDisease never reportedOIE, 2009
UgandaNo information availableOIE, 2009
ZambiaDisease never reportedOIE, 2009
ZimbabweDisease never reportedOIE, 2009

North America

CanadaRestricted distributionOIE, 2009
GreenlandDisease never reportedOIE, 2009
MexicoDisease not reportedNULLAguirre et al., 1992; OIE, 2009
USARestricted distributionOIE, 2009
-FloridaPresentTurell, 1993; Gamble et al., 1994; Morris et al., 1994; Day and Stark, 1996; Day and Stark, 1996; Dunbar et al., 1998
-GeorgiaMoore, 1991; Brown et al., 1993; Elvinger et al., 1994; Elvinger et al., 1996; Elvinger et al., 1996; Elvinger and Baldwin, 1999
-KentuckyPresentPoonacha et al., 1998
-LouisianaPresentTully et al., 1992; Hugh-Jones and Samui, 1993; Veazey et al., 1994
-MarylandPresentMoore, 1991; Olsen et al., 1997
-MassachusettsPresentMoore, 1991; Villari et al., 1994; Alstad and Pearson, 1995; Moncayo et al., 2000
-MichiganPresentAlstad and Pearson, 1995; Ross and Kaneene, 1995
-MississippiPresentCalisher et al., 1990; McGee et al., 1992; Hugh-Jones and Samui, 1993; Goddard and Currier, 1995
-New JerseyPresentCrans, 1991; Moore, 1991; Crans, 1994; Reinert, 1995; Crans et al., 1996
-New YorkPresentKrenick, 1991; Moore, 1991; Howard et al., 1996
-North CarolinaPresentMoore, 1991; Ficken et al., 1993; Wages et al., 1993; Barnes, 1994
-OhioPresentNasci et al., 1993
-South CarolinaPresentMoore, 1991; Letson et al., 1993; Wright, 1993; Fowler, 1997

Central America and Caribbean

BelizeDisease not reportedOIE, 2009
Costa RicaNo information availableOIE, 2009
CubaDisease not reportedOIE, 2009
Dominican RepublicDisease not reportedOIE, 2009
El SalvadorDisease not reportedOIE, 2009
GuadeloupeNo information availableOIE, 2009
GuatemalaDisease not reported2000Ubico and McLean, 1995; OIE, 2009
HaitiDisease never reportedOIE, 2009
HondurasDisease never reportedOIE, 2009
JamaicaNo information availableOIE, 2009
MartiniqueDisease never reportedOIE, 2009
NicaraguaDisease never reportedOIE, 2009
PanamaNo information availableNULLAndreadis et al., 1998; OIE, 2009

South America

ArgentinaDisease not reported1988Sabattini et al., 1991; OIE, 2009
BoliviaAbsent, reported but not confirmedOIE, 2009
BrazilDisease not reportedOIE, 2009
-AmazonasWidespreadVasconcelos et al., 1991
-ParanaWidespreadFernández et al., 2000
-Sao PauloPresentFerreira et al., 1994; Romano-Lieber and Iversson, 2000
ChileDisease never reportedOIE, 2009
ColombiaDisease not reportedOIE, 2009
EcuadorDisease never reportedOIE, 2009
French GuianaDisease not reportedOIE, 2009
PeruDisease never reportedOIE, 2009
UruguayDisease not reportedOIE, 2009
VenezuelaDisease not reportedOIE, 2009

