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
International Common Names
cattle plague, rinderpest in pigs- exotic, rinderpest in ruminants- exotic
Rinderpest is an acute to subacute contagious viral disease of large ruminants of the order Artiodactyla that can cause morbidity and mortality rates in excess of ninety per cent, though inapparent infections also occur. The disease is characterized by necrosis and erosions in the gastrointestinal tract that result in severe diarrhoea and dehydration. It is caused by a morbillivirus, a member of a group of enveloped viruses forming a separate genus within the family Paramyxoviridae. Viruses in this genus included rinderpest virus (RPV) causing disease in cattle and other large ruminants, peste des petits ruminants virus (PPRV) causing disease in sheep and goats, canine distemper virus (CDV) which causes disease in many types of carnivore (primarily dogs, but also large cats, seals and mustelids), the human pathogen measles virus (MV), while other members of the genus cause disease in marine mammals. Members of the genus are closely related antigenically and are distinguished from the other paramyxoviruses by their lack of neuraminidase activity and their common use of the cell surface protein CD150 (aka Signalling Lymphocytic Activation Molecule (SLAM)) as the primary receptor.
In terms of economic losses in domestic animals, rinderpest has been the most important member of the group. For centuries it has been the most economically important viral disease of cattle, and the history of the disease, and various efforts to combat it, have filled whole books (see especially Spinage, 2003; Barrett et al., 2006). Most importantly, rinderpest has become the first veterinary disease to be eradicated from the world. The disease, although still notionally on the list of diseases notifiable to the World Organisation for Animal Health (in French, the Office International des Epizooties, and hence abbreviated everywhere as 'OIE'), was declared eradicated by the Food and Agriculture Organisation of the UN (FAO) and the OIE in 2011 (FAO and OIE, 2011). There have been no confirmed cases of rinderpest in livestock or wildlife anywhere in the world since 2001.
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.
Rinderpest has been eradicated from nature, and the only known rinderpest virus that exists is stored in laboratories or vaccine manufacturers.
= 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||Last reported||1972||OIE, 2012|
|Benin||Last reported||1987||OIE, 2012|
|Botswana||Last reported||OIE, 2012|
|Burkina Faso||Last reported||1988||OIE, 2012|
|Burundi||Last reported||1934||OIE, 2012|
|Cameroon||Last reported||1986||OIE, 2012|
|Cape Verde||Disease never reported||OIE, 2012|
|Central African Republic||Last reported||1983||OIE, 2012|
|Chad||Last reported||1983||OIE, 2012|
|Comoros||Disease never reported||OIE, 2012|
|Congo||Disease never reported||OIE, 2009|
|Congo Democratic Republic||Last reported||1961||OIE, 2012|
|Côte d'Ivoire||Last reported||1986||OIE, 2012|
|Djibouti||Last reported||1985||OIE, 2012|
|Egypt||Last reported||1986||OIE, 2012|
|Eritrea||Last reported||1995||OIE, 2012|
|Ethiopia||Last reported||1995||OIE, 2012|
|Gabon||Disease never reported||OIE, 2012|
|Gambia||Last reported||1965||OIE, 2012|
|Ghana||Last reported||1988||OIE, 2012|
|Guinea||Last reported||1967||OIE, 2012|
|Guinea-Bissau||Last reported||1967||OIE, 2012|
|Kenya||Last reported||2001||OIE, 2012; OIE, 2003|
|Lesotho||Disease not reported||OIE, 2009|
|Libya||Last reported||1963||OIE, 2012|
|Madagascar||Disease never reported||OIE, 2009|
|Malawi||Disease never reported||OIE, 2012|
|Mali||Last reported||1986||OIE, 2012|
|Mauritius||Disease never reported||OIE, 2012|
|Morocco||Disease never reported||OIE, 2012|
|Mozambique||Last reported||1896||OIE, 2012|
|Namibia||Last reported||1905||OIE, 2012|
