Selected content from the Animal Health and Production Compendium (© CAB International 2013). Distributed under license by African Union – Interafrican Bureau for Animal Resources.
Whilst this information is provided by experts, we advise that users seek veterinary advice where appropriate and check OIE manuals for recent changes to regulations, diagnostic tests, vaccines and treatments.
This work is licensed under a Creative Commons Attribution-NonCommercial-NoDerivs 3.0 Unported License.
Identity Pathogen/s Overview Distribution Distribution Map for Africa Distribution Table for Africa Hosts/Species Affected Host Animals Systems Affected Epidemiology Impact: Economic Zoonoses and Food Safety Pathology Diagnosis Disease Course Disease Treatment Prevention and Control References Links to Websites OIE Reference Experts and Laboratories Images
Preferred Scientific Name
Rift Valley fever
International Common Names
fièvre de la Vallée du Rift, la fièvre de la Vallée du Rift
Rift Valley fever virus
A new disease syndrome, named Rift Valley fever (RVF), was first described in 1930 in the Rift Valley of Kenya, although it may have occurred earlier. One of the plagues of Egypt reported in the Bible had features, which suggested that it might have been due to RVF. The first outbreak was identified in sheep, where there were very large numbers of abortions with many deaths in newborn lambs and older animals. Several thousand were affected. These were sheep of a breed imported into Kenya, the indigenous animals kept nearby did not show any disease. People associated with the infected sheep became ill, with a dengue/malarial like disease. The disease was shown to be due to a virus, which was transmitted by mosquitoes. The losses ceased within a few days, when the sheep were moved to a higher altitude above the Rift Valley. Subsequently, the disease occurred irregularly at 3- to 10-year intervals in Kenya. It was then recognised in South Africa in 1950-51, when an epizootic involved much of the country.
Sheep, goats, cattle, camels and man have been affected in the many subsequent epizootics, which have occurred throughout the Ethiopian faunal region. An extension beyond this range RVF has occurred to Egypt, where a dramatic epizootic in 1997 resulted in much human disease and huge losses amongst the domestic animal populations. RVF cause an acute episode of human disease, with at least 600 deaths and more than 60,000 severe clinical cases. The total morbidity was thought to be measurable in hundreds of thousands, and the resources of the hospitals in the affected areas were severely strained by the numbers of cases presenting daily. Most cases were thought to arise from mosquito bites, but many of the human cases followed close contact with infected animals, and the aerosol route of infection appeared to be responsible.
RVF can be considered to be an emerging zoonotic disease of importance, affecting the domestic livestock populations in Africa, especially where these have been improved by the introduction of genotypes originating in Europe and elsewhere. Indigenous animals appear to have a degree of resistance to the virus. The human disease component has been highly significant in recent East African and West African epizootics.
This disease is on the list of diseases notifiable to the World Organisation for Animal Health (OIE). The distribution section contains data from OIE's WAHID database on disease occurrence. Please see the AHPC library for further information on this disease from OIE, including the International Animal Health Code and the Manual of Standards for Diagnostic Tests and Vaccines. Also see the website: www.oie.int.
The presence of Rift Valley fever (RVF) virus or specific antibody to the virus has been detected in nearly all of the sub Saharan African countries. Clinical disease in man or domestic animals has been reported in the following countries:
Kenya, South Africa ,Uganda, Tanzania, Zambia, Zimbabwe, Botswana, Malawi, Nigeria, Mozambique, Sudan, Senegal, Zaire (Congo Democratic Republic), Angola, Mauritania, Madagascar. Central African Republic, Ethiopia, Somalia, Egypt
Specific antibody to RVF has been found in most other African countries without any manifestation of a disease problem. These include :
Mali, Gambia, Cameroon, Mali, Chad, Guinea, Burkina Faso.
Zaire, Niger, Angola, Côte d'Ivoire.
RVF virus and/or RVF specific antibody have been detected across the wide range of biotopes present in the African continent. RVF virus is present in the central African tropical forests, the wet and dry bushed and wooded savannah grasslands, the semi desert and edge of the desert, the highland plateaux with their temperate type forest, bushed grasslands and the coastal forest zones. Epizootic disease occurs wherever livestock numbers (especially exotic) are greatest and this tends to be in the bushed and savannah grassland zones, and wherever there are floodplains or temporary water pools occurring after rains. These occur right up to the edge of the Sahara. Irrigation sites create large populations of mosquitoes at certain seasons and allow huge amplification of RVF virus. RVF has occurred from the Cape to Cairo, but not as yet in the Mahgreb countries of North Africa , nor further into the Middle East from Egypt.
There have been reports from Russia of the occurrence of RVF in Afghanistan and northern India, and a suspicion of RVF in southern India. The latter was a result of misinterpretation of the serological cross reactions, which occur amongst Phlebotomus fever group viruses, and may also be the explanation for the reports from the north. Certainly, no massive animal nor human epizootic, which might have been due to RVF, has ever been recorded outside Africa.
For current information on disease incidence, see OIE's WAHID Interface.
The Republic of South Africa (RSA) is the only country in Africa that has consistently reported outbreaks of RVF during the past four years (AU-IBAR, 2011). It is still not clear whether the outbreaks reported in neighbouring Botswana (in 2010) and Namibia (in 2010 and 2011) were linked to the situation in RSA. During 2011, RVF was reported to AU-IBAR by only 4 countries - RSA (74), Mauritania (3), Namibia (2) and Comoros (1) (see table below). Although the number of RVF affected countries in 2011 remained the same as that of 2010, there was a remarkable reduction in the number of affected epidemiological units from 330 in 2010 to 80 in 2011 (with 92.5% of the outbreaks occurring in South Africa).
*NS: Not Specified
Though available data may not be enough to give full details of RVF progression in RSA over the past four years, the zoonotic nature of the disease and the magnitude of its spread in that country during the past few years justify its special consideration. The number of outbreaks has steadily increased from 34 in 2008 to 41 in 2009 followed by an eight fold jump to 330 in 2010. However, the number of reported outbreaks declined remarkably five-fold in 2011 with only 74 as compared to the 330 epidemiological units affected in 2010. Although not clearly indicated in their monthly reports, this outcome may be attributed to the coordinated control measures put in place by the RSA.
= Present, no further details = Widespread = Localised
= Confined and subject to quarantine = Occasional or few reports
= Evidence of pathogen = Last reported... = Presence unconfirmed
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 information for individual references may be available in the Animal Health and Production Compendium. A table for worldwide distribution can also be found in the Animal Health and Production Compendium.
|Country||Distribution||Last Reported||Origin||First Reported||Invasive||References||Notes|
|Algeria||Disease never reported||OIE, 2012|
|Angola||Disease never reported||OIE, 2012|
|Benin||No information available||OIE, 2009|
|Botswana||Disease never reported||OIE, 2009|
|Burkina Faso||No information available||OIE, 2009|
|Burundi||Disease not reported||OIE, 2012|
|Cameroon||Last reported||2009||OIE, 2012|
|Cape Verde||Disease never reported||OIE, 2012|
|Central African Republic||Disease not reported||OIE Handistatus, 2005|
|Chad||No information available||OIE, 2009|
|Comoros||Restricted distribution||AU-IBAR, 2011|
|Congo||No information available||OIE, 2009|
|Congo Democratic Republic||Disease not reported||OIE Handistatus, 2005|
|Côte d'Ivoire||Disease not reported||OIE Handistatus, 2005|
|Djibouti||Disease not reported||OIE, 2012|
|Egypt||Last reported||2004||OIE, 2012|
|Equatorial Guinea||No information available||OIE, 2012|
|Eritrea||Disease not reported||OIE, 2009|
|Ethiopia||Disease not reported||OIE, 2009|
|Gabon||Disease never reported||OIE, 2012|
|Gambia||No information available||OIE, 2009|
|Ghana||Disease not reported||OIE, 2012|
|Guinea||Disease not reported||OIE, 2009|
|Guinea-Bissau||No information available||OIE, 2009|
|Kenya||Last reported||2007||OIE, 2012|
|Lesotho||Disease never reported||OIE, 2012|
|Libya||Disease never reported||OIE, 2012|
|Malawi||Disease not reported||OIE, 2012|
|Mali||Disease never reported||OIE, 2012|
|Mauritania||Restricted distribution||OIE, 2012|
|Mauritius||Disease never reported||OIE, 2012|
|Morocco||Disease never reported||OIE, 2012|
|Mozambique||Last reported||1963||OIE, 2012|
|Nigeria||Disease never reported||OIE, 2012|
|Réunion||Disease never reported||OIE Handistatus, 2005|
|Rwanda||Disease not reported||OIE, 2009|
|Sao Tome and Principe||Disease not reported||OIE Handistatus, 2005|
|Seychelles||Disease not reported||OIE, 2012|
|Sierra Leone||Disease not reported||OIE, 2012|
|Somalia||No information available||OIE Handistatus, 2005|
|South Africa||Present, no further details||AU-IBAR, 2011|
|Sudan||Last reported||2008||OIE, 2012|
|Swaziland||Last reported||2008||OIE, 2012|
|Togo||Disease never reported||OIE, 2012|
|Tunisia||Disease never reported||OIE, 2012|
|Uganda||Disease never reported||OIE, 2012|
|Zambia||Disease not reported||OIE, 2009|
|Zimbabwe||Last reported||2010||OIE, 2012|
Man is an important disease host, other disease hosts are the domestic ruminants, cattle sheep and goats. There are distinct differences in susceptibility amongst different breeds and strains. Those breeds of Bos taurus and of European wool sheep that are exotic to Africa, tend to be highly susceptible and the indigenous breeds relatively insusceptible. The wild ruminant species found in the enzootic areas are not disease hosts, no clinical RVF is seen when disease occurs in domestic ruminants in the same habitat. However, many develop antibody to RVF. Camels, the Asiatic buffalo and horses have a transient viraemia and pigs are much less susceptible. Primates have been extensively used in laboratory studies of RVF and most species have shown a susceptibility, similar to that seen in man. Laboratory rodents and hamsters are highly susceptible and are used extensively in laboratory studies of RVF. Distinct genetic differences in susceptibility have been elegantly demonstrated to occur in rodents, as in the domestic animals. Certain inbred strains are highly susceptible, and others are resistant. The differences are related to one or more genes. There is evidence to show that rats do become infected during periods of epizootic RVF, but generally only a small proportion of their populations. All the susceptible animals develop viraemia, which varies in titre and duration according to the level of susceptibility. It has not proved possible to infect avian species, and RVF has occasionally been isolated from bats. They are thought to be accidental hosts fed upon by RVF vector mosquitoes.