Europe

AlbaniaNo information availableOIE, 2009
AustriaDisease not reportedOIE, 2009
BelarusDisease not reportedOIE, 2009
BelgiumDisease not reportedOIE, 2009
BulgariaDisease never reportedOIE, 2009
CroatiaDisease never reportedOIE, 2009
CyprusDisease never reportedOIE, 2009
Czech RepublicDisease never reportedOIE, 2009
DenmarkDisease never reportedOIE, 2009
EstoniaDisease never reportedOIE, 2009
FinlandDisease never reportedOIE, 2009
FranceDisease never reportedOIE, 2009
GermanyDisease never reportedOIE, 2009
GreeceDisease not reportedOIE, 2009
HungaryDisease never reportedOIE, 2009
IcelandDisease never reportedOIE, 2009
IrelandDisease never reportedOIE, 2009
ItalyNo information availableOIE, 2009
LatviaDisease never reportedOIE, 2009
LiechtensteinDisease not reportedOIE, 2009
LithuaniaDisease never reportedOIE, 2009
LuxembourgDisease never reportedOIE, 2009
MacedoniaDisease never reportedOIE, 2009
MaltaDisease never reportedOIE, 2009
MontenegroDisease never reportedOIE, 2009
NetherlandsDisease never reportedOIE, 2009
NorwayDisease never reportedOIE, 2009
PolandNo information availableOIE, 2009
PortugalDisease never reportedOIE, 2009
RomaniaDisease never reportedOIE, 2009
Russian FederationDisease never reportedOIE, 2009
SerbiaNo information availableOIE, 2009
SlovakiaDisease not reportedOIE, 2009
SloveniaDisease never reportedOIE, 2009
SpainDisease never reportedOIE, 2009
SwedenDisease never reportedOIE, 2009
SwitzerlandDisease not reportedOIE, 2009
UKDisease not reportedOIE, 2009
UkraineDisease not reportedOIE, 2009

Oceania

AustraliaDisease never reportedOIE, 2009
French PolynesiaDisease never reportedOIE, 2009
New CaledoniaDisease never reportedOIE, 2009
New ZealandDisease never reportedOIE, 2009

Pathology

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No gross lesions are found post-mortem. Four of 9 unweaned pigs inoculated with the eastern equine encephalitis isolant in cell culture fluid or 10% brain suspension developed CNS signs similar to those seen in a naturally infected pig in southern Georgia, USA, in 1972. Brain lesions in the pigs were characterized by neutrophils in perivascular cuffs and in necrotic areas (Pursell et al., 1972). Histological features in an experimentally infected calf conformed with those described for eastern equine encephalitis in the horse. (Pursell et al., 1976.). Histological lesions in birds include inflammation and haemorrhage or necrosis of visceral organs, which differs from the central nervous system disease seen in mammals (Dein et al., 1986; Tully et al., 1992; Guy et al., 1994).

Diagnosis

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Speculative diagnosis of alphavirus infection may be based on signs of encephalitis associated with past history and the appropriate seasonal conditions. Laboratory diagnosis via virus isolation or serology is necessary. Virus isolation is carried out using blood or CNS tissue, the latter being preferable due to the uncertainty of viraemia when signs of encephalitis are noted. Cell culture, chick embryos and suckling mice may also be used for isolation. The virus is subsequently identified via virus neutralization, haemagglutination inhibition (HI), immunofluorescence, ELISA or polymerase chain reaction (PCR). Serological diagnosis is best performed using paired serum samples in an HI test. Results must be interpreted with care as animals with previous alphavirus infection may respond to later infections with broadly cross-reactive antibodies. Previous vaccinations can also affect serology results (Biberstein and Zee, 1990).

Diagnosis of eastern equine encephalitis in avian species is relatively difficult compared with other species as there are no characteristic histological lesions in the avian that would serve to distinguish between eastern equine encephalitis infections and those of Newcastle disease or highly pathogenic avian influenza virus, for example. However, virus isolation and/or HI are still used for definitive diagnosis in birds. An immunohistochemistry (IHC) assay has been developed for detection of eastern equine encephalitis virus antigen in formalin-fixed, paraffin-embedded tissues of horses, pheasants and a partridge. The IHC assay was based on standard streptavidin-biotin-binding technology, using a commercial kit (Histostatin-SP Kit, Zymed Laboratories, CA) and a mono-specific polyclonal primary antibody preparation derived from murine ascitic fluid. It was shown to be a rapid, effective test for confirming diagnosis (Patterson et al., 1996), including isolation in two cases of eastern equine encephalitis in ring-necked pheasants (Williams et al., 2000).

