Rift Valley Fever
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Also Known As: RVF
Caused By: Rift Valley Fever Virus — RVFV
Rift Valley Fever (RVF) is a viral zoonotic disease belonging to the family Bunyaviridae in the Phlebovirus genus, possessing a segmented negative sense RNA genome. The disease has an episodic occurrence reemerging ever 5-25 years and is seasonal in its occurrence  . The occurrence of non immune animal populations every 5-25years combined with the introduction of RVF (due to rainfall) accounts for the explosive cyclical nature of the disease . RVF primarily affects animals but can infect humans and has the capacity to cause severe disease in both.
RVF has a wide economic impact due to livestock loss and trade restrictions as well as public health implications. It is a notifiable disease.
RVF virus was first identified in 1831 in the Rift Valley in Kenya during an investigation on a sheep farm and has since spread throughout Sub Saharan Africa emerging into North Africa in the 1970’s. the outbreak in Egypt in 1977-78 is considered to be the largest outbreak with 200,000 human cases reported  . In September 2000 it was reported for the first time outside of Africa, in Saudi Arabia and Yemen, probably introduced through infected livestock or mosquitoes  . The increase in cases in South Africa may be due to the end of an inter epizootic period  . Outbreaks are frequently reported though there is no evidence that it has spread to previously uninfected countries in the last 10 years, though it is hard to monitor changes in disease occurrence due to the cyclical occurrence of epidemics . Most recently RVF was reported in Mauritania in November 2012 . See a map detailing current outbreaks.
A number of mosquito species (Aedes, Culex, Mansonia, Anopheles) are implicated as vectors of RFV, the most important being Aedes and Culex spp. They are responsible for both maintenance and amplification of RVF.
Mosquitoes can be infected via feeding on infected animals. Vertical transmission can also occur (particularly in Aedes spp); female infected mosquitoes lay virus infected eggs leading to a new generation of infected mosquitoes. Vertical transmission is important in the survival of the virus as the eggs laid by the female can survive for many months in dry conditions, hatching after a period of rain and so increasing spread post rainfall leading to epizootics . Once animal infection has occurred mosquitoes are then responsible for amplifying infection. Aedes mosquito numbers decrease following rain but Culex tend to breed in more permanent water sites, hence the continuation of virus spread. 
RVF affects 4 areas: 
Dambos (e.g. East Africa) – Shallow depressions, often near rivers, that fill with water during the rainy season. Vertical transmission in mosquitoes occurs here.
Semi-arid (e.g. Senegal, Mauritania) – At temporary water points. It is unknown how the virus persists here, presumably either via vertical transmission or reintroduction of virus through visiting herds.
Irrigated areas (e.g. Nile Delta and Senegal River Valley). Yearlong viral transmission occurs here as the permanent water favours Culex breeding.
Temperate and Mountainous (e.g. Madagascar) – Virus is transmitted by vectors through cattle movement.
No outbreaks have been reported in urban areas.
Zoonotic transmission occurs through direct or indirect contact with infective blood or organs through slaughter, assisting with births and carcass disposal amongst other means. Faecal shedding of virus also occurs, as does spread through nasal and ocular secretions. Aerosol infection has also occurred within laboratory workers . Consuming unpasteurised or uncooked milk has also been associated with infection and seropositivity   . Mosquito bites have resulted in infection, and blood feeding flies also have the potential to transmit infection. There is no evidence of human to human transmission.
Virus particles are shed in milk but animals have not been infected via suckling or ingestion of milk  .
It is currently unknown if there are animal reservoirs of RVF between outbreaks. The namaqua rock rat and bats have been implicated and have been shown to be capable of infection but the potential impact of this is unknown.  Low levels of circulation between livestock or wild ruminants and mosquitoes (sylvatic cycle) is also likely to occur.
Outbreaks occur after heavy rain and flooding due to favourable breeding conditions for mosquitoes.