|Bos indicus (zebu)|
|Bos taurus (cattle)||Domesticated host|
|Camelus dromedarius (dromedary camel)||Domesticated host|
|Capra hircus (goats)||Domesticated host|
|Ovis aries (sheep)||Domesticated host|
Blood and Circulatory System - Large Ruminants
Blood and Circulatory System - Small Ruminants
Digestive - Large Ruminants
Digestive - Small Ruminants
Multisystem - Large Ruminants
Multisystem - Small Ruminants
Nervous - Large Ruminants
Nervous - Small Ruminants
Reproductive - Large Ruminants
Reproductive - Small Ruminants
Respiratory - Large Ruminants
Respiratory - Small Ruminants
Incidence in man
Sporadic cases occur in man in forest and forest edge zones, which are identified occasionally at hospitals and clinics. During the epizootic periods in Africa, cases occur amongst those people associated with the animal disease hosts, such as farmers, veterinarians and abattoir staff. Most cases are of the mild variety with occasional more severe hepatitis and other complications. Only 1-2% of these become severe enough to warrant referral to hospital with complications, such as severe hepatitis, ocular/retinal damage or encephalitis. The fatal haemorrhagic cases are rarely seen throughout most of the traditional epizootic areas for the disease in southern and eastern Africa.
The situation has been different, when the disease has occurred in the arid and semi-arid zones and in the first epizootic in Egypt. Many fatal haemorrhagic cases were seen in the Egyptian epizootic and have also occurred when Rift Valley fever (RVF) has behaved in an epizootic manner in the semi arid zones of East and the Horn of Africa. They are only thought to represent less than 0.5-1% of cases. Haemorrhagic cases are not seen in humans, when the epizootics are simultaneously affecting many more domestic animals in the high potential higher rainfall climatic zones, with apparently much greater amplification of the virus with greater numbers of vectors. Nutritional, intercurrent disease such as malaria or schistosomiasis, or other factors may determine the outcome of RVF infection in man and account for this difference. Post epizootic prevalence of antibody is of the order of 5-40% in the general population, whilst higher prevalence may be detected in selected high risk groups.
Incidence in domestic animals
In epizootic periods
A high prevalence of antibody may be detected hosts, notably, cattle, sheep and goats, following periods of epizootic RVF activity. The highest figures are usually obtained from cattle, where 60-95% have been found to have IgM to RVF virus. Similar figures have been found in sheep but generally a lower prevalence is found amongst goats and camels. There are differences between epizootics in the same areas. For example, 90% sero-conversion may occur in one epizootic and then 15-25% in another. These figures are related to the level of challenge by infected vectors and reflect the epidemiological factors prevailing at the time.
In inter-epizootic periods (IEP)
Epizootics persist for 1-3 years before virus activity dies out. Virus activity may then not be detected for very long periods. In enzootic zones, such as central Africa or coastal forest, there may be some virus activity most years depending upon the rainfall and mosquito populations. Sporadic cases in humans have been reported in inter-epizootic periods and 1-5% of domestic animals in such areas may be found to sero-convert. In the bushed and wooded grasslands where epizootics are most severe, no virus activity may be detected for many years in the IEP.
In Egypt, RVF virus activity has disappeared in the 2-3 years after epizootics.
Seasonal distribution and periodicity
The disease follows the rainy seasons in most countries in Africa, and where this has a bimodal character, the epizootics may follow either of the rainy seasons. In West Africa for example, this results in RVF virus activity being detected from August to December. In Egypt, most virus activity has occurred between May and October, when the onset of frosts result in a fall in the vector populations. Peak mosquito populations are seen in June, July and August. In East Africa, RVF virus activity follows the March/April rains, with most cases in between May and August, but it may also follow the short rains in October, with cases in November through to January.
There may be some emergence of RVF-infected Aedes mosquitoes in most years in areas of high rainfall in Africa. These tend to be the central African tropical forest zones, and coastal and temperate type high altitude forest. The occurrence of sporadic clinical cases in humans and virus isolations from mosquitoes in forest situations have been consistent with this hypothesis.
In areas with relatively high rainfall, which have been excised from forest or are adjacent, some RVF virus activity may occur every 2-3 years. In the drier bushed and wooded grasslands, where most domestic animals are found, the epizootics occur in 5- to 15-year cycles and little or no RVF virus activity may be detected between epizootics. The RVF virus activity is usually detected in association with riverine flood water plains, dambo formations or water pans, whenever these are flooded for prolonged periods. In the arid and semi arid zones, a similar picture exists, with little or no virus activity between epizootics. That it occurs is evident, as 5-15% of adult cattle or camels (which live for many years) may have IgG antibody to RVF. Epizootic RVF in such areas occurs more rarely, and in the Horn of Africa recently, the interval was 35 years, although 15-30 years might be a more reliable range. In the Sahelian zones of Mauritania and Senegal, some RVF emergence may occur each year if the water pans become flooded.
Risk factors for human infection
The major risk factor is an association with domestic animals. Farmers, shepherds, veterinarians and abattoir workers are more susceptible than other members of society. An association with milk and milkers, can be correlated with their close contact with animals and their discharges. Milk is not a vehicle for infection in epidemics. Laboratory workers handling the carcases in post mortem rooms have also been infected. The close association of families, who share common housing with their animals, increases the risk of RVF infection. This has been evident in Egypt.
There does not appear to be a sex or specific age group more susceptible to the disease amongst populations closely associated with animals such as the nomadic pastoralists, herders and those milking cattle. Where sheep or goats have become infected, cases are suspected to have originated from ritual slaughter, where raw warm flesh is handled and eaten. Apparent differences in attack rates are usually explicable on such a basis.
An important observation from Egypt and elsewhere is that RVF is a rural and semi rural disease and does not cause problems in heavily populated towns and cities. This was notably the case in Egypt, where cases could be found in peri-urban situations but not urban. There was no evidence for the amplification of RVF virus in the urban situations.
Early work in Kenya showed a close association of epizootic RVF activity with periods of prolonged and heavy rainfall. The climate in much of Africa depends upon the characteristics of the inter-tropical convergence zone (ITCZ). This is the area of convergence of hot, dry air currents from the north of the continent with warm, humid air from the south. Its density and magnitude determine the levels of cloud cover and precipitation.
Initially, prediction of RVF virus activity was made on the basis of the occurrence of 2-5 times the mean annual rainfall, which caused flooding of grassland water pans or dambo formations. It was noticed that such flooded habitat, allowed huge mosquito populations to develop at periods when clinical RVF occurred. A statistic was derived from the actual rainfall figures, the number of rain days/month and the rolling mean, which correlated well with epizootics of RVF over 25 years.
The advent of Remote Sensing Satellite Imagery (RSSD) derived from Landsat, SPOT and later Synthetic Aperture Radar allowed the use of more sophisticated tools to predict RVF epizootic activity over much wider areas. Ground truth data was generated to correlate with changes in the Cold Cloud Density (CCD) and the Normalised Differentiated Vegetation Index (NDVI). This is a measure of the intensity of green vegetation. The NDVI index could then be correlated with the conditions, which allowed the emergence of the Aedes mosquito vectors of RVF. Pilot studies with the NDVI values in other parts of Africa have allowed Risk Assessments for epizootic RVF to be made, by calculating the difference between the normal, expected and the actual rainfall. The initial results appear to be promising and may be applicable on a much wider basis to similar biotopes in other parts of Africa. Ocean temperature changes have now been shown to be correlated with the RSSD data and provide a further tool for the prediction of epizootic RVF activity. Climatic changes usually occur on a sub-regional basis rather than continental.
Periodicity of epizootics
The periodicity of RVF epizootics, which occur at 5- to 15-year intervals in most epizootic areas, prompted research into possible vertebrate reservoir hosts for the virus during the inter-epizootic periods (IEPs). Wild and domestic ruminant populations, rodents, birds and bats have all been investigated. No hosts have been identified which could satisfactorily explain the virus persistence. Cryptic virus cycles could have occurred in indigenous livestock species, and amplify the virus to the point where it manifests as human disease, in the absence of any obvious disease in their domestic ruminants. Longitudinal serological studies have not shown that this has been happening. There did not appear to be any virus activity in most of the epizootic areas in the IEPs. Intensive longitudinal studies in areas severely affected during epizootics would show no evidence for any virus activity during the IEP. Some virus activity was shown to occur most years in the very high rainfall forest and forest edge zones in both southern, eastern and central Africa. The disease in domestic animals requires the presence of large numbers of susceptible genotypes to become clinically evident.
Evidence from field studies
The answer to this problem of virus persistence was suggested by artificial flooding of the dambo (water pan) formations in an epizootic area in the Central Highlands of Kenya. Many millions of larvae hatched and these were virtually entirely of Aedes mcintoshi (previously Aedes linneatopennis). RVF virus was isolated from adult Aedes mcintoshi mosquitoes (including males), which developed in the laboratory from larvae collected from the field. The hypothesis was presented that RVF virus, persisted throughout the IEP in the eggs of the floodwater-breeding Aedes mosquitoes. Trans-ovarian transmission of the virus provides an explanation for the persistence of the virus through the IEP and for its simultaneous emergence throughout epizootic areas, which experience similar climatic conditions. This has been observed in all RVF epizootics. Areas separated by a thousand kilometres or more will show clinical cases of RVF virtually at the same time.