Eastern equine encephalitis virus was detected by indirect immunofluorescence technique in a crossbred cow, aged 1.5 years, which had been recumbent for 2 days. The cow could stand with assistance but circled to the left when walking (McGee et al., 1992).

Reverse-transcription-PCR (RT-PCR) tests have been developed for specific detection of equine encephalitis viruses and in New Jersey in 1994, the efficacy of the RT-PCR was evaluated as an early warning tool for the presence of eastern equine encephalitis virus activity in wild populations of Culiseta melanura. The technique proved to be efficient as a warning tool and detected viral activity earlier at sites from which horse and emu deaths were later reported (Mahmood and Crans, 1995; Linssen et al., 1999). Arboviral RNA can also be directly identified by RT-PCR in field collected pools of infected mosquitoes (Huang et al., 2001). Linssen et al. (2000) developed an eastern equine encephalitis-specific RT-PCR to amplify a 464-base pair region of the E2 gene exclusively from 10 different eastern equine encephalitis strains from South and North America, with a sensitivity of about 3000 RNA molecules. Specificity was confirmed in a subsequent nested PCR by the amplification of a 262-bp fragment which increased the sensitivity to approximately 30 RNA molecules.

A PCR has been shown to detect eastern equine encephalitis virus RNA from Culex pipiens pipiens, Culex pipiens form molestrus and Aedes aegypti in the laboratory and from field collected samples of Culex melanura and from the 1990 vector surveillance programme in New Jersey, USA. Confirmation that PCR products were of viral origin was obtained by hybridization to viral RNA and by internal restriction endonuclease digestion (Monroy and Webb, 1993).

Johnson et al. (2000) developed a monoclonal antibody based capture ELISA for detecting an anti-arboviral immunoglobulin G for eastern equine encephalitis. The virus was detected in infected formalin-fixed horse and emu tissues and infected chicken embryo fibroblasts. Results of in-situ hybridization using a digoxigenin-labelled 40-base DNA probe complementary to a conserved region of the virus RNA compared favourably with results of both virus isolation and serum neutralization tests. Gregory et al. (1996) had previously concluded that an ELISA technique may be useful for diagnosing infection in various animals species, especially when fresh tissue are not available for analysis. An IgM test can be used to identify recent infection.

Indirect immunofluorescence and double-antibody ELISA detected eastern equine encephalitis virus within 6-8 h of inoculation of the viruses maintained in Vero and XL (Xenopus laevis)-2 cell cultures; better results were obtained with XL-2 cultures. (Pelegrino et al., 1993).

In a human case, eastern equine encephalitis virus was recovered directly from a 14-year old American patient’s CSF in A549 and MRC-5 cell cultures (Sotomayor and Josephson, 1999).