|Niger||Last reported||1986||OIE, 2012|
|Nigeria||Last reported||1987||OIE, 2012|
|Réunion||Last reported||1902||OIE Handistatus, 2005|
|Rwanda||Last reported||1932||OIE, 2012|
|Sao Tome and Principe||Disease not reported||OIE Handistatus, 2005|
|Senegal||Last reported||1978||OIE, 2012|
|Seychelles||Disease not reported||OIE, 2012|
|Sierra Leone||Last reported||1958||OIE, 2012|
|Somalia||Last reported||1983||OIE, 2012|
|South Africa||Last reported||1904||OIE, 2012|
|Sudan||Last reported||1998||OIE, 2012|
|Swaziland||Last reported||1898||OIE, 2012|
|Tanzania||Last reported||1997||OIE, 2012|
|Togo||Last reported||1986||OIE, 2012|
|Tunisia||Disease never reported||OIE, 2012|
|Uganda||Last reported||1994||OIE, 2012|
|Zambia||Last reported||1896||OIE, 2012|
|Zimbabwe||Last reported||1898||OIE, 2012|
Rinderpest virus infects a wide variety of vertebrates. Some of these, including rabbits, hamsters, mice, giant rats (Cricetomys gambianus), ferrets, and susliks (Citellus mongoliscus ramosus) are usually only infected experimentally, and then often only by using strains of virus adapted to them. In the field, only artiodactyla are naturally infected, although dogs fed infected meat may develop antibodies to the virus, suggesting subclinical infection. Amongst domestic stock, cattle and buffaloes (Bubalus bubalus) are especially susceptible and are more frequently infected than other species. Sheep, goats and Asiatic pigs are also susceptible and may develop clinical disease, though this has only been recorded experimentally. European breeds of pig undergo subclinical infection. Although some early reports indicated that camels are susceptible to clinical disease, more recent experimental studies have shown only mild or subclinical disease in this species. Contact transmission from cattle to camels occurs under experimental conditions, but is probably rare in the field.
Infection of wild artiodactyls with strains largely maintained in cattle causes a wide spectrum of clinical disease, ranging from very severe in African buffalo (Syncerus caffer), giraffe (Giraffa camelopardalis), eland (Taurotragus oryx) and kudu (Tragelaphus strepciceros, T. imberbis) through increasingly less severe syndromes in other antelopes to mild or atypical in impala (Aepyceros melampus) and subclinical in hippopotami (Hippopotamus amphibius). There is also variation in susceptibility to clinical disease between breeds or races of a species, especially cattle. Most European cattle breeds (Bos taurus) are more susceptible than Bos indicus breeds. African humpless cattle, such as the Ankole in East Africa, are notoriously susceptible in comparison to East African zebus. Because Japanese black cattle reacted so severely to goat-adapted vaccines that were sufficiently attenuated for other cattle, the virus had to be further attenuated in rabbits and embryonated chickens' eggs.
Vectors and intermediate hosts are not involved in the transmission of rinderpest.
|Bos grunniens (yaks)||Domesticated host, Wild host|
|Bos indicus (zebu)||Domesticated host|
|Bos taurus (cattle)||Domesticated host|
|Bubalus bubalis (buffalo)||Domesticated host|
|Capra hircus (goats)||Domesticated host|
|Ovis aries (sheep)||Domesticated host|
Digestive - Large Ruminants
Digestive - Small Ruminants
Multisystem - Large Ruminants
Multisystem - Small Ruminants
Rinderpest virus (RPV) can infect a wide variety of vertebrates. In the field, only artiodactyla are naturally infected, although dogs fed infected meat may develop antibodies to the virus suggesting subclinical infection, and various laboratory species can be infected experimentally. Amongst domestic stock, cattle and buffalo (Bubalus bubalus) are especially susceptible and are more frequently infected than other species. Sheep, goats and Asiatic pigs are also susceptible and may develop clinical disease. European breeds of pig and camels undergo subclinical or mild infection.