This mosquito species or other floodwater-breeding Aedes of the Aedes (Neomelaniconium) group, have been found throughout much of the epizootic range of RVF in Africa. Other floodwater-breeding Aedes species are more prevalent and may be more important in the different regions. In Senegal, for example, Aedes ochraeus, A. dalzieli and A. vexans appear to be more common in the water pans. No true enzootic system has been identified in Egypt, and the disease is thought to recur following fresh introduction of virus. In Egypt, Aedes caspius and Culex pipiens are though to be the most important vector species. Culex pipiens is also extremely common in many African RVF epizootic areas. These species feed predominantly upon human and domestic ruminant hosts.
The persistence of water for extended periods of time in the flooded dambo or water pans allows further mosquito species to colonise and breed in this habitat. The very rapid disappearance of the water within 7-14 days will not allow generation of epizootic RVF, although some emergence of RVF-infected vectors may have occurred, and further oviposition cycles initiated. It is the secondary cycles, which involve a wide range of mosquito species, many of which are capable of transmitting RVF, which facilitate amplification to epizootic proportions. Species which are thought to be important in RVF transmission on the basis of laboratory transmission experiments and field isolations are listed in the Vectors section of the datasheet
Evidence of vector competence
Many of the vector mosquito species have been demonstrated to replicate RVF in a biological cycle in laboratory experiments, and to be capable of transmitting the virus. Thus members of Aedes (Neomelaniconion) and Aedes (Stegomyia), Culex, Mansonia, Anopheles and Eretmapodites, have all been shown to transmit the virus.
The vector competence has not been determined for all of these species, but most have tested positive for RVF virus when caught and assayed in epizootic situations.
Humans, other primates, cattle, sheep, goats, camels, wild and domestic buffaloes have been shown experimentally to replicate RVF virus to a level where infection of other feeding mosquitoes would be likely to occur. However, there is considerable variation in the response to infection with RVF virus in most animal species. Some breeds and strains are much more susceptible than others. Species such as the horse, which develop lower levels of viraemia, are not present in sufficient numbers in epizootic areas to play any role in amplification of the virus. The same applies to dogs and cats, which play no role in epizootic situations, although they do develop viraemia and disease. In most situations, cattle, sheep, goats or camels are likely to outnumber other potential hosts by 10 to 1, or greater. Rodents and bats have been shown to become infected by RVF in serological investigations. There are higher proportions positive for RVF after periods of epizootic activity (2-15%). Most of the vector mosquitoes which have been studied do not show any preference to feed upon these hosts, when ruminants are available. They may play a more important role in virus maintenance in the tropical forest zones. Those avian species which have been examined are insusceptible and do not develop a viraemia. A suggestion that RVF is spread by the migration of the Sudan dioch (Quelea quelea aethiopica) along the Rift Valley from north to south was not supported by experiment.
Other transmission mechanisms
RVF has been isolated from Culicoides, which do not replicate RVF virus biologically though they can transmit the virus mechanically. Other biting flies have also been shown to be capable of transmitting RVF mechanically. With virus titres of 109 per 0.1 ml of blood in the animals, it is clear that this may be an important component of the transmission cycles. Culicoides feed in huge numbers at crepuscular periods, and the abdomens of cattle and camels have been seen to be black with thousands of feeding mosquitoes in epizootic conditions. RVF virus transmission following interrupted feeds is highly likely to occur, Stomoxys, Tabanids, Glossina (tsetse flies) may all play a role in epizootic situations.
The ritual slaughter of animals moribund with RVF, in Muslim societies, has been associated with infection of all involved. Family slaughter amongst nomadic pastoralists also accounts for many cases of RVF. Post mortem examinations in the field and at laboratories have resulted in many cases amongst farmers, veterinarians and laboratory workers. Abattoir staff in epizootic areas experience much exposure to RVF virus and up to 40% have been shown to have antibody to RVF after epizootics. Infected blood and discharges from haemorrhaging patients may also serve as a source of infection. Milk is not a vehicle for infection. Nosocomial infections from patients to medical staff and others on hospitalisation, which are so important with Ebola virus, do not occur with RVF. Animal to animal infection by contact, by aerosol or other means has been demonstrated experimentally and may occur in epizootics.
The field and experimental data all support a conclusion that trans-ovarial transmission of RVF virus in floodwater-breeding mosquitoes, which have a single generation cycle, provides the explanation for the enzootic-epizootic character of RVF seen in Africa. The epizootics appear to be triggered by periodic cycles of prolonged rainfall, although other factors may be involved. The capacity to make some prediction of the risk of RVF epizootic activity exists by using Remote Sensing Satellite Imagery (RSSD) data.
Ecological dynamics - climate and weather
The climate and weather play a major role in determining the emergence of the infected vector populations and the amplification of the virus. Critical features are the development of greater cloud cover and density, accompanied by regular and significant precipitation. This increases the level of the water table and creates the pre-epizootic conditions, which can be monitored and identified by RSSD. This rise in the water table results in an increase in vegetation quantity and green density, which can be quantitated by RSSD as the Normalised Differentiated Vegetation Index (NDVI), which is based upon the ratio of brown/green vegetation density readings from the satellites. The critical NDVI value which was found to be associated with the emergence of RVF-infected mosquitoes in Kenya and Zambia was 0.45. This may not apply in other biotopes. Flooding occurs as the water table rises in dambos and water pans, or as a result of water spillage from rivers and water-courses to cover the riverine flood plains. This flooding, if it persists, allows the emergence of the RVF reservoir, the mosquito vectors.
Ecological dynamics - the vector
The floodwater-breeding Aedes mosquitoes emerge as larvae within 2-4 days of flooding and as adults 6-8 days later, according to the prevailing temperatures. The range of movement from the site of emergence is dependent largely upon the prevailing winds. The overall dispersion for males and females is about 0.15 km. An emerging population survived for approximately 45 days in an experimental study, but this may be greater at times of an RVF epizootic, when the cloud cover is greater and the vegetation and humidity create habitat, which favours longer survival. Aedes mcintoshi and other floodwater-breeding Aedes spp. have been shown to have a feeding preference for domestic cattle, although humans and other ruminant species are also fed upon. Huge populations of Aedes and other mosquito species may be seen in association with the flooded areas, and these persist whilst the flood remains, although the species population structure changes. The Aedes spp. oviposit in the water and vegetation associated with the flooded areas. Most other species oviposit at the waters edge or in small isolated flooded pockets.
The extension of RVF, from the enzootic and epizootic areas in sub-Saharan Africa into Egypt, has posed problems for epidemiologists. The movement of insect vectors both within and beyond continental boundaries has been well documented for insect pests, for malarial mosquitoes and other insect-borne viruses. The appearance of RVF in Egypt on two occasions has followed exceptional but short periods, when convection and low level air currents could have transported infected insect vectors from active foci of RVF virus activity 500 km to the south. The prevailing air currents are generally in directly the opposite direction. A period 5-6 days, when this reversal occurred in 1993, was followed 14-21 days later by RVF cases in Aswan. The disease then spread up the Nile to the delta within 3 months, again, it is suggested, by insect movements. Egypt provides a very receptive area for RVF amplification in summer, as there are large populations of Culex spp. and other mosquitoes. The virus persists for 1-3 years and then it is not possible to show any residual foci of RVF virus activity.
Ecological dynamics - the vertebrate hosts
RVF as a clinical disease is manifested in domestic animals in Africa, wherever they have been imported to improve livestock production. Early outbreaks were apparent in the huge flocks of wool sheep in eastern and southern Africa. New cattle (Bos taurus) breeds imported into the continent to improve milk and beef production proved to be highly susceptible to RVF virus. Abortion and newborn deaths were the prominent signs in these exotic cattle, although mortality has been seen in all age groups. The indigenous Bos indicus breeds of cattle and the hair sheep and goats owned by the pastoralists in many parts of Africa are relatively resistant to RVF. They develop a short period of viraemia, with or without a transient febrile reaction, and while some may abort, the numbers are usually not remarkable. The genotype of an animal is a major determinant of the outcome of an RVF infection. RVF virus activity in areas where the domestic animals are resistant may only be signalled by human cases. Quite extensive virus amplification may be revealed by serological investigations.
The distribution of the resistant genotypes is thought to be within the limits of enzootic virus activity in wetter forest derived areas and grasslands. In the epizootic areas to the north in some semi-arid and arid zones in the Sahel, and possibly also in the south of the continent, proportions of susceptible genotypes are thought to increase.
The impact of RVF in susceptible animals in epizootic form is sufficiently dramatic to drive diagnostic activities, which should be able to identify the cause. Virus isolation and identification where possible (vide supra), is a rapid and straightforward procedure. Frequently, the facilities to do this may not be available, and the imuno-staining of fixed liver specimens or examination of the sera taken from aborted animals are the best strategies to make a diagnosis. Sera can be examined for IgM to RVF virus in an ELISA test system. This is the method most widely used and kits for this purpose should be available. An emergency preparedness with such a diagnostic capacity, should be established at national laboratories in all the target countries in Africa. These activities will give a good indication of the level of RVF virus activity in an outbreak.
Sporadic or isolated cases would clearly be missed if surveillance were driven only by gross evidence of clinical disease. However, cases have not been detected by intensive longitudinal studies in inter-epizootic periods in many epizootic areas, but have been identified in forest or forest edge situations, usually as human disease.
The use of sentinel cattle sited in known enzootic and epizootic areas for RVF is the recommended strategy for surveillance for RVF activity. The sera are collected 2-3 months after the seasonal rains, and may then be tested by ELISA for IgM or by other methods such as virus-serum neutralization assayed in a micro-titre system. It has been useful in areas where indigenous cattle predominate and the highly susceptible disease hosts are not available. This is the case in countries such as Mauritania, Senegal, Sudan and Ethiopia, where human disease is the main problem in RVF epizootic periods. Where the cattle are of susceptible breeds, RVF virus activity is likely to be clinically evident.