List of Symptoms/Signs

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SignLife StagesType
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed Pigs:Piglet Diagnosis
Digestive Signs / Diarrhoea Sign
Digestive Signs / Dysphagia, difficulty swallowing Sign
Digestive Signs / Excessive salivation, frothing at the mouth, ptyalism Sign
Digestive Signs / Grinding teeth, bruxism, odontoprisis Sign
Digestive Signs / Tongue weakness, paresis, paralysis Sign
Digestive Signs / Vomiting or regurgitation, emesis Sign
General Signs / Ataxia, incoordination, staggering, falling Cattle & Buffaloes:All Stages,Other:All Stages Diagnosis
General Signs / Decreased, absent thirst, hypodipsia, adipsia Sign
General Signs / Dysmetria, hypermetria, hypometria Sign
General Signs / Fever, pyrexia, hyperthermia Sign
General Signs / Forelimb weakness, paresis, paralysis front leg Sign
General Signs / Generalized weakness, paresis, paralysis Cattle & Buffaloes:All Stages,Pigs:Piglet Diagnosis
General Signs / Head, face, ears, jaw weakness, droop, paresis, paralysis Sign
General Signs / Inability to stand, downer, prostration Sign
General Signs / Increased mortality in flocks of birds Poultry:All Stages Sign
General Signs / Opisthotonus Sign
General Signs / Reluctant to move, refusal to move Sign
General Signs / Tetraparesis, weakness, paralysis all four limbs Sign
General Signs / Trembling, shivering, fasciculations, chilling Sign
General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift Sign
General Signs / Weight loss Sign
Nervous Signs / Abnormal behavior, aggression, changing habits Sign
Nervous Signs / Circling Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:Piglet Diagnosis
Nervous Signs / Coma, stupor Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:Piglet Diagnosis
Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:Piglet Diagnosis
Nervous Signs / Excitement, delirium, mania Sign
Nervous Signs / Head pressing Sign
Nervous Signs / Head tilt Sign
Nervous Signs / Hyperesthesia, irritable, hyperactive Sign
Nervous Signs / Propulsion, aimless wandering Sign
Nervous Signs / Seizures or syncope, convulsions, fits, collapse Sign
Nervous Signs / Tremor Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:Piglet Diagnosis
Ophthalmology Signs / Blindness Sign
Ophthalmology Signs / Nystagmus Sign
Reproductive Signs / Agalactia, decreased, absent milk production Cattle & Buffaloes:Cow Diagnosis
Reproductive Signs / Decreased, dropping, egg production Poultry:Mature female Diagnosis
Skin / Integumentary Signs / Alopecia, thinning, shedding, easily epilated, loss of, hair Sign
Skin / Integumentary Signs / Pruritus, itching skin Sign

Disease Course

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Alphavirus infections can range from inapparent to severe fatal disease. Infection typically occurs following the bite of an infected insect, with primary virus replication occurring locally and in adjacent lymph nodes. The primary viraemia, which persists for several days, serves to infect various target organs throughout the body; it is followed by secondary viraemia. The encephalitis virus may gain entry into the central nervous system if the viraemia is of sufficient level. The virus appears to enter the nervous tissue via exposed nerve endings or neuromuscular junctions, resulting in a necrotizing encephalitis in one to three weeks. Lesions often develop throughout the grey matter of the brain and include neuronal degeneration, vascular cuffing, infiltration of neutrophils and lymphocytes, microglial proliferation and haemorrhage. Macroscopic lesions in the central nervous system may be absent or consist of necrosis and haemorrhage. Microscopically, a necrotizing encephalitis is characterized by perivascular cuffing and infiltration of cells, gliosis, haemorrhage, neuronal necrosis and swollen endothelial cells. Additional necrotic lesion may be found in non-neural tissues, including lymphoid tissues (with myeloid depletion of bone marrow, spleen and nodes), pancreas, adrenal cortex, liver, myocardium and small blood vessels.

Upon infection an inflammatory response ensues within 4 to 5 days and antibody production begins. Production of interferon and activation of natural killer cells starts, possibly playing a role in limiting early virus replication and dissemination. Antibody development appears to be important in limiting virus replication and spread, as well as preventing re-infection. Antibodies develop to all viral proteins and exhibit activities such as neutralization, inhibition of haemagglutination and complement fixation. Peak antibody titres develop typically within 2 weeks. This antibody may be effective in neutralizing virus, enhancing virus clearance, lysing virus in the presence of complement or lysing infected cells via complement or killer cells. In addition to systemic production, there may also be local antibody production in the CNS. Cell-mediated immunity also appears to play a role in virus clearance and protective immunity. Cytotoxic T cells can be identified as early as 3 to 4 days post infection. While most studies are of an indirect nature (in vitro and T cell transfer in mice), cytotoxic T cells probably contribute to both recovery and protective immunity (Biberstein and Zee, 1990).