Infection of wild artiodactyls with strains largely maintained in cattle causes a wide spectrum of clinical disease, ranging from very severe in African buffalo (Syncerus caffer), giraffe (Giraffa camelopardalis), eland (Taurotragus oryx) and kudu (Tragelaphus strepciceros, T. imberbis), through increasingly less severe syndromes in other antelopes, to mild or atypical in impala (Aepyceros melampus) and subclinical in hippopotami (Hippopotamus amphibius). There is also variation in susceptibility to clinical disease between breeds or races of a species, especially cattle. Most European cattle breeds (Bos taurus) are more susceptible than Bos indicus breeds. African humpless cattle, such as the Ankole in East Africa, are notoriously susceptible in comparison to East African zebus.
Infected animals excrete infectious virus in their ocular, nasal, oral and vaginal secretions and faeces. Excretion begins 1 or 2 days before the onset of fever, the first clinical sign, and continues for 9 to 10 days after the start of pyrexia. Highest titres of virus are excreted during the early stages of clinical disease when epithelial lesions, especially those in the mouth, are developing to their maximum extent. Subsequently, the titres of excreted virus wane as antibody develops. Recovered cows may abort an infected foetus some weeks after apparent recovery, with virus excretion in their uterine and vaginal discharges.
The fragility of the virus ensures that most infectivity survives for only a few hours outside the host, though some may persist under favourable conditions for up to 2 to 4 days. Carcass decomposition inactivates the virus within 1 to 3 days.
Spread of RPV was affected almost exclusively by contact between infected and susceptible animals. Transmission by infected aerosols probably only occurs under ideal conditions of close proximity and gentle air currents, i.e. amongst housed animals. There is no carrier state in rinderpest and recovered animals do not excrete infectious RPV and are not involved in the maintenance and transmission of the disease. The virus is not transmitted by arthropods and the potential for transmission through abortion is limited. Consequently, RPV has a short direct cycle of infection and is spread by close contact. Under experimental conditions regular contact transmission can be difficult to achieve.
In the field, rinderpest was maintained by large, heterogeneous populations of animals with a sufficient supply of new susceptibles. In Africa in recent times the endemic areas have been those with large cattle populations belonging to nomadic or semi-nomadic people, which ensures good mixing of the population, especially when restricted by the availability of water during dry seasons.
In highly susceptible populations rinderpest behaved in epidemic fashion with the virus infecting virtually all susceptible individuals and causing severe clinical disease in most age groups. Endemic rinderpest, however, was much milder and was maintained by young animals usually less than 2 years old that have lost their maternal immunity. Intermediate patterns also existed.
Wildlife played an important role in rinderpest. In Asia wildlife have been described with clinical disease and such infected animals can transmit infection to other susceptible species, including domestic stock. However, the sizes and densities of wildlife populations are low and they are not considered to be involved in the maintenance of the virus in Asia. In Africa, however, the greater population sizes and densities, the larger number of susceptible species, and the frequency with which the disease used to be reported in wildlife have lead to considerable study of rinderpest in these species. Until the 1960s a widely held view was that wildlife could maintain the virus independently of cattle, though some authorities considered cattle to be the main reservoir of infection. However, when cell-culture-attenuated vaccine led to the eradication of the disease from cattle in Maasailand [East Africa] in the early 1960s, clinical disease also disappeared from wildlife. The absence of antibodies in wildebeest and other species born after 1963 supported this and as a consequence opinion changed to the view that wildlife could not maintain the virus, which is still widely held today. This view was also supported by the total eradication of rinderpest in all countries through sustained vaccination of livestock cattle and buffalo followed by surveillance of the livestock animals.
Rinderpest causes both direct and indirect economic losses on domestic livestock enterprises. Direct losses from high levels of mortality and loss of production, indirect losses from reduced international trade, decreased sales during quarantine restrictions and the continual high costs of control by vaccination in infected areas. The direct losses caused by rinderpest can be enormous. In the eighteenth century alone, some 200 million cattle may have died in north-western Europe, and it has been estimated that as many as half a million cattle died during the last epidemic in Britain in the 1860s. Africa south of the Zambezi lost over 2.5 million cattle during the Great African Pandemic in the 1890s and, despite the availability of an effective vaccine, over a million cattle may have died in Africa during the second continental pandemic in the 1980s. During a recent epidemic of rinderpest in the north of Pakistan which had been free of the disease for 60 years, perhaps 10% of the area's estimated half million cattle died. Individual villages lost nearly all of their stock and one valley lost 7,000 of its population of 12,000 animals in just 4 months.