Predisposing climatic factors
Rainfall data may be used as a predictive tool for epizootic RVF, either by actual recordings or as a calculation from Cold Cloud Density (CCD) data derived from Remote Sensing Satellite Imagery (RSSD). The monthly analysis of this and Normalised Differentiated Vegetation Index (NDVI) data in the epizootic areas in eastern and parts of southern Africa are probably the most valuable tools for risk assessment of RVF virus activity. Currently FAO/EMPRES and some regional and national centres are doing this analysis on a monthly basis. The intention is to create an awareness and emergency preparedness for RVF. When the risk assessment is high, strategic preventive vaccination may be carried out and larvicidal and mosquito control strategies put into place. Whether the RSSD methodology will be applicable to the RVF problems in the Sahelian zones, has not yet been evaluated and sentinel cattle serology may currently be the best method.
The emergence of the floodwater-breeding mosquito species in epizootic areas is dependent upon the climatic factors, which are monitored by Remote Sensing Satellite Imagery. Longitudinal sampling of mosquitoes in such situations cannot be recommended as a useful surveillance activity for RVF virus activity. An observation of their emergence is obtained too late for it to drive any preventive measures.
RVF epizootics may be signalled by a sudden onset of many cases of a fatal haemorrhagic fever in man. This has a serious impact in small communities poorly served by communication and other services. Such episodes have been experienced in Mauritania and Senegal, Egypt, Kenya, Somalia and Ethiopia in the Horn of Africa. There is generally a period of 4-6 weeks before the cause of the problem is identified. No vaccine is available for use in these communities against RVF infection.
An epizootic of RVF affecting susceptible dairy cattle can result in abortions in up to 80% of pregnant animals with many (20-40%) deaths of calves of up to 3 months of age, 10-15% mortality may occur in animals of up to 1 year of age, with some deaths occurring in adults. The consequential losses from the abortions are considerable as milk production is drastically reduced, following the disruption of lactation patterns.
Camel herds suffer principally from abortions, when most of those pregnant may abort. This has a serious impact on the food security of the herders, for whom camel milk is major source of dietary protein.
The wool sheep, which have been imported particularly into South Africa and into Kenya and other exotic breeds introduced to improve production, are highly susceptible to RVF. The greatest impact may be seen in such flocks, where virtually all animals abort and all the young lambs die in a period of a few weeks. Many, but not all, of the indigenous sheep and goat breeds in RVF enzootic areas of sub-Saharan Africa show a degree of resistance to the virus and do not suffer in this manner. To the north however, in the Sahelian zones and in Egypt, most of the small ruminant breeds are susceptible to RVF.
The greatest overall loss produced by epizootics of RVF, results from the imposition of a total ban on livestock trade from RVF-infected areas. A recent RVF epizootic in the Horn of Africa was followed by the cessation of the lucrative trade in small ruminants to Arabic countries to the north and north-east. This resulted in serious economic losses of the order of 50-75 million dollars per year to the populations in the region, who were totally dependent upon this income. The first RVF epizootic in Egypt was said to have cost the livestock industry in Egypt some US $82 million in direct and indirect losses. The OIE regulations recommend the banning of livestock exports from an RVF infected country for 3 years following an outbreak.
A high risk of infection exists to all personnel associated with RVF-infected animals. These include the farmers and their staff, including those milking cattle or camels, veterinarians, and laboratory workers, abattoir staff, particularly those involved in the killing, skinning and cutting of fresh carcases. Those involved at later stages, who deal with the "set" meat as butchers and when cooking, do not become infected. As the virus is acid sensitive, the changes in pH associated with the setting process result in falls in the virus titre. The procedure of Halal, practised at the Muslim religious festivals and in homes, is particularly dangerous and has been an undoubted source of many cases of RVF during epizootics in such areas.
Great danger exists at the religious festivals at Mecca, for the ritual slaughter of a sheep is an integral part of the ceremony for many pilgrims. The sheep are imported to Mecca for this purpose in huge numbers (millions). The rapid transportation time from ports in the north of the Horn of Africa, allows of a very real risk that animals could be slaughtered at a viraemic stage and result in high levels of aerosol transmission of RVF amongst the very high populations that occur at Mecca at such times. No importation should ever be allowed when epizootic RVF virus activity is occurring in the Horn of Africa.
The virus does not persist in set meat carcases or frozen meat for any great length of time and no cases of RVF have been attributed to infection from such sources.
The mortality, which takes place in young animals, frequently occurs at the viraemic stage of the disease, and no specific lesions may be evident. The signs associated with viraemia, such as widespread petechial and ecchymotic haemorrhages on serous surfaces and organs will be seen, with extravasation of blood tinged fluids in the body cavities.
Virtually all older animals dying from RVF will show some degree of liver enlargement, inflammation and necrosis. This may be engorged with many foci of necrosis, 0.5 to 2.0 mm in diameter, bronzed with necrotic and engorged liver tissues which are jaundiced, then mottled yellow-brown or become completely yellow as jaundice becomes severe. The necrosis of the liver is not restricted to centri or peri-lobular areas but is a pan-necrosis, which affects all zones in a focal or a wider area. Early changes include cloudy swelling followed by hyaline degeneration. There is a massive necrosis with haemorrhage of the liver parenchyma with infiltration of lymphocytes, polymorphonuclear leukocytes and histiocytes, the original architecture of the liver is often unrecognisable. The gall bladder is often oedematous with many haemorrhages scattered throughout the tissues, and the mucosa may be necrotic and ulcerated. The carcase lymph nodes are enlarged, and oedematous, with superficial haemorrhages; the germinal centres of the follicles may show some necrosis.
The spleen may or may not be enlarged, but there are usually extensive subcapsular haemorrhages, with necrotic changes and cellular infiltration, mainly of polymorphonuclear cells. The kidneys show congestion with usually some petechial haemorrhages. There are degenerative changes in the tubular cells with varying degrees of cloudy swelling, necrosis and protein casts, which depend upon the stage at which death occurred. The heart usually shows sub-epicardial and endocardial haemorrhages. The lungs are hyperaemic with sub-pleural and other haemorrhages, often a mild interstitial pneumonia and on occasion emphysema and oedema. A haemorrhagic gastro-enteritis is commonly encountered, with foci of haemorrhagic necrosis and extravasation of blood into the mucosa and muscular tissues. The serous surface of the bowels may be covered with petechial and ecchymotic haemorrhages. Frank haemorrhage occurs into the lumen of the bowel with some necrosis of the mucosa.
In acute cases, the visible mucous membranes of the mouth and vulva will be cyanotic and haemorrhages may be visible in hairless areas such as the axilla and groin, and also the udder and scrotum. Necrotic changes may also be seen in these areas. There is a subcutaneous congestion and sometimes oedema.
Encephalitis has not been reported in the epizootics of RVF, which occur in domestic animals, it may however occur in a small proportion of animals and be overlooked. In experiments, it manifests as abnormalities of movement, feeding and posture, with other neurological signs. These signs also follow intra-cerebral inoculation with the neuro-adapted strains. It is commonly seen as a serious complication of human cases in epizootics.
Foetuses may be aborted at all stages of pregnancy and virus may be isolated from the placenta, foetal liver and other tissues. An unusual syndrome has been observed following the vaccination of ewes at an early stage of pregnancy with the Smithburn neurotropic strain (SNS) vaccine. This produces an hydranencephaly/arthrogryopsis syndrome in 10-15% of those vaccinated and the ewes show hydrops amnii and usually have a prolonged period of gestation. Late vaccination may result in neurological disease in the lambs and the virus may be re-isolated from the brain. It is for these reasons that the SNS vaccine is recommended for use only in non-pregnant animals.
The clinical impact of an RVF epizootic, with its sudden onset, acute prostrating disease in man and abortion and neonatal deaths in domestic animals should prompt a strong suspicion of RVF. Its occurrence in a country for the first time has invariably been followed by delays in making a confirmatory diagnosis. RVF virus belongs to the Group of P-4 pathogens, which require high security virological laboratory facilities to work with the virus.
Virus isolation and identification
The old classical methods of virus isolation in suckling mice and hamsters , where deaths occur within 56-72 h may be used if tissue cultures are not available. Their livers and spleens may then be used for the identification of the antigen in agar gel precipitation tests, in antigen capture ELISA tests, as antigen for complement fixation, haemagglutination, fluorescent antibody or other test system. Antigen capture ELISA systems may be used with the liver and spleen tissue, which usually contain high titres of virus.
The inoculation of tissue cultures with viraemic blood, spleen or liver homogenates allows a specific diagnosis by fluorescent antibody or immuno-peroxidase staining methods in 36–72 h, when viral antigen can be identified in the cytoplasm of infected cells. The primary inoculation and identification in tissue culture, with baby hamster kidney (BHK), Vero or Aedes pseudoscutellaris cell lines is a sensitive, rapid and safe procedure which is recommended for RVF diagnosis.
Specific identification of the isolate to distinguish it from other viruses can be made in a virus serum neutralization test by a microplate or plaque assay. This is necessary to distinguish the isolate from other Phleboviruses. Sandfly fever, however, which is a possible Phlebovirus isolate, is not associated with disease in domestic animals.
In areas where communications are poor, liver samples may be transported in formol-saline over long distances without deterioration. These can then be processed by an immuno-staining method using avidin-biotin or other method to demonstrate RVF antigen in the liver tissue. This avoids the need for P-4 Biohazard facilities and may be used in any laboratory. Capture ELISA using monoclonals and polymerase chain reaction (PCR) methodology has now been developed and are both are available for the diagnosis of RVF. The capture ELISA systems do have a role to play in countries without virological facilities or laboratories, but where ELISA testing is established for other diseases.
The serological tools, which have been used for RVF for many years, have now been largely replaced by ELISA systems. A neutralisation test in mice was used extensively 50 years ago and more recently plaque or micro-titre tissue culture assays, for the identification of isolates, epidemiological and other studies. The availability of ELISA test systems, which detect RVF IgM and IgG have now largely replaced the previous methods for RVF investigations. The presence of IgM in large numbers of animals, which have recently aborted or shown clinical signs supports a presumptive diagnosis of epizootic RVF virus activity. However, IgM antibody persists for up to 9 months after an infection.