Eastern equine encephalitis has caused fatal disease in ratites including ostriches (Smith, 1993), with infection having a short incubation period (20 h to 25 h), acute haemorhagic enterocolitis and a viraemia of up to two days. Birds aged from twenty to thirty-six months and juveniles are affected principally, with a morbidity rate of up to 70% and mortality of up to 87% (OIE, 2001, Tully and Shane, 1996). Multifocal areas of coagulative necrosis (hepatitis) in the liver have also been described in emus with eastern equine encephalitis (Veazey et al., 1994).

In horses, infection may result in an inapparent or mild infection (febrile response and mild depression) with no obvious signs of disease. Virus invasion of the nervous system is associated with clinical disease. Horses experiencing CNS involvement often move about aimlessly and may walk into obstacles. Later there is severe depression associated with unusual stances. Varying degrees of muscular paralysis may be observed prior to death, which typically occurs 2-4 days after onset of clinical signs. Horses that recover may have permanent CNS damage.

Eastern equine encephalitis cases were studied in man between 1988 and 1994. 36 patients were studied with 57 CT scans and 23 MRI scans from 22 patients. Mortality rate was 36% and 35% of survivors were moderately or severely disabled. Among MRI scans, results were abnormal for all 8 comatose patients and for 3 non-comatose patients who subsequently became comatose. CT results were abnormal in 21 of 32 patients with readable scans. Abnormal findings included focal lesions in basal ganglia , thalami and brain stem. Cortical lesions, meningeal enhancement and periventricular white-matter changes were less common (Deresiewicz et al., 1997).

Epidemiology

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Culiseta melanura is the documented enzootic vector of eastern equine encephalitis virus. Other species have been ranked as epidemic vectors (most to least likely): Culex salinarius, Anopheles quadrimaculatus, Aedes canadensis, Coquillettidia perturbans, Aedes vexans, Anopheles punctipennis, in test of susceptibility to per os infection and potential salivary transmission for eastern equine encephalitis (Vaidyanathan et al., 1997). Eastern equine encephalitis virus isolated from Aedesalbopictus in Florida, USA, in 1991 did infect and was transmitted by females of Aedes albopictus and Aedes taeniorhynchus in the laboratory (Turell, 1993). Scott and Lorenz (1998) demonstrated that eastern equine encephalitis virus reduces survival and reproduction (fitness) of Culiseta melanura, which is required for transmission of eastern equine encephalitis virus in North America. Mosquito virulence was not measurably attenuated in virus isolates recovered 55 years apart. The virus did not affect the ability of mosquitoes to obtain a blood meal (from chickens) or the rate of mosquito oocyte development.

The mosquito is of major importance in the epidemiology of eastern equine encephalitis. The cycle of infection is between water birds and mosquitoes in freshwater swamps, the mosquitoes then transfer the virus to humans and horses but both are considered 'dead-end' hosts as they do not develop sufficient viraemia to infect mosquitoes. Pheasants and water birds, rodents and other small mammals, reptiles and amphibians may also carry and transmit the virus. Peak times of the disease are routinely recorded in late summer with an abrupt halt when mosquito numbers are reduced due to adverse conditions such as cold or drought. Outbreaks of disease attributable to eastern equine encephalitis virus were evaluated in horses in Michigan, USA. Data on eastern equine encephalitis virus vectors, wild bird reservoir hosts, and incidental hosts, including horses and man were obtained from census reports and medical records compiled between 1942 and 1991. Michigan had all the elements required to sustain eastern equine encephalitis virus on a state-wide basis. Regions of Michigan with specific patterns for water drainage, specific mosquito species and areas with higher amounts of precipitation were associated with outbreaks of disease attributable to the virus in horses. Evaluation of environmental patterns, weather conditions and vector and reservoir host distributions may be useful to identify areas in Michigan and elsewhere in which horses and humans are at increased risk for an outbreak of the disease (Ross and Kaneene, 1996).