RVF can also be spread by the introduction of infected livestock into previously unaffected areas where mosquitoes are present.
A large number of animal hosts are susceptible, many producing high enough levels of viraemia to then infect mosquitoes.
RVF causes severe disease in animals, mainly cattle, sheep, goats and camels, with sheep being more susceptible. Bos Taurus cattle and other European breed imported into Africa appear highly susceptible to RVF.
Age is an important factor in determining the severity of the disease, young stock are more susceptible – 90% of infected lambs die whereas in adult sheep mortality can be as low as <10% . Small ruminants are also more susceptible. Pigs are resistant to low doses of RVF but high doses can cause viraemia  . During an outbreak in Egypt RVF virus was also isolated from horses as well as camels .
Other species (e.g. dogs and cats) have been infected experimentally and have become viraemic. The only species that are resistant are reptiles, birds and amphibians .
RVF has an incubation period of 1-6 days (12-36 hrs in lambs). Once in the lymph nodes viral replication occurs which leads to viraemia and systemic infection. Spontaneous abortions are seen as the hallmark of RVF outbreaks. Pregnant animals can abort at any stage often with 100% of stock aborting.
Newborn lambs and kids are highly susceptible to RVF, presenting with pyrexia and anorexia shortly followed by death 24-36hrs after infection.  In newborn lambs hepatocytes of the liver are the predominant target cell with hepatic necrosis being a significant post mortem finding. Other organs affected include the gall bladder (haemorrhage and oedema), gastrointestinal tract haemorrhage, lymph node haemorrhage, cutaneous haemorrhage and haemothorax. 
Signs in older lambs, kids, calves and adults vary from acute to subclinical (20-70% mortality), Signs can include fever (lasts 24-96hrs), weakness, bloody diarrhoea, abdominal pain, photosensitivity, anorexia, excessive salivation and decreased milk production. Signs in adult cattle are most often subclinical with less than 10% mortality.  
Camels display signs similar to those seen with Pasteurellosis infection, though infection can also be subclinical or asymptomatic. Abortions can also occur. During the 2010 outbreak in Mauritania 2 forms of disease were observed in camels; a hyperacute form causing sudden death in <24hrs and an acute form causing fever, ataxia, respiratory signs, icterus, oedema, foot lesions and neurological signs. If haemorrhagic signs were observed death occurred in a few days.
Differential diagnosis should include: bluetongue, Wesselbron disease, Enterotoxaemia of sheep, Ephemeral fever, Brucellosis, Vibrosis, Trichomonosis, Nairobi sheep disease, Heartwater, Ovine enzootic abortion, plant toxicity, bacterial septicaemias, Rinderpest, Anthrax.
Humans develop malarial-like disease. High risk individuals include farmers, veterinarians and abattoir staff. Mild disease is most common but severe hepatitis, encephalitis and ocular damage can develop. The usual presentation is of sudden onset fever, myalgia, biphasic behaviour and gastrointestinal disease.
Following infection viraemia is often high (though short lived) so the virus can be easily detected in the blood shortly after. Tissue samples can also be used in dead animals or aborted foetuses.
The nucleocapsid protein is used as the antigen of choice in serological assays. Blood samples can be used to detect the virus during the early phase using virus propagation, antigen detection and RT-PCR.
During the acute stages ELISA or EIA can be used to confirm the presence of IgM antibody to the virus, which allows recent infections to be diagnosed. ELISA’s based on recombinant RVF virus proteins have been developed which negates the need for biosecure facilities and are used in a number of species..Cross reactions may occur with other phleboviruses
RT-PCR is the standard method used in most laboratories as it has a high sensitivity. This is useful for rapid diagnosis and can also be used to detect RVF virus in mosquito pools.