Assay of RVF
The virus may be assayed in laboratory mice and hamsters, and these systems were extensively used until cell cultures became available. Laboratory mice were inoculated either by intra-cerebral (ic) or intra-peritoneal (ip) routes. Suckling mice would be used for the ic and 4-6 week old mice for ip inoculations. Hamsters of 4-8 weeks of age are highly susceptible to RVF virus and may be inoculated by the ip route. Virus titres from infected viraemic animals may be 106.5 to 9.5 LD50's per 0.1 ml of blood in these assays. Note that P-4 facilities are required in the laboratory to handle RVF virus. However, vaccinated personnel may carry out RVF work in Class III Biohazard Units, with great care in their technique and disposal and sterilization procedures.
Tissue culture systems are now used extensively for assay of RVF virus, and those commonly employed are BHK 21 C13, Vero, L (* mouse fibroblast), foetal rhesus monkey, Aedes pseudoscutellaris , primary/secondary ovine or bovine kidney, testis or muscle cells. Assays may be made by titrations in bottles, tubes or microtitre systems, and as plaque assays under agar overlays. The titres obtained in Vero or BKK cells are likely to be at least 1 log10 less than those obtained in laboratory animals, such as hamsters or mice.
RVF in man presents as a sudden onset of fever, usually of a biphasic character, severe myalgia, headaches with retro-orbital pain; colic, vomiting and diarrhoea may also occur at this stage. The initial febrile period is of 2-3 days and the second after a remission of 1-2 days. This may last for 3-7 days and then there is a complete recovery in the majority of cases. Many of these cases may also show signs of hepatitis and recovery is usually accompanied by some degree of debility, which may persist for several months. More severe complications develop in 1-4% of cases, but the figure varies from outbreak to outbreak and may reflect the inter-current nutritional and disease status of the population at risk. There was a suggestion that the mortality in the first Egyptian epizootic of RVF was higher than had ever been encountered in Sub-Saharan Africa. This may have reflected inter-current liver or other chronic disease (such as schistosomiasis), and a greater susceptibility appears to exist in areas hyper-endemic for malaria, especially in semi-arid zones.
The complications are evident as a severe hepatitis with jaundice, which may be followed by a haemorrhagic syndrome. Many of these cases are severely ill and fatalities are common in the severe haemorrhagic cases. This occurs fairly early in the course of the disease. The complications, which develop 1-4 weeks later, are of retinal damage and encephalitis. These manifest as progressive degrees of blindness and various signs of encephalitis, including paralysis of one or more limbs.
In domestic animals
The incubation period is short, usually 1-3 days, when a febrile reaction occurs of 40.5 to 41.5ºC. Young animals experience a peracute form of the disease, when anorexia and listlessness is closely followed by collapse and death. In older lambs and adults the febrile reaction is accompanied by severe depression, anorexia, raised respiratory and heart rate, occasionally colic, and vomiting (which is rare in ruminants), a sero-sanguineous and later muco-purulent discharge from the nostrils (which may also be stained with ruminal contents), haemorrhagic gastro-enteritis, followed by recumbency and often death. The mortality is from 10-60% in these cases, according to the age structure of the flock. In less susceptible age groups and breeds, a necrotizing encephalitis may develop in a few animals 2-3 weeks after the period when acute infections were occurring. Neurological disease is manifested by changes in posture and gait. Affected herds and flocks will usually show ill thrift associated with the severe liver damage for several months following recovery. Photosensitization has been observed as a complication in such animals, a consequence of the hepatitis.
Abortion in pregnant sheep, goats or cattle is the most obvious presenting sign in a flock or herd. Often 60-80% of animals at all stages of pregnancy will abort over 2-3 weeks. RVF produces an acute inflammation of the maternal caruncles, which results in necrosis of the villi and cotyledons. Foetal death is a direct consequence of these lesions.
Morbidity in many epizootics has been found to be extremely high with 85-95% of all animals becoming affected in a relatively short time. Indigenous breeds of cattle or sheep held in close association, with those which are showing clinical signs, may exhibit no evidence of RVF. Serology would, however, show that most had seroconverted to RVF. In dairy herds the febrile reaction is accompanied by a fall in milk yield, some depression, anorexia with salivation, a foetid diarrhoea which is often blood stained, and sero-sanguineous nasal discharges. Many of the superficial lymph nodes will be enlarged. A stomatitis with erosions and haemorrhage and necrotic changes of the hairless areas of the skin may also be seen. Abortion may occur at this stage or up to 10 days later.
Other disease hosts
Camel herds experience abortion storms during periods of epizootic RVF virus activity. Adult animals do not show any clinical signs of RVF and experimental inoculation has shown only brief periods of viraemia. There are camel deaths amongst neonatal foals during these epizootic periods, which may be due to RVF infection. The horse develops a transient viraemia with no clinical signs. The Asiatic buffalo (Bubalis bubalis) was challenged by RVF virus during the first Egyptian outbreak, 20% were seropositive and there were reports of some abortions. Apart from this they showed no obvious clinical disease. The African buffalo (Syncerus caffer) has been shown to develop viraemia after inoculation with RVF virus, and one pregnant animal was observed to abort. No clinical disease problem has been identified in this species in the field however. Many wild ruminant species have shown evidence of RVF infection by serology after epizootics. Neither disease nor mortality have been observed in wild game animals during epizootic periods, when cattle and sheep, which share the same grazing areas were sick and dying.
Dogs and cats develop viraemia after experimental inoculations but are not thought to be other than accidental hosts in epizootics. Primates become viraemic and develop antibody to RVF, the rhesus monkey shows a moderately severe reaction with 25% mortality in one study. The African green monkey (Cercopithecus aethiops) and baboons (Papio anubis) show fever with some clinical malaise for 1-2 days after experimental inoculation with RVF, but very few have been shown to have had natural infections. Most rodent species are susceptible to RVF but no obvious mortality has been seen in wild rodent populations during RVF epizootics. Antibody studies show that only a small proportion (less than 2-12%) of the populations are exposed to RVF in epizootics and they are not thought to play any significant role as reservoir or amplification hosts. Birds are insusceptible and whilst antibody has been found in bats, the significance of this is probably not important. Bats and rodents may be more important in the tropical forest and forest edge situations where ruminants are not present.
Ribavirin treatment at an early stage of the disease has been shown to modify the course of the disease in experiments with primates (rhesus monkeys). The administration of RVF immune globulins has a protective effect, if given very early in the course of the disease. Alpha interferon has also been tested in primates. No trials have been reported in human cases in the field.
Larvicides provide a most promising area for control in certain of the RVF epizootic areas, where the Aedes mosquito breeding sites can be clearly defined, and are of limited extent. This is the case in parts of eastern and southern Africa and in Mauritania and Senegal. In the latter for example, clay deposits have formed pans in the sandy Sahel, which fill regularly if there is sufficient rainfall. These can be treated with slow release preparations of larvicides, either before or when they have flooded. Both methoprene, a hormonal larval growth inhibitor and Bacillus thurigiensis slow release preparations are available commercially and pilot studies have shown that they can be extremely effective. They could be applied by community health workers to the local flood-water zones after suitable instruction and extension inputs.
Insecticides have been used during periods of epizootic RVF virus activity. Thermal fogging and ground application by spraying have both given good results, when used in the Egyptian epizootics. They are not readily applicable to the situation in most of the RVF epizootic areas in Africa.
Immunization and vaccines
A modified live virus vaccine has been developed from the Smithburn neurotropic strain (SNS) of RVF, which has lost the viscerotropic characteristics of the virus following serial passage in suckling mouse brain. It does retain some foeto-tropic characteristics, however, and is not recommended for use in pregnant sheep of susceptible breeds. The vaccine may be prepared in roller cultures of Vero or BHK cells and grown to a high titre. It is extremely cheap to prepare, is immunogenic for sheep, goats and cattle, and protects against abortion. The vaccine has been used in the face of an epizootic in pregnant cattle of highly susceptible breeds with no complications. In sheep however, some 5-15% of pregnant animals may show foetal abnormalities and/or abortion. Most farmers prefer to take this risk rather than suffer the 90% abortion and the high death rates caused by the epizootics of RVF, others vaccine routinely after lambing. Several million doses were used in Egyptian animals without complications. The immunity is lifelong, but revaccination is often carried out after 2-3 years. Despite its disadvantages, this remains the most widely used vaccine against RVF in Africa.
Inactivated RVF vaccines have been prepared in South Africa and Egypt, where they have been widely and successfully used, especially in cattle. These prevent abortion in laboratory studies and in the field, but other evidence has accrued during epizootics in Egypt and Africa, which suggests that the level of protection has been inadequate in these situations. Animals that had been given two inoculations with inactivated vaccines still aborted in the epizootic situations. These vaccines are expensive and require at least two inoculations to produce the desired level of protection. This is often not a cost effective strategy nor possible in many epizootic situations. A modified live strain is more suitable.
A mutagen has been used to induce a change in a strain of RVF (MP 12), which is of greatly reduced virulence. Experiments show that this is protective against disease and abortion after needle challenge and does not have the foeto-tropic disadvantage of the Smithburn strain. Laboratory development has proceeded to the point, where extensive field trials are required. Further development of the strains has, however, been slow and it has still not been adequately tested in the field.
An experimental vaccine has been prepared for use in laboratory personnel to protect them against RVF. This is a formalinised tissue culture vaccine, and 3 inoculations result in the development of high titres of neutralising antibody to RVF virus in most people. The vaccine has been extensively used in personnel handling RVF virus in many countries, it generates a good antibody response after 2-3 inoculations with protection against laboratory infection. No vaccine is available for widespread use and the protection of human populations at risk from RVF during epizootics.