The mosquito vector must ingest an infective blood meal. The alphaviruses are biologically transmitted so the vector must be actually infected. The vector must obtain a blood meal from a host that is sufficiently viraemic. The level of viraemia required to infect the vector is dictated primarily by virus strain, the feeding insect or both. Upon ingestion, the virus initiates infection in the gut with eventual distribution to the salivary gland, where replication provides a ready source of virus to infect additional hosts upon feeding. The time required for this process is the extrinsic incubation period. Once infected, the vector remains infected for life. On pheasant farms, transmission may also occur by birds pecking each other. The overwintering reservoir of eastern equine encephalitis is probably wild swamp birds, though other mammalian and reptilian hosts may also play a role since virus has been isolated during winter months from cats, dogs, mice, foxes and skunks. (Biberstein and Zee, 1990).

Thirty six isolates of eastern equine encephalitis virus were obtained from 8 species of mosquitoes collected from 5 September to 18 October during an epizootic in southeastern Connecticut, USA. These included Culiseta melanura (19 isolates), Culex pipiens (8), Culiseta morsitans (3), Aedes sollicitans (2), Aedes cantator (1), Aedes trivittatus (1), Aedes vexans (1) and Coquillettidia perturbans (1). Isolations from Aedes cantator and Aedes trivittans were new to North American records, and those from Aedes cantator and Aedes solicitans represented the first infections of human-biting, salt-march mosquitoes with eastern equine encephalitis virus in Connecticut. With one exception, eastern equine encephalitis-infected Culiseta melanura were found at all sites where eastern equine encephalitis virus was isolated. Large numbers of eastern equine encephalitis isolates from Culiseta melanura and the collection of infected mosquitoes in residential woodlots and coastal salt marshes away from traditional red maple or white cedar swamp habitats, reaffirmed the importance of local populations of this mosquito for viral amplification and dispersal from swamp foci. No human or horses cases of eastern equine encephalitis were reported although this represented the largest number of isolations for eastern equine encephalitis ever recovered from field-collected mosquitoes in Connecticut (Andreadis et al., 1998).

In the laboratory Aedes sollicitans was more susceptible to infection with eastern equine encephalitis and viral dissemination than Aedes taeniorhynchus when fed on a chick with a viraemia of 107 +/- 0.1 pfu/ml. Infection rates in adults were not affected by rearing in salt concentrations ranging from fresh water to brackish water containing 2.4% sea salts. When fed on the same viraemic 6-day-old chicken, all 48 Aedes albopictus, reared in fresh water, became infected (Turell, 1998).

Geographic information system technology and remote sensing have identified wetlands as the most important element, accounting for up to 72.5% of the observed variation in the host-seeking populations of Aedes canadensis, Aedes vexans and Culiseta melanura when examining landscape features determining the risk of eastern equine encephalitis transmission in Massachusetts, USA. Stepwise linear regression demonstrated deciduous wetlands to be the specific wetland category contributing to the major class models (Moncayo et al., 2000). Population data in southeastern Massachusetts between 1982-90, showed Coquillettidiea pertubans, Aedes canadensis and Culex salinarius are more likely vectors of eastern equine encephalitis in Massachusetts than Aedes vexans, Anopheles punctipennis and Anopheles quadrimaculatus (Moncayo and Edman, 1999).

Tengelsen et al. (2001) studied immune responses to vaccination and challenge against eastern equine encephalitis in emus. An unvaccinated pen-mate became infected in the absence of mosquito vectors, presumably as a result of direct virus transmission between birds. Unvaccinated challenged birds developed viraemia (> 10 9 pfu/ml blood) and shed virus in faeces, oral secretions and regurgitated material.

Japanese quail (Coturnix coturnix) would facilitate sentinel-based eastern equine encephalitis surveillance programmes because of their small size and low cost, susceptibility to eastern equine encephalitis infection without mortality, and detectability of both virus and virus-neutralizing antibodies (Komar and Spielman, 1995).