Virus neutralisation tests (VNT) are very specific and sensitive and can be performed in a biosecure laboratory. They are also the prescribed test for international trade, though it cannot differentiate between vaccinated and infected animals. It is the only method to detect functional antibodies though a low level of cross reaction to some other phleboviruses has been observed. Plaque reduction neutralisation assays are the most commonly used VNTs and involve incubating the virus and heat inactivated serum allowing the virus to infect. Four to six days later the presence of cytopathic plaques is observed.
Haemagglutination inhibition (HI) and complement fixation assays are available but show extensive cross reactivity with other phlebovirus species. HI assays are used in non endemic areas but animals previously infected with other phleboviruses may show a positive result. Immunofluorescence can also be used.
Definitive confirmation can be carried out by virus isolation, however due to the zoonotic risk this can only be carried out in biosecure facilities.
Histopathology on tissue samples will show cytopathology and immunostaining can be used to identify RVF antigen in cells. On post mortem during the viraemic stage, widespread petechiae and ecchymoses on serous surfaces and organs will be seen and present in the body cavities. In older animals, the liver is enlarged and inflamed, with many foci of necrosis which are bronzed and jaundiced. The gall bladder may also be distended and haemorrhagic. Lymph nodes are enlarged and their germinal centres may be necrotic on closer examination. Extensive subcapsular haemorrhage in the spleen is usual. Renal changes include oedema and congestion. Epicardial and endocardial haemorrhages are often present on the heart.
No treatment is available.
Herd monitoring has been used extensively in Africa as a means of building up a picture of disease spread as well as to identify risk areas. Sentinel herds in representative areas are used for monitoring. Blood is collected initially and, providing no seropositivity is detected, animals are sampled at the beginning of the rainy season and every four to six weeks up to the end of the rainy season. This allows the early detection of any epidemics. Monitoring is also important as RVF outbreaks in animals precede outbreaks in human populations, as well as identifying increases in vector numbers.
During an outbreak, control at slaughterhouses to reduce zoonotic spread. Contact with animals, both direct and indirect, should be avoided. Movement restrictions should be enforced to prevent movement of the disease from diseased areas to RVF free areas; however movement restrictions within endemic countries have had little impact on the spread of RVF.
Forecasting is key as outbreaks occur following a period of heavy rain, thus is rain is forecast preventative measures can be implemented beforehand to help prevent/ lessen the impact of an outbreak.
Mosquito breeding sites should be reduced through drainage and larvicidal measures. Methoprene spraying, larvicidal toxins and controlled burning can be used, though low level aerial spraying has been shown to have little effect.
Education of risk factors and mosquito bite prevention, appropriate clothing, use of insect repellent and mosquito nets should be undertaken. Local populations should be educated as to the risks of eating raw meat and milk products (though a fall in pH destroys the virus so some uncooked meat may be safe).
Modified live attenuated and inactivated virus vaccines are available.
The live attenuated Smithburn vaccine only requires one dose but may cause spontaneous abortion in pregnant stock. This vaccine has adverse effects in newborn kids and lambs and teratogenic effects or abortion in pregnant cows, ewes and goats. Protection is conferred to offspring via suckling. Antibody titres post vaccination are higher in sheep than cattle. One dose will protect for three years.
The formalin inactivated virus vaccine (with Aluminium Hydroxide adjuvant) requires multiple doses to achieve immunity, and annual boosters are needed as it is less immunogenic.It is safe to use in pregnant ewes. Biosecurity is also a consideration when using this vaccine as reversion to virulence is a possibility.
Vaccines should be administered prior to outbreak to prevent an epizootic. Vaccination during an outbreak may worsen the situation, especially if multi dose vials are used, due to the possibility of viraemic animals being vaccinated and then the same vial being used on a healthy animal, thus spreading the virus. The use of the Smithburn vaccine is restricted in non endemic areas and during outbreaks due to the possibility of reassortment and a return to virulence.
More recently a naturally attenuated virus vaccine has been developed (Clone 13) that is marketed in South Africa and Namibia. Testing in pregnant ewes and lambs demonstrated safety and it has been shown to be as effective as the Smithburn vaccine in conferring immunity.  The vaccine has however shown to cause neurological disease and paralysis in some vaccinated mice.