African Union-Interafrican Bureau for Animal Resources, 2011. Panafrican Animal Health Yearbook 2011. Pan African Animal Health Yearbook, 2011:xiii + 90 pp. http://www.au-ibar.org/pan-african-animal-health-yearbook
Alexander RA, 1951. Rift Valley fever in the Union. Journal of the South African Veterinary Medical Association, 22:105-112.
Ali AM, Kamel S, 1978. Epidemiology of RVF in domestic animals in Egypt. Journal of the Egyptian Public Health Association, 53:255-263.
Anderson GW, Slone TW, Peters CJ, 1987. Pathogenesis of Rift Valley fever virus in in-bred rats. Microbial Pathogenesis, 2:283-293.
Anderson GWJr, Saluzzo JF, Ksiazek TG, Smith JF, Ennis W, Thureen D, Peters CJ, Digoutte JP, 1989. Comparison of in vitro and in vivo systems for propagation of Rift Valley fever virus from clinical specimens. Research in Virology, 140(2):129-138; 11 ref.
Arborio M, Hall WC, 1989. Diagnosis of a human case of Rift Valley fever by immunoperoxidase demonstration of antigen in fixed liver tissue. Research in Virology, 140(2):165-168; 8 ref.
Art D, André P, 1991. Clinical and epidemiological aspects of listeriosis in Belgium 1985-1990. Zentralblatt für Bakteriologie, 275(4):549-556; 23 ref.
Barnard BJH, 1979. Rift Valley fever vaccine antibody and immune response to in cattle to a live and an inactivated vaccine. Journal of the South African Veterinary Association, 50:155-157.
Barnard BJH, Botha MJ, 1977. An inactivated Rift Valley fever vaccine. Journal of the South African Veterinary Association, 48:45-48.
Baskerville A, Hubbard KA, Stephenson JR, 1992. Comparison of the pathogenicity for pregnant sheep of Rift Valley fever virus and a live attenuated vaccine. Research in Veterinary Science, 52(3):307-311; 12 ref.
Battles JK, Dalrymple JM, 1988. Genetic variation among geographic isolates of Rift Valley fever virus. American Journal of Tropical Medicine and Hygiene, 39(6):617-631; 32 ref.
Brubaker JF, Turell MJ, 1998. Effect of environmental temperature on the susceptibility of Culex pipiens (Diptera: Culicidae) to Rift Valley fever virus. Journal of Medical Entomology, 35(6):918-921; 21 ref.
Capstick PB, Gosden D, 1962. Neutralising antibody response in sheep to pantropic and neurotropic Rift Valley fever virus. Nature, 195:583-4.
Cash P, Robeson G, Erlich BJ, Bishop DHL, 1981. Biochemical characterisation of Rift Valley fever and other Phlebotomus fever group viruses. In: Swartz TA, Klingberg MA, Godblum N, ed. Rift Valley fever. Contributions to Epidemiology and Statistics. Vol 3, Basel, Switzerland: S. Karger, 1-20.
Chartier C, Chartier F, 1988. Sero-epidemiological survey of infectious abortion of small ruminants in Mauritania. Revue d'élevage et de Médecine Vétérinaire des Pays Tropicaux, 41(1):23-34; 73 ref.
Coakley W, 1963. The effect of recently isolated strains of Rift Valley fever virus on lamb testis cell cultures. J. Path. Bact., 86:530-532.
Coakley W, 1965. Alteration in virulence of Rift Valley fever virus during serial passage in lamb testis cells. J. Path. Bact., 89:123-131.
Coakley W, Pini A, Gosden D, 1967. Experimental infection of cattle with Pantropic Rift Valley fever virus. Research in Veterinary Science, 8:399-405.
Coakley W, Pini A, Gosden D, 1967. The immunity induced in cattle and sheep by inoculation of neurotropic or pantropic Rift Valley fever viruses. Research in Veterinary Science, 8:406-414.
Coetzer JAW, 1977. The pathology of Rift Valley fever. Lesions occurring in natural cases in new born lambs. Onderstepooort Journal of Veterinary Research, 44:205-212.
Coetzer JAW, Barnard BJH, 1977. Hydrops amnii in sheep associated with hydranencephaly and arthrogryposis, with Wesselbron and Rift Valley fever viruses as aetiological agents. Onderstepoort Journal of Veterinary Research, 44:119-126.
Dar O, McIntyre S, Hogarth S, Heymann D, 2013. Rift Valley fever and a new paradigm of research and development for zoonotic disease control. Emerging Infectious Diseases, 19(2):189-193. http://wwwnc.cdc.gov/eid/article/19/2/pdfs/12-0941.pdf
Darwish MA, Imam IZE, Omar F, 1978. A sero-epidemiological study for Rift Valley fever virus in humans and domestic animals in Egypt. Journal of the Egyptian Public Health Association, 53:153-162.
Daubney R, Hudson JR, 1932. Rift Valley fever. Lancet, 1:611-612.
Daubney R, Hudson JR, Garnham PC, 1931. Enzootic hepatitis or Rift Valley fever. An undescribed virus disease of sheep, cattle and man from East Africa. J. Path. Bact., 34:545-579.
Davies FG, 1975. Observations on the epidemiology of Rift Valley fever in Kenya. Journal of Hygiene, 75:219-230.
Davies FG, 1990. Rift Valley fever in the Sudan. Transactions of the Royal Society of Tropical Medicine and Hygiene, 84(1):141; 6 ref.
Davies FG, Adday PAK, 1979. Rift Valley fever: a survey for antibody to the virus in bird species commonly found in situations considered to be enzootic. Transactions of the Royal Society of Tropical Medicine and Hygiene, 72:213-214.
Davies FG, Clausen B, Lund LJ, 1972. The pathogenicity of Rift Valley fever virus for the baboon. Transactions of the Royal Society of Tropical Medicine and Hygiene, 66:363-365.
Davies FG, Highton RB, 1980. Possible vectors of Rift Valley fever in Kenya. Transactions of the Royal Society of Tropical Medicine and Hygiene, 74:815-816.
Davies FG, Karstad L, 1981. Experimental infection of the African buffalo (Syncerus caffer) with the virus of Rift Valley fever. Tropical Animal Health Production, 13:185-188.
Davies FG, Kilelu E, Linthicum KJ, Pegram RG, 1992. Patterns of Rift Valley fever activity in Zambia. Epidemiology and Infection, 108(1):185-191; 15 ref.
Davies FG, Koros J, Mbugua H, 1985. Rift Valley fever in Kenya: the presence of antibody to the virus in camels (Camelus dromedarius). Journal of Hygiene, 94(2):241-244; 11 ref.
Davies FG, Linthicum KJ, 1986. The Sudan dioch (Quelea quelea aethiopica) and Rift Valley fever. Transactions of the Royal Society of Tropical Medicine and Hygiene, 80(1):171-172; 6 ref.
Davies FG, Linthicum KJ, James AD, 1985. Rainfall and epizootic Rift Valley fever. Bulletin of the World Health Organization, 63(5):941-943; 20 ref.
Davies FG, Logan TM, Binepal Y, Jessen P, 1992. Rift Valley fever virus activity in East Africa in 1989. Veterinary Record, 130(12):247-248; 11 ref.
Dell'Aquila S, Pilla AM, Catillo G, Scardella P, Taibi L, 1993. Milk yield in dairy sheep of the Comisana (C), Langhe (L), Massese (M) and Sardinian (S) breeds, and crosses L x C, L x M and L x S: second lactation. Zootecnica e Nutrizione Animale, 19(2):95-102; 13 ref.
Digoutte JP, Jouan A, Le Guenno B, Riou O, Philippe B, Meegan J, Ksiazek TG, Peters CJ, 1989. Isolation of the Rift Valley fever virus by inoculation into Aedes pseudoscutellaris cells: comparison with other diagnostic methods. Research in Virology, 140(1):31-41; 31 ref.
Easterday BC, 1965. Rift Valley fever. Advances in Veterinary Science, 10:65-127.
Easterday BC, McGavran MH, Rooney JR, Murphy LC, 1962. The pathogenesis of Rift Valley fever in lambs. American Journal of Veterinary Research 23:470-479.
Easterday BC, Murphy LC, 1963. Studies on Rift Valley fever in laboratory animals. Cornell Veterinarian, 53:423-433.
Eisa M, Kheir El Sid ED, Shomei AM, Meegan JM, 1980. An outbreak of Rift Valley fever in the Sudan - 1976. Transactions of the Royal Society of Tropical Medicine and Hygiene, 745:417-418.
Eisa M, Obeid HMA, El Sawi ASA, 1977. Rift Valley fever in the Sudan. Bulletin of Animal Health and Production in Africa, 25:343-355.
El Akkad AM, 1978. Rift Valley fever outbreak in Egypt, October-December 1977. Journal of the Egyptian Public Health Association, 53:123-128.
El-Rahim IHAA, El-Hakim UA, Hussein M, 1999. An epizootic of Rift Valley fever in Egypt in 1997. Revue Scientifique et Technique - Office International des épizooties, 18(3):741-748; 41 ref.
Emmens RL, Murray MD, 1983. Bacterial odours as oviposition stimulants for Lucilia cuprina (Wiedemann) (Diptera: Calliphoridae), the Australian sheep blowfly. Bulletin of Entomological Research, 73(3):411-415; 9 ref.
Fagbami AH, Tomori O, Fabiyi A, Isoun TT, 1975. Experimental Rift Valley fever in West African dwarf sheep. Research in Veterinary Science, 18:334-335.
Fontenille D, Traore-Lamizana M, Diallo M, Thonnon J, Digoutte JP, Zeller HG, 1998. New vectors of Rift Valley fever in West Africa. Emerging Infectious Diseases, 4(2):289-293; 17 ref.
Fontenille D, Traore-Lamizana M, Zeller H, Mondo M, Diallo M, Digoutte JP, 1995. Short report: Rift Valley fever in western Africa: isolations from Aedes mosquitoes during an interepizootic period. American Journal of Tropical Medicine and Hygiene, 52(5):403-404; 5 ref.