The losses in endemic areas are far less severe, but nevertheless there is continual mortality in young stock. In southern Sudan, one of the remaining endemic foci in Africa, the virus affects nearly all unprotected young stock, with an estimated case mortality of up to 40%.
For countries in the Horn of Africa the export of livestock, primarily to the Middle East, is a major source of income. In the 1980s, Yemen and Saudi Arabia placed a ban on the importation of livestock from Somalia after it was accused of exporting rinderpest to them. At the time livestock export was the number one source of income for Somalia and the ban had a severe effect on the national economy.
This disease is not a zoonosis.
A proportion of infected cattle show slight lymphocytosis before the onset of pyrexia. This is followed by marked lymphopenia, caused by lymphoid necrosis, which in most cases lasts throughout the acute clinical stage of the disease. During convalescence, lymphocyte levels slowly return to normal over a period of days to weeks. The number of neutrophils remain relatively unaltered, though juvenile forms are not infrequent during the terminal stages of fatal infection. However, a degree of neutropenia that parallels the decline in lymphocyte levels has been reported. Eosinophils may also disappear from the blood during the early stages of clinical disease, returning to normal levels some 2 to 3 weeks later. In severe cases the excessive loss of water causes haemoconcentration.
Serum aspartate transaminase and blood urea nitrogen levels increase during severe cases of disease. Serum chloride levels fall markedly in terminal illness, and other electrolytes may decrease in absolute terms, although this can be masked by haemoconcentration. Blood clotting may be impaired in severely affected animals. Serum protein levels may be lowered, especially in fatally infected animals. In cattle recovering from experimental infections a rise in serum globulins was attributed to the specific humoral response to the virus, but since the challenge material was citrated blood this may need re-interpretation in the light of known responses to heterologous tissue antigens.
The lesions of rinderpest are a direct result of virus-induced cytopathology. Generally, the severity of the lesions is directly related to the virulence of the strain of virus involved. Complications may arise during convalescence through re-activation of latent pathogens, especially protozoa.
The overall appearance at necropsy is similar for most species that die of typical severe rinderpest. The carcass is dehydrated, sometimes emaciated, and usually soiled with fluid faeces. The eyes are sunken and often encrusted with mucopurulent discharge and the cheeks may show signs of epiphora. Erosions with or without necrotic material may be found throughout the mouth but predeliction sites are the gums, lips, buccal papillae, dorsal and ventral aspects of the tongue and the soft palate. The erosions often extend into the pharynx, anterior oesophagus, rumen (especially the pillars), the reticulum and omasum. Necrotic areas, some of which may penetrate the leaves of the omasum, are sometimes present.
The folds of the abomasum are congested and oedematous and often show necrosis, erosions and haemorrhage along the edges. The fundus of the abomasum may have small discrete erosions that increase in size towards the pylorus where whole areas of mucosa may become desquamated. The early necrotic lesions are pale-greyish, whereas the erosions are often red as a result of congestion of the underlying lamina propria. Haemorrhage may occur from the raw surfaces. The abomasum is almost invariably severely affected, whereas the small intestine frequently shows less involvement. Congestion, oedema and erosions may occur on the margins of mucosal folds of the anterior duodenum and terminal ileum. The Peyer's patches, being lymphoid tissue, are severely affected and are swollen, dark red to almost black as a result of haemorrhage and may slough completely leaving deep ulcer-like areas. Large erosions are commonly found on the ileocaecal valve. In the large intestine, marked oedema and congestion accompanied by petechiae or larger haemorrhages occur, particularly along the crests of longitudinal folds of the mucosa. This can be very striking in the colon and rectum, meriting the description 'zebra striping'. In acute cases, the gut has little content other than desquamated necrotic epithelium, blood, and fibrin exuding from exposed lamina propria.
The urinary and gall bladders are frequently congested and haemorrhagic with occasional erosions. The vaginal mucosa may be congested and have small erosions.