Guy et al. (1995), inoculated tom turkeys with eastern equine encephalitis virus and examined their semen for the presence of the virus and its ability to transmit infection by artificial insemination. Mild depression and inappetance were observed. Toms were viraemic on days 1-2 days after inoculation and virus was shed in semen on days 1-5. Semen collected from eastern equine encephalitis-virus inoculated toms on days 1-2 was inseminated into turkey breeder hens. Eastern equine encephalitis virus was detected in one of the 10 hens after insemination. It was therefore concluded that semen is a potential vehicle for transmission of eastern equine encephalitis virus (Guy et al., 1995.)

A role is suggested for glossy ibis (Plegadis falcinellus) in the epidemiology of eastern equine encephalitis in New Jersey, USA, based upon blood infection rates (22.3%, n=130). They can come into contact with the vector species Culiseta melanura and Aedes sollicitans. The snowy egret (Egretta thula) was also found to be infected sometimes (1.8%, n=163) (Crans et al., 1991).

Impact: Economic

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The costs of eastern equine encephalitis virus infection are known to be substantial, but are infrequently recorded. The costs in man were calculated for a series of residents in eastern Massachusetts, USA. The majority of the expense was hospital costs. In the early years the costs were US $0.4 million, with a projected total cost of US $3 million in a residual case, including lost personal income (Villari et al., 1994). Costs of insecticide applications can cost as much as US $1.4 million depending on the size of area treated (Center for Disease Control, 2001).

Zoonoses and Food Safety

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Eastern equine encephalitis is a zoonotic disease, but a vaccine is available for humans. In a vaccination study in emus (Tengelsen et al., 2001), infected emus shed eastern equine encephalitis virus in secretions and excretions, making them a direct hazard to pen-mates and attending humans. Reasonable precautions should therefore be taken when handling livestock in areas of potential outbreaks of the disease.

Disease Treatment

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There is no treatment for eastern equine encephalitis virus, but the symptoms in horses can be treated supportively. Non-steroidal anti-inflammatory drugs may control pyrexia, inflammation and discomfort, for example, phenylbutazone, flunixin meglumine. Dimethyl sulfoxide can be useful in reducing inflammation and providing some analgesia and mild sedation. Convulsions may be controlled with pentobarbital, diazepam phenobarbital or phytoin. Secondary bacterial infections require appropriate antibiotics. Fluid solution, orally or intravenously, should be employed for fluid balance. Laxatives minimize the risk of gastrointestinal impaction, and oral or parenteral supplementation should be undertaken if anorexia persist for more than 48 hours. All animals should be provided with extra bedding, and protective leg wraps and head protection may be used to reduce self-induced trauma.

Prevention and Control

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Tengelsen et al. (2001) studied the humoral immune response of emus vaccinated with equine polyvalent and monovalent eastern equine encephalitis virus vaccines. All birds vaccinated were fully protected against an otherwise lethal dose of eastern equine encephalitis virus.

Serum-virus neutralizing antibodies were detected in serum and colostrum of 18 [Yorkshire x Hampshire] x Duroc sows vaccinated during pregnancy with commercially available vaccines (Encevac with Havlogen Encephalomyelitis Vaccine, Haver, Mobay Corp. and Encephaloïd, Fort Dodge Laboratories) against eastern equine encephalitis and antibodies were detected in serum from nearly all piglets from vaccinated sows following colostrum uptake. Serum-virus neutralizing antibody test titres were measured in colostrum and piglets at the next farrowing and additional vaccination of sows before the third farrowing led to elevated SVN titres in serum, colostrum and all pigs. Six piglets from vaccinated sows challenged at 8 to 9 days of age with 1x106 TCID50 eastern equine encephalitis virus did not develop disease.