Gear J, de Meillon B, Le Roux AF, Kofsky BA, 1955. Rift Valley fever in South Africa. A study of the 1953 outbreak in the Orange Free State, with special reference to the vectors and possible reservoir hosts. South African Medical Journal, 29:514-518.
Gerdes GH, 2004. Rift Valley fever. Revue Scientifique et Technique - Office International des Épizooties, 23(2):613-623.
Gonzalez JP, Guenno Ble, Some MJR, Akakpo JA, 1992. Serological evidence in sheep suggesting phlebovirus circulation in a Rift Valley fever enzootic area in Burkina Faso. Transactions of the Royal Society of Tropical Medicine and Hygiene, 86(6):680-682; 18 ref.
Hatheway CL, 1990. Toxigenic clostridia. Clinical Microbiology Reviews, 3(1):66-98; 355 ref.
Henderson BE, McCrae AWR, Kirya BG, Ssenkubuge Y, Sempala SDK, 1972. Arbovirus epizootics involving man, mosquitoes and vertebrates at Lunyo, Uganda, 1968. Annals of Tropical Medicine and Parasitology, 66:343-355.
Hoogstraal H, Meegan JM, Khalil GM, Adham FK, 1979. The Rift Valley fever epizootic in Egypt 1977-1978. Ecological and entomological studies. Transactions of the Royal Society of Tropical Medicine and Hygiene, 73:624-629.
Imam IZE, El Karamany R, Darwish MA, 1979. An epidemic of Rift valley fever in Egypt. Isolation of the virus from animals. Bulletin of the World Health Organization, 57:441-443.
Joshi MV, Umarani UB, Pinto BD, Joshi GD, Paranjape SP, Banerjee K, 1995. Prevalence of antibodies to Rift Valley fever (or closely related) virus in sheep and goats from Rajasthan, India: a preliminary report. Indian Journal of Virology, 11(1):59-60; 7 ref.
Kitchen SF, 1950. The development of neurotropism in Rift valley fever virus. Annals of Tropical Medicine and Parasitology, 44:132-145.
LaBeaud AD, Cross PC, Getz WM, Glinka A, King CH, 2011. Rift Valley fever virus infection in African buffalo (Syncerus caffer) herds in rural South Africa: evidence of interepidemic transmission. American Journal of Tropical Medicine and Hygiene, 84(4):641-646. http://www.ajtmh.org
Lecatsas G, Weiss KW, 1968. Electron microscopic studies on BHK-21 cells infected with Rift Valley fever virus. Arch. Ges. Virusforshung., 25:58-64.
Linthicum KJ, Anyamba A, Tucker CJ, Kelley PW, Myers MF, Peters CJ, 1999. Climate and satellite indicators to forecast Rift Valley fever epidemics in Kenya. Science (Washington), 285(5426):397-400; 21 ref.
Linthicum KJ, Bailey CL, Davies FG, Tucker CJ, 1987. Detection of Rift Valley fever viral activity in Kenya by satellite remote sensing imagery. Science, USA, 235(4796):1656-1659; 17 ref.
Linthicum KJ, Bailey CL, Tucker CJ, Angleberger DR, Cannon T, Logan TM, Gibbs PH, Nickeson J, 1991. Towards real-time prediction of Rift Valley fever epidemics in Africa. Preventive Veterinary Medicine, 11(3-4):325-334; 12 ref.
Linthicum KJ, Bailey CL, Tucker CJ, Mitchell KD, Logan TM, Davies FG, Kamau CW, Thande PC, Wagateh JN, 1990. Application of polar-orbiting, meteorological satellite data to detect flooding of Rift Valley fever virus vector mosquito habitats in Kenya. Medical and Veterinary Entomology, 4(4):433-438; 9 ref.
Linthicum KJ, Davies FG, Kairo A, 1985. Rift Valley fever virus (family Bunyaviridae, genus Phlebovirus). Isolations from Diptera collected during an inter-epizootic period in Kenya. Journal of Hygiene, 95(1):197-209; 39 ref.
Linthicum KJ, Logan TM, Bailey CL, Dohm DJ, Moulton JR, 1989. Transstadial and horizontal transmission of Rift Valley fever virus in Hyalomma truncatum.. American Journal of Tropical Medicine and Hygiene, 41(4):491-496; 23 ref.
Logan TM, Linthicum KJ, Davies FG, Binepal YS, Roberts CR, 1991. Isolation of Rift Valley fever virus from mosquitoes (Diptera: Culicidae) collected during an outbreak in domestic animals in Kenya. Journal of Medical Entomology, 28(2):293-295; 16 ref.
Logan TM, Linthicum KJ, Wagateh JN, Thande PC, Kamau CW, Roberts CR, 1990. Pretreatment of floodwater Aedes habitats (dambos) in Kenya with a sustained release formulation of methoprene. Journal of the American Mosquito Control Association, 6:736-738.
MacGavran MH, Easterday BC, 1963. Rift Valley fever hepatitis. Light and electron microscopic studies in the mouse. American Journal of Pathology, 42:587-607.
Mackenzie RD, Findlay GM, 1936. The production of neurotropic variants of Rift Valley fever virus. Lancet, 1:140-144.
Manohar BM, Sundararaj A, Sheela PRR, Elankumaran S, Albert A, Venugopalan AT, 1995. A report on an outbreak of Rift Valley fever-like disease in sheep in Tamil Nadu. Indian Veterinary Journal, 72(6):662-664; 3 ref.
Mariner JC, Morrill J, Ksiazek TG, 1995. Antibodies to hemorrhagic fever viruses in domestic livestock in Niger: Rift Valley fever and Crimean-Congo hemorrhagic fever. American Journal of Tropical Medicine and Hygiene, 53(3):217-221; 31 ref.
Maurice Y, 1967. Premieres constatations serologiques sur l'incidence de la maladie de Wesselbron et de la fievre de la Vallee du Rift chez les ovins et les ruminants sauvages du Tchad et du Cameroun. Revue d'Elevage et de Médecine Vétérinaire des Pays Tropicaux, 20:359-405.
Mazanowski A, Bernacki Z, 1998. Evaluation of meat traits of intensively reared crossbred geese from genetic reserve flocks compared with White Koluda geese. Roczniki Naukowe Zootechniki, 25(4):159-174; 16 ref.
Mazanowski A, Smalec E, 1998. Rearing performance of 12-week-old crossbreds of ganders and geese from genetic reserve flocks compared with White Koluda. Roczniki Naukowe Zootechniki, 25(4):191-205; 17 ref.
McIntosh BM, 1972. Rift Valley fever. Vector studies in the field. Journal of the South African Veterinary Association, 43:391-395.
McIntosh BM, Jupp PG, 1981. Epidemiological aspects of Rift Valley fever in South Africa, with reference to vectors. Basel, Switzerland: Karger. Contr. Epid. Biostatist., 3:92-99.
McIntosh BM, Jupp PG, Anderson D, Dickenson DB, 1973. Rift Valley fever: attempts to transmit the virus with seven species of mosquito. Journal of the South African Veterinary Association, 44:57-60.
McIntosh BM, Jupp PG, Dos Santos I, Barnard JH, 1980. Vector studies on Rift Valley fever virus in South Africa. South African Medical Journal, 58:127-132.
Meegan JM, Bailey CL, Monath TP, 1989. Rift Valley fever. The arboviruses:-epidemiology and ecology. Boca Raton, Florida; USA: CRC Press, Inc., IV:51-76.
Meegan JM, Khalil GM, Hoogstraal H, Adham FK, 1980. Experimental transmission and field isolation studies implicating Culex pipiens as a vector of Rift valley fever virus in Egypt. American Journal of Tropical Medicine and Hygiene, 29:1405-1410.
Meegan JM, Yedloutschnig RJ, Peleg BA, Shy J, Peters CJ, Walker JS, Shope RE, 1987. Enzyme-linked immunosorbent assay for detection of antibodies to Rift Valley fever virus in ovine and bovine sera. American Journal of Veterinary Research, 48(7):1138-1141; 13 ref.
Metselaar D, Henderson BE, Kirya J, Tukei MP, de Geus A, 1974. Isolation of arboviruses in Kenya, 1966-1971. Transactions of the Royal Society of Tropical Medicine and Hygiene, 68:114-123.
Mims CA, 1956. Absorption and multiplication of the virus. British Journal of Experimental Pathology, 37:110-119 & 37:120-128 & 37, Incomplete virus, 37:129-143.
Mims CA, 1956. Rift Valley fever in mice. General features of the infection. British Journal of Experimental Pathology, 37:99-109.
Mims CA, 1957. Rift Valley fever in mice. The histological changes in the liver in relation to virus multiplication. Australian Journal of Experimental Biology and Medical Science, 35:595-604.
Molinuevo HA, 1995. Productivity of Criollo x Aberdeen-Angus back-crossed cattle. Revista Argentina de Producción Animal, 15(3/4):909-911; 9 ref.
Morrill JC, Carpenter L, Taylor D, Ramsburg HH, Quance J, Peters CJ, 1991. Further evaluation of a mutagen-attenuated Rift Valley fever vaccine in sheep. Vaccine, 9(1):35-41; 20 ref.
Morrill JC, Czarniecki CW, Peters CJ, 1991. Recombinant human interferon- modulates Rift Valley fever virus infection in the Rhesus monkey. Journal of Interferon Research, 11(5):297-304; 31 ref.
Morrill JC, Jennings GB, Johnson AJ, Cosgriff TM, Gibbs PH, Peters CJ, 1990. Pathogenesis of Rift Valley fever in rhesus monkeys: role of interferon response. Archives of Virology, 110(3-4):195-212; 34 ref.
Morrill JC, Knauert FK, Ksiazek TG, Meegan JM, Peters CJ, 1989. Rift Valley fever infection of rhesus monkeys: implications for rapid diagnosis of human disease. Research in Virology, 140(2):139-146; 12 ref.
Morrill JC, Mebus CA, Peters CJ, 1997. Safety and efficacy of a mutagen-attenuated Rift Valley fever virus vaccine in cattle. American Journal of Veterinary Research, 58(10):1104-1109; 23 ref.