The mucosa of the upper respiratory tract, including the larynx, is congested and usually covered with mucopurulent exudate. Petechiae are frequent and necrotic, erosive lesions may extend from the nares to the larynx. The tracheal mucosa is frequently congested. Congestion and emphysema may be seen in the lungs, whereas secondary bronchopneumonia may complicate chronic cases.
Although regularly described in early reports, skin lesions are now rarely seen, although they are reputedly common in domestic buffalo. The exudative dermatitis would seem to develop from macular to pustular lesions, but the role of secondary bacterial infections such as Dermatophilus congolensis needs clarification.
Although RPV has a predilection for lymphoid tissues, there are usually few visible changes to the superficial and visceral lymph nodes. These may show congestion, oedema, and a few petechiae. The nodes of animals that die after a prolonged clinical course may be shrunken and may show greyish radial streaks in the cortex, presumably due to haemorrhage. The spleen and haemolymph nodes appear normal or slightly enlarged.
Histopathological lesions become more easily detectable with increasing severity of clinical disease, implying that the pathology is directly related to the ability of a strain to multiply rapidly in the tissues.
The essential histopathology of rinderpest is widespread necrosis of lymphocytes throughout the lymphoid tissues, together with syncytia and intracytoplasmic and (less frequently) intranuclear inclusion bodies. The histology in cattle is similar with lytic destruction of lymphoid tissues, especially germinal centres, sometimes accompanied by an increase in the numbers of macrophages. In acute cases lymph nodes are virtually devoid of cells, with just a reticular stroma containing eosinophilic material remaining.
The early epithelial lesions in the squamous epithelium of the digestive tract are associated with the formation of syncytia and eosinophilic intracytoplasmic inclusions in the stratum spinosum. Infected epithelial cells become necrotic and slough off, leaving clearly demarcated erosions. The erosions heal rapidly unless complicated by secondary infections, which may rarely cause them to ulcerate.
Typical rinderpest is an acute febrile disease with mortality reaching close to 100% with some of the more virulent strains of the virus, such as the Saudi/81 strain, whereas less virulent strains with typical mortalities of 20% or less were circulating in eastern Africa towards the end of the eradication programme. Classic rinderpest is divided into five stages. After a short incubation period (3-5 days) the prodromal phase is seen in which there is a rapid rise in temperature. This is followed by the mucosal phase in which severe mouth lesions are seen and there is a copious nasal and ocular mucopurulent discharge. The affected animals become depressed and anorexic and on post-mortem examination many necrotic lesions of the epithelium are seen throughout the digestive system. This is followed by the diarrhoeal phase where there is severe bloody diarrhoea, the animal is prostrated and dies from dehydration and weakness. In non-fatal cases there follows the fifth phase in which the animals recover and which may take many weeks. During convalescence pregnant animals may abort. With mild strains the incubation period can extend up to 15 days, and most of the clinical signs are less severe than in disease caused by virulent strains, and some may be absent. Because of the immunosuppressive nature of morbilliviruses, secondary bacterial and concurrent parasitic infections may influence the outcome of the disease and it is sometimes difficult to assess the contribution of virus infection.
The clinical and laboratory diagnosis of rinderpest is described in detail in several handbooks and reports. Now that the virus has been eradicated from nature, it is necessary to be extremely careful about making a presumptive diagnosis of rinderpest on the basis of the clinical signs and gross pathology. Because of the economic and political significance that re-emergence of the disease would have, it is essential to obtain laboratory confirmation of a suspected diagnosis of rinderpest as soon as possible. This will allow immediate steps to be taken to control the disease and restrict losses. In particular, because of the similarity in symptoms and the overlap in their susceptible hosts, distinguishing between rinderpest and peste des petits ruminants (PPR) is very important, the latter still being widespread.
The collection of adequate quantities of appropriate specimens greatly increases the chances of an accurate laboratory diagnosis. A thorough clinical examination should be made of animals in suspected herds and six or seven animals in the early acute stage of the disease with fever, mouth lesions and lachrymation should be selected for sampling. Animals that are dead, moribund or have had diarrhoea and mucopurulent discharges for more than 3 days are less reliable sources of virus or antigen as the levels of these decline with the onset of antibody development.