A live attenuated vaccine MP12 has been shown to be safe in newborns, lambs and pregnant cows and ewes. Malformation has occurred when the vaccine was administered to sheep during the first trimester (days 35-56) and viral shedding has been documented in macaques following MP12 vaccination. Colostrum from vaccinated ewes gives temporary immunity to lambs.
An R566 strain has been developed from MP12 and Clone 13 and has shown to confer immunity in laboratory experiments. 
Viral vector vaccines using sheep pox and lumpy skin disease viruses have been shown to give protection, and have the advantage that the diseases exist in the same habitats and could potentially confer protection to two diseases with one vaccination but the use is restricted to countries with sheep pox and lumpy skin disease due to the use of the vectors. A vaccine using Newcastle disease virus as a vector has also been developed.
A recombinant virus vaccine has been found to be safe and effective in pregnant and non pregnant ewes, even when challenged with the virus. This vaccine has the advantage that it would allow differentiation between vaccinated and previously infected animals using a DIVA ELISA test (as would vector vaccines).
DNA and virus particle based vaccines are currently being developed and have demonstrated some level of protection against RVF virus. Plant derived subunit vaccines are also being researched.
Vaccine storage and transport is an issue within developing countries; breaking of the cold chain may result in ineffective vaccines being administered. However, a study has shown that the use of a formalin inactivated vaccine transported 200km at ambient temperatures had no adverse effect on antibody responses.
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- ↑ Public Health England factsheet accessed June 21, 2013
- ↑ 2.00 2.01 2.02 2.03 2.04 2.05 2.06 2.07 2.08 2.09 2.10 2.11 OIE Rift Valley Fever factsheet accessed June 21, 2013
- ↑ 3.0 3.1 Imam, Z. E., Karamany R. El., Darwish, M.A. (1979) An epidemic of Rift Valley fever in Egypt 2. Isolation of the virus from animals Bull World Health Organ. 1979; 57(3): 441–443.
- ↑ 4.0 4.1 4.2 4.3 4.4 4.5 4.6 WHO Rift Valley Fever accessed June 23, 2013
- ↑ 5.0 5.1 5.2 [EFSA Panel on Animal Health and Welfare (AHAW); Scientific Opinion on Rift Valley fever. EFSA Journal 2013;11(4):3180. [48 pp.] doi:10.2903/j.efsa.2013.3180.]
- ↑ 6.00 6.01 6.02 6.03 6.04 6.05 6.06 6.07 6.08 6.09 6.10 6.11 6.12 6.13 6.14 6.15 6.16 Lagerqvist, N, Rift Valley fever virus vaccine strategies, Karolinska Institutet 2013
- ↑ LaBeaud AD, Muiruri S, Sutherland LJ, Dahir S, Gildengorin G, Morrill J, Muchiri EM, Peters CJ, King CH. (2011) Postepidemic analysis of Rift Valley fever virus transmission in northeastern kenya: a village cohort study, PLoS Negl Trop Dis. 2011 Aug;5(8).
- ↑ Mohamed M, Mosha F, Mghamba J, Zaki SR, Shieh WJ, Paweska J, Omulo S, Gikundi S, Mmbuji P, Bloland P, Zeidner N, Kalinga R, Breiman RF, Njenga MK, (2010) Epidemiologic and clinical aspects of a Rift Valley fever outbreak in humans in Tanzania, 2007, Am J Trop Med Hyg. 2010 Aug;83(2 Suppl):22-7. doi: 10.4269/ajtmh.2010.09-0318.
- ↑ Oelofsen MJ, Van der Ryst E. (1999) Could bats act as reservoir hosts for Rift Valley fever virus? Onderstepoort J Vet Res. 1999 Mar;66(1):51-4.