Morvan J, Saluzzo JF, Fontenille D, Rollin PE, Coulanges P, 1991. Rift Valley fever on the east coast of Madagascar. Research in Virology, 142(6):475-482; 25 ref.
Muller R, Saluzzo JF, Lopez N, Dreier T, Turell M, Smith J, Bouloy M, 1995. Characterization of clone 13, a naturally attenuated avirulent isolate of Rift Valley fever virus, which is altered in the small segment. American Journal of Tropical Medicine and Hygiene, 53(4):405-411; 33 ref.
OIE Handistatus, 2002. World Animal Health Publication and Handistatus II (dataset for 2001). Paris, France: Office International des Epizooties.
OIE Handistatus, 2003. World Animal Health Publication and Handistatus II (dataset for 2002). Paris, France: Office International des Epizooties.
OIE Handistatus, 2004. World Animal Health Publication and Handistatus II (data set for 2003). Paris, France: Office International des Epizooties.
OIE, 2004. Rift Valley fever in Saudi Arabia. Serological findings (follow-up report No. 1: end of the outbreak). Disease Information, 17(49).
OIE, 2005. World Animal Health Publication and Handistatus II (data set for 2004). Paris, France: Office International des Epizooties.
OIE, 2009. World Animal Health Information Database - Version: 1.4. World Animal Health Information Database. Paris, France: World Organisation for Animal Health. http://www.oie.int
OIE, 2012. World Animal Health Information Database. Version 2. World Animal Health Information Database. Paris, France: World Organisation for Animal Health. http://www.oie.int/wahis_2/public/wahid.php/Wahidhome/Home
Paweska JT, Mortimer E, Leman PA, Swanepoel R, 2005. An inhibition enzyme-linked immunosorbent assay for the detection of antibody to Rift Valley fever virus in humans, domestic and wild ruminants. Journal of Virological Methods, 127(1):10-18. http://www.sciencedirect.com/science/journal01660934
Pini A, Lund LJ, Davies FG, 1970. Detection of Rift Valley fever virus by the fluorescent antibody technique in organs of experimentally infected animals. Research in Veterinary Science, 11:82-85.
Pittman PR, Liu CT, Cannon TL, Makuch RS, Mangiafico JA, Gibbs PH, Peters CJ, 1999. Immunogenicity of an inactivated Rift Valley fever vaccine in humans: a 12-year experience. Vaccine, 18(1/2):181-189; 46 ref.
Prehaud C, Lopez N, Blok MJ, Obry V, Bouloy M, 1997. Analysis of the 3 terminal sequence recognized by the Rift Valley fever virus transcription complex in its ambisense S segment. Virology (New York), 227(1):189-197; 51 ref.
Pretorius A, Oelofsen MJ, Smith MS, Ryst Evan der, 1997. Rift Valley fever virus: a seroepidemiologic study of small terrestrial vertebrates in South Africa. American Journal of Tropical Medicine and Hygiene, 57(6):693-698; 24 ref.
Randall R, Binn LN, Harrison VR, 1964. Immunisation against Rift Valley fever virus. Studies on the immunogenicity of lyophilised formalin inactivated vaccine. Journal of Immunology, 92:293-299.
Rice RM, Erlick BJ, Rosato RR, Eddy GA, Mohanty SB, 1980. Biochemical characterisation of Rift Valley fever virus. Virology, 105:256-260.
Rippy MK, Topper MJ, Mebus CA, Morrill JC, 1992. Rift Valley fever virus-induced encephalomyelitis and hepatitis in calves. Veterinary Pathology, 29(6):495-502; 18 ref.
Rossi CA, Turell MJ, 1988. Characterization of attenuated strains of Rift Valley fever virus. Journal of General Virology, 69(4):817-823; 13 ref.
Sall AA, Zanotto PMde A, Zeller HG, Digoutte JP, Thiongane Y, Bouloy M, 1997. Variability of the NS protein among Rift Valley fever virus isolates. Journal of General Virology, 78(11):2853-2858; 40 ref.
Saluzzo JF, Chartier C, Bada R, Martinez D, Digoutte JP, 1987. Rift Valley fever in West Africa. Revue d'élevage et de Médecine Vétérinaire des Pays Tropicaux, 40(3):215-223; 27 ref.
Samany R, 1997. Rift Valley fever surveillance in Madagascar. Bulletin - Office International des épizooties, 109(1):43-44.
Samui KL, Inoue S, Mweene AS, Nambota AM, Mlangwa JED, Chilonda P, Onuma M, Morita C, 1997. Distribution of Rift Valley fever among cattle in Zambia. Japanese Journal of Medical Science & Biology, 50(2):73-77; 4 ref.
Scott GR, 1963. Pigs and Rift Valley fever. Nature, 200:920-921.
Scott GR, Coakley W, Roach RW, Cowdy NR, 1963. Rift Valley fever in camels. J. Path. Bact., 86:229-231.
Scott GR, Heisch RB, 1959. Rift Valley fever and Rift Valley rodents. East African Medical Journal, 36:665-667.
Shope RE, Peter CJ, Davies FG, 1982. The spread of Rift Valley fever and approaches to its control. Bulletin of the World Health Organization, 60:299-304.
Shope RE, Peters CJ, Walker JS, 1980. Serological relation between Rift Valley fever virus and the viruses of the Phlebotomus fever serogroup. Lancet, 1:886-887.
Smithburn KC, 1949. Rift Valley fever: the neurotropic adaptation of the virus and the experimental use of this modified virus as a vaccine. British Journal of Experimental Pathology, 30:1-16.
Smithburn KC, Haddow AJ, Gillett JD, 1948. Rift Valley fever, isolation of the virus from wild mosquitoes. British Journal of Experimental Pathology, 29:107-121.
Soumaré POL, Freire CCM, Faye O, Diallo M, Oliveira JVCde, Zanotto PMA, Sall AA, 2012. Phylogeography of Rift Valley Fever virus in Africa reveals multiple introductions in Senegal and Mauritania. PLoS ONE, 7(4):e35216. http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0035216
Tesch RB, Peters CJ, Meegan JM, 1982. Studies on the antigenic relationships among Phleboviruses. American Journal of Tropical Medicine and Hygiene, 31:149-155.
Thiongane Y, Thonnon J, Fontenille D, Zeller H, Akapo JA, Gonzalez JP, Digoutte JP, 1997. Epidemiosurveillance of Rift Valley fever in the Rift Valley in Senegal. épidémiologie et Santé Animale, No. 31/32:02.A.08; 3 ref.
Thonnon J, Picquet M, Thiongane Y, Lo M, Sylla R, Vercruysse J, 1999. Rift Valley fever surveillance in the lower Senegal River Basin: update 10 years after the epidemic. Tropical Medicine and International Health, 4(8):580-585; 15 ref.
Turell MJ, Bailey CL, 1987. Transmission studies in mosquitoes (Diptera: Culicidae) with disseminated Rift Valley fever virus infections. Journal of Medical Entomology, 24(1):11-18; 23 ref.
Turell MJ, Presley SM, Gad AM, Cope SE, Dohm DJ, Morrill JC, Arthur RR, 1996. Vector competence of Egyptian mosquitoes for Rift Valley fever virus. American Journal of Tropical Medicine and Hygiene, 54(2):136-139; 20 ref.
Turell MJ, Spielman A, 1992. Nonvascular delivery of Rift Valley fever virus by infected mosquitoes. American Journal of Tropical Medicine and Hygiene, 47(2):190-194; 10 ref.
Weiss KE, 1957. Rift Valley fever: a review. Bull. Epiz. Dis. Afr., 5:431-458.
Woodall JP, 1964. The viruses isolated from arthropods at the East African Virus Research Institute in the 26 years ending December 1963. Proc. E. Afr. Acad., 2:141-146.
Yedloutschnig RJ, Dardiri AH, Walker JS, Peters CJ, Eddy GA, 1979. Immune response of steers, goats and sheep to an inactivated Rift Valley fever vaccine. Proceedings of the United States Animal Health Association 83rd Annual Meeting, 83:253-260.
Yedloutschnig RJ, et al., 1981. Abortion in vaccinated cattle and sheep after challenge with Rift valley fever virus. Veterinary Record, 109:383-384.
Zeller HG, Bessin R, Thiongane Y, Bapetel I, Teou K, Ala MG, Atse AN, Sylla R, Digoutte JP, Akakpo JA, 1995. Rift Valley fever antibody prevalence in domestic ungulates in Cameroon and several West African countries (1989-1992) following the 1987 Mauritanian outbreak. Research in Virology, 146(1):81-85; 17 ref.
Zeller HG, Fontenille D, Traoré-Lamizana M, Thiongane Y, Digoutte JP, 1997. Enzootic activity of Rift Valley fever virus in Senegal. American Journal of Tropical Medicine and Hygiene, 56(3):265-272; 44 ref.
Zeller HG, Rakotoharinadrasana HT, Rakoto-Andrianarivelo M, 1998. Rift Valley fever in Madagascar: infection risks for the abattoir staff in Antananarivo. Revue d'élevage et de Médecine Vétérinaire des Pays Tropicaux, 51(1):17-20; 18 ref.
(http://www.oie.int, accessed 5 June 2013)
Dre Michèle Bouloy
Unité de génétique moléculaire des Bunyavirus
Département de Virologie
25 rue du Dr Roux
75724 Paris cedex 15
Tel: +33-1 126.96.36.199 Fax: +33-1 188.8.131.52
Dr Baratang Alison Lubisi
Onderstepoort Veterinary Institute
Agricultural Research Council
Private Bag X05
Tel: +27-12 529 91 17 Fax: +27-12 529 94 18
Date of report: 03/06/2013
© CAB International 2013. Distributed under license by African Union – Interafrican Bureau for Animal Resources.
This work is licensed under a Creative Commons Attribution-NonCommercial-NoDerivs 3.0 Unported License.