From each selected animal, whole blood should be collected for serum antibody assay, and in anti-coagulant for virus isolation from leukocytes, a biopsy from a superficial lymph node, debris from oral lesions, and ocular and nasal swabs for virus isolation and antigen or nucleic acid detection. If possible, two or more animals should be killed for necropsy examination and collection of up to three universal bottles of spleen and mesenteric lymph nodes. All specimens should be collected and bottled aseptically, kept cool on ice (but not frozen) and transported as rapidly as possible to a diagnostic laboratory.
Glycerol should not be used as a preservative because it inactivates RPV. The use of protease inhibitors and cold storage increases the survival of RPV antigens in tissue suspensions and reduces the degradation of RNA.
At the laboratory, suspensions of solid tissues are prepared in physiological saline or cell-culture medium, the buffy coat is removed from the whole blood and the serum separated from the clotted blood. Thirty per cent tissue suspensions (w/v) for antigen detection can be prepared by most techniques including grinding with sand in a mortar, but 10 per cent suspensions for attempted virus isolation can best be prepared in Ten Broeck or similar grinders. Many of the laboratory tests for RPV were developed before PPRV became widespread; now that RPV is eradicated and PPRV is common, diagnostic tests have to be able to distinguish between these two viruses.
The first procedure that was usually carried out was to detect viral antigen using specific rabbit hyperimmune serum against RPV. The most commonly used assay was the agar-gel immunodiffusion test (AGID) which was simple and easy to read, but not highly specific. While it can be used in the field with swabs and gum debris and can give a result within 2 hours if the micro-version is used, there will be significant cross-reaction with PPRV with AGID and all tests using polyclonal antisera. Counter-immunoelectrophoresis is quicker and more sensitive than AGID but requires more sophisticated equipment, as does immunofluorescence and immunoperoxidase staining, which are both very sensitive, though time-consuming. Although once widely used, complement fixation and conglutinating complement absorption tests are too complicated in comparison with more recently developed tests. If classically prepared rabbit hyperimmune sera were unavailable, sera could be prepared using other immunizing techniques in rabbits or in goats or cattle. A positive test result in any of the tests used to be taken as confirming rinderpest. None of these tests can discriminate reliably between RPV and PPRV
Where cell cultures are unavailable, specimens could be inoculated into known immune and susceptible cattle, making sure that these are isolated from other susceptible animals. Where facilities are available, attempts should be made to isolate the virus in cell culture. Suspensions prepared from swabs, gum debris, buffy coats or lymphoid tissues are inoculated onto growing monolayers of primary or secondary bovine kidney cells in tubes or flasks. Vero cells are also suitable, especially the Vero cell derivatives expressing SLAM protein (Erlenhofer et al., 2001; Hashimoto et al., 2002). After 12-24 hours adsorption the cells are washed, re-fed with maintenance medium and incubated at 37°C. Typical cytopathic effects develop within 3 to 14 days, occasionally longer, and consist initially of foci of round and refractile cells with cytoplasmic processes and small syncytia, followed by generalization throughout the monolayer with distinct syncytium formation. Negative test cultures should be passaged at least once. The virus can be identified by inoculating sample materials into tubes containing antiserum or specific monclonal antibodies to RPV or by examining fixed monolayers using immunofluorescent or immunoperoxidase techniques. Preferably, specific molecular techniques (see below) can be used.
If antigen detection and virus isolation are negative then convalescent animals should be bled again 2 to 4 weeks later. Assays for serum antibodies should demonstrate a four-fold or greater increase in antibody titre in recovered cases. Virus neutralization in microplates was most commonly used for this, although several other techniques such as measles virus haemagglutination inhibition, indirect immunofluorescence, ELISA and counter-immuno-electrophoresis were alternatives.