- ↑ 10.0 10.1 10.2 10.3 10.4 10.5 10.6 10.7 FAO Recognising Rift Valley Fever accessed June 23, 2013
- ↑ Ahmed B. Ould El Mamy, Mohamed Ould Baba, Yahya Barry, Katia Isselmou, Mamadou L. Dia, Ba Hampate, Mamadou Y. Diallo, Mohamed Ould Brahim El Kory, Mariam Diop, Modou Moustapha Lo, Yaya Thiongane, Mohammed Bengoumi, Lilian Puech, Ludovic Plee, Filip Claes,Stephane de La Rocque, and Baba Doumbia, (2011) Unexpected Rift Valley Fever Outbreak, Northen Mauritania, Emerging Infectious Diseases • www.cdc.gov/eid • Vol. 17, No. 10, October 2011
- ↑ FAO Signs of Rift Valley Fever accessed June 23, 2013
- ↑ 13.0 13.1 13.2 Lagerqvist N, Moiane B, Bucht G, Fafetine J, Paweska J.T., Lundkvist Å and Falk K.I. 2012. Stability of a formalin‐inactivated Rift Valley fever vaccine: evaluation of a vaccination campaign for cattle in Mozambique. Vaccine 30(46):6534‐40.
- ↑ Botros B, Omar A, Elian K, Mohamed G, Soliman A, Salib A, Salman D, Saad M, Earhart K. (2006) Adverse response of non-indigenous cattle of European breeds to live attenuated Smithburn Rift Valley fever vaccine. J Med Virol. 2006 Jun;78(6):787-91.
- ↑ Kortekaas J, Zingeser J, de Leeuw P, de La Rocque S, Unger H, Moormann RJ.,2011 Rift Valley Fever Vaccine Development, Progress and Constraints. Emerg Infect Dis. 2011 Sep;17(9)
- ↑ GALVmed Rift Valley Fever vaccination strategy accessed June 23, 2013
- ↑ Dungu B, Louw I, Lubisi A, Hunter P, von Teichman BF, Bouloy M (2010). Evaluation of the efficacy and safety of the Rift Valley Fever Clone 13 vaccine in sheep., Vaccine. 2010 Jun 23;28(29):4581-7
- ↑ von Teichman B, Engelbrecht A, Zulu G, Dungu B, Pardini A, Bouloy M (2011). Safety and efficacy of Rift Valley fever Smithburn and Clone 13 vaccines in calves. . Vaccine. 2011 Aug 5;29(34):5771-7
- ↑ Vialat P., Billecocq A., Kohl A., and Bouloy M. (2000). The S segment of Rift Valley fever phlebovirus (Bunyaviridae) carries determinants for attenuation and virulence in mice. J Virol 74:1538‐1543.
- ↑ Hunter P, Erasmus BJ, Vorster JH. (2002) Teratogenicity of a mutagenised Rift Valley fever virus (MVP 12) in sheep,. Onderstepoort J Vet Res. 2002 Mar;69(1):95-8.
- ↑ Bird BH, Maartens LH, Campbell S, Erasmus BJ, Erickson BR, Dodd KA, Spiropoulou CF, Cannon D, Drew CP, Knust B, McElroy AK, Khristova ML, Albariño CG, Nichol ST (2011) Rift Valley fever virus vaccine lacking the NSs and NSm genes is safe, nonteratogenic, and confers protection from viremia, pyrexia, and abortion following challenge in adult and pregnant sheep, J Virol. 2011 Dec;85(24):12901-9
- ↑ Kortekaas J, Zingeser J, de Leeuw P, de La Rocque S, Unger H, Moormann RJ Rift Valley Fever Vaccine Development, Progress and Constraints. Emerg Infect Dis. 2011 Sep;17(9)
The datasheet was accessed on 8 June 2011.
This article has been expert reviewed by Nick Lyons MA VetMB CertCHP MRCVS
Date reviewed: July 8, 2012
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