Inactivated vaccines against eastern equine encephalitis are available commercially. Attenuated eastern equine encephalitis virus vaccines have not proven satisfactory. The vaccines licensed for use in the USA are prepared using combinations of eastern, western and Venezuelan equine encephalitis. In addition, tetanus toxoid and inactivated influenza virus have been combined with eastern equine encephalitis and western or eastern, western and Venzuelan equine encephalitis. Early vaccines were produced from virus propagated in embryonating chicken eggs and inactivated with formalin. Current vaccines are prepared from virus propagated in cell culture, and inactivated with formalin or monoethylamine. Standard strains of eastern equine encephalitis viruses that were isolated over 20 years ago have been used for vaccine production and have been proven to produce good immunity. According to OIE guidelines for vaccine preparation for horses, strains of eastern equine encephalitis virus that differ antigenically and in molecular structure have been identified from different geographical regions. However, the North American and Caribbean isolates appear to be similar. Viruses that are selected must be immunogenic and replicate to high titres in cell culture. Primary chicken embryo fibroblasts and Vero cells have been used for propagation of viruses used for vaccine production. The fibroblasts should be prepared from specific pathogen free embryos. Other susceptible cell lines could also be used. If a cell line is used, the master cell stock is tested to confirm the identity of the cell line, species of origin, and freedom from extraneous agents. If primary cell cultures are used, a monolayer from each batch of each subculture should be tested for extraneous agents including bacteria, fungi, mycoplasma and viruses. The master seed virus should also be tested to ensure freedom from bacteria, fungi, mycoplasma and extraneous viruses. The vaccines are administered by i.m. or intradermal routes in the cervical region in 2 doses given 2-4 weeks apart. Annual revaccination is recommended. All foals vaccinated before 1 year of age should be revaccinated before the next vector season.

Inactivated eastern equine encephalitis vaccine (PE-6) is available to man. Serological and recent molecular analyses of eastern equine encephalitis viruses have demonstrated marked difference between the 2 antigenic varieties of eastern equine encephalitis virus designated North American and South American (Strizki and Repik, 1995).

A population of whooping cranes (Grus americana) were protected from disease in Maryland, USA in 1989, by an eastern equine encephalitis vaccination programme (Olsen et al., 1997).

Vaccines developed for eastern equine encephalitis via inactivation are safe and efficacious and may also be used by bird farms if diluted. (Biberstein and Zee, 1990). Vaccines for horses and humans are safe and stimulate effective immunity and are also used in ratites with good results in endemic areas (OIE, 2001; Tully and Shane, 1996).

Husbandry Methods and Good Practice

Control of alphavirus infections can be achieved through vaccination and pest management programmes. Vector control can be approached through eliminating mosquito breeding sites by water control or spraying programmes. In the case of domestic bird farms, the use of tightly screened, insect-proof, rearing pens and locating pens away from freshwater swamps is also advisable. The practice of trimming the birds beaks can also minimize mechanical (pecking) transmission from bird to bird (Biberstein and Zee, 1990).

Aerial mosquito adulticide applications have been used in response to eastern equine encephalitis outbreaks and have targeted swamp habitats of the primary enzootic vector of the virus, Culiseta melanura. Organophosphate insecticide naled (Dibrom 14) has been insecticide of choice in these regions. Analyses of 11 years (1984-94) of mosquito collection data from Cicero Swamp (Onondaga county) and Toad Harbor Swamp (Oswego County) in relation to applications of naled showed it was successful in achieving short-term reductions in mosquito abundance. However, populations of Culiseta melanura increased 15-fold at Cicero Swamp. Preventive applications had no noticeable impact on the enzootic amplification of eastern equine encephalitis virus and isolations of virus following preventative applications resulted in additional spraying (Howard and Oliver, 1997).

Eastern equine encephalitis is a notifiable disease in Middle Eastern countries (Yehya, 1993).

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Links to Websites

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WebsiteURLComment
Center for Disease Control - EEE Fact Sheethttp://www.cdc.gov/ncidod/dvbid/arbor/eeefact.htm
ProMed Digest - EEEhttp://www.promedmail.orghttp://www.promedmail.org/pls/askus/f?p=2400:1202:36928::NO::F2400_P1202_CHECK_DISPLAY,F2400_P1202_PUB_MAIL_ID:X,14448
Rutgers University - EEE and horseshttp://www.rci.rutgers.edu/~insects/heee.htm

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