A number of ELISA tests have been developed. Original indirect tests based upon whole virus antigens have largely been replaced by a range of competition ELISAs (cELISAs) that use monoclonal antibodies to different viral antigens such as the H or N protein, and may also use purified or recombinant antigens. The cELISA has the advantage that laboratories without cell-culture can test thousands of sera, which was often required in the eradication programme, and the sensitivity and specificity of these tests was confirmed during extensive field use. During the early antibody response, serum contains significant levels of IgM to RPV, the detection of which confirm the diagnosis, though this approach is rarely used.
Histopathology is not sufficiently specific to confirm a diagnosis of rinderpest, but demonstration of syncytia and viral inclusions is supportive.
Nucleic-acid techniques including hybridization with probes and polymerase chain reactions (PCR) are capable of detecting minute quantities of RPV RNA in tissues and secretions, and are now often a routine choice for confirmation in reference laboratories and in many national diagnostic centres as the technology has spread. PCR-based techniques offer the vital advantage of being completely specific for RPV as opposed to PPRV, as well as providing amplified viral RNA for nucleotide sequencing in order to establish the virus sub-type or lineage for epidemiological purposes. A 'penside' test based, in a similar manner to tests used to confirm pregnancy in women, upon a specific monoclonal antibody based latex bead agglutination was developed for use with rinderpest (Bruning et al., 1999).
All conditions that cause stomatitis and/or enteritis in domestic stock could be clinically confused with rinderpest, a matter that is altered now with the eradication of the virus in nature. In cattle, difficulties could arise in distinguishing rinderpest from mucosal disease (MD), malignant catarrhal fever, infectious bovine rhinotracheitis (particularly when caused by strains that induce diarrhoea), papular stomatitis, Jembrana disease and foot-and-mouth disease. In small ruminants, peste des petits ruminants (PPR) and Nairobi sheep disease can resemble rinderpest. Infection with Campylobacter spp., Treponema hyodysenteriae and Salmonella spp. needs to be considered when investigating possible rinderpest in pigs.
In practice, only MD in cattle and PPR in small ruminants regularly presented a problem. The clinical signs and gross pathology in cattle with MD can be indistinguishable from rinderpest and diagnosis requires laboratory confirmation. However, mucosal disease usually affects very few animals in a herd, whereas morbidity rates in rinderpest are much higher. Agar-gel immunodiffusion applied to tissue suspensions could rapidly differentiate the two diseases. Immunohistochemical techniques could be used on frozen sections of mesenteric lymph node or on formalin-fixed tissues to distinguish between rinderpest and MD. Failing this, virus isolation with subsequent virus identification had to be attempted, with follow-up studies to detect rising antibody titres.
The differentiation of PPR from rinderpest is more difficult. Useful epidemiological evidence is provided by the absence of disease in cattle. The virus cross-reacts serologically with RPV and is difficult to differentiate with hyperimmune polyclonal sera. Fortunately, contemporary studies have produced monoclonal antibodies and nucleic-acid techniques that clearly distinguish between PPRV and RPV. Even in countries that have previously been free of PPR it is unwise to assume that a rinderpest-like syndrome in small ruminants is not PPR.
Infected animals mount a vigorous response against the virus. Interferon is produced within 2 days of infection, enabling attenuated vaccines to protect cattle very rapidly against challenge by virulent virus. Viral antigens are produced in large amounts throughout the lymphoid tissues and affected epithelia and stimulate an effective antibody response which begins 2 to 5 days after the onset of clinical disease in virulent infections, and some 6 to 10 days after infection with mild or avirulent strains. The early response consists predominantly of IgM antibodies which can be detected by virus neutralization (VN), ELISA, and also, for a period of a few months, by immunoprecipitation, complement fixation and measles virus haemagglutination inhibition. At the same time IgG antibodies are produced but these persist for much longer, usually for life and are usually measured by VN or ELISA tests. High titres (102-103 log10 VN50) of neutralizing antibodies are produced within 2 to 3 weeks of infection and remain high for several months, after which they may decline slowly, but usually remain at easily detectable levels (in excess of 1 log10 VN50) for the rest of the animal's life. Even when neutralizing antibodies decline to very low or undetectable levels the animals are clinically immune, although limited replication of the virus may occur in tissues such as the tonsils before the stimulation of an anamnestic response. The antibody responses of naturally infected cattle and those vaccinated with live tissue culture virus vaccine are indistinguishable.
Secretory antibody is found in nasal secretions of convalescent cattle but its persistence is presumably limited to only a few months after recovery, and the role it plays in preventing re-infection is unknown.
Very few sequence changes are needed to alter the virulence of rinderpest virus; the genome of the cell-culture-adapted vaccine strain differs by less than 0.55% from the virulent virus from which it was derived. Nothing is known concerning the molecular factors that determine the virulence/attenuation of different rinderpest virus strains and it is possible that changes in pathogenic phenotype can occur on passage through different animal species. Selection of a mild form could well be a means whereby the virus evades detection for many years.
The ability to manipulate the rinderpest genome through rescue of live virus from DNA copies of the virion RNA meant that it became possible to address questions concerning virus attenuation and pathogenicity by directly altering virus genes. It was hoped that site-specific mutagenesis, swapping of genes between mild and virulent isolates and the insertion/deletion of genes would lead to an understanding of factors which determine the host range, tissue tropism and pathology of the virus. In practice, although such studies provided information on the natural receptor for the virus (Baron, 2005) and the underlying causes of the stability of the vaccine strain (Baron et al., 2005; Banyard et al., 2005), continued study of the virus will be necessarily restricted in the future, and further animal studies are unlikely.
Experimental infections can be established by all routes of parenteral inoculation and, more variably, by intranasal or conjunctival installation. Natural infection usually occurs via the upper respiratory tract following inhalation of virus-containing aerosols or the oropharynx after ingestion of infected material. Primary multiplication has not been demonstrated in the invaded epithelium but, following intranasal and contact challenge, virus can be recovered within 24 hours from the pharyngeal lymph nodes and tonsils and, to a lesser degree, from other lymph nodes draining the head and upper respiratory tract. In vivo infectivity is closely associated with mononuclear leukocytes and is not readily detected in plasma and other body fluids.
Following primary multiplication in draining lymph nodes, viraemia enables the virus to infect and replicate in lymphoid tissues throughout the body. This increases the viraemia, which then transports the virus to epithelial tissues, especially to those of the alimentary tract where virus-induced cytopathic effects produce the typical lesions of the disease.
There is an inverse relationship between increasing attenuation and the degree of viral multiplication in cattle lymph nodes. Virulent strains of RPV have a greater ability to infect lymphoid cells and mononuclear phagocytes and may grow to higher titres in these cells than do strains which induce mild disease. The cell-culture-attenuated variant of the Kabete 'O' strain of RPV, which is the most commonly used vaccine, only produces low levels of infectivity in lymphoid tissues and is barely detectable in the blood. These low levels of viraemia are probably one reason why attenuated and very mild strains cause so little epithelial damage. The virus has a predilection for T rather than B or null lymphocytes, which corresponds with the distribution of the primary receptor (SLAM).
During disease the virus is also found in non-lymphoid organs, such as the lungs, liver and kidneys where the antigen-bearing cells are usually associated with reticuloendothelial and perivascular connective tissue.
Virulent strains of RPV are excreted from epithelial tissues 1 or 2 days before the appearance of fever or lesions, but the amount of excreted virus increases dramatically as the lesions develop and only starts to decline when the immune response becomes detectable some 4 to 6 days after the start of fever. The virus is usually undetectable by 12-14 days after the start of fever. At the height of virus excretion, 3 to 6 days after the start of pyrexia, virus titres of up to 105 tissue culture infectious doses (TCID50)/swab and up to 106 TCID50/g, respectively can be found in nasal secretions and faeces from cattle infected with virulent strains. This copious output of virus explains why the disease can be so contagious despite the fragility of RV. The diarrhoea and oculo-nasal discharge probably help to increase the transmissibility of the virus by forming infectious aerosols, and by causing greater contamination of the environment.
The severity of the cytopathology caused by the virus before the onset of antibody development influences the course of the disease. Virulent strains cause severe lesions before being restrained by the immune response and such animals, if sufficiently damaged, still die despite high titres of antibody and low or undetectable amounts of virus. The persistence of immunity in recovered animals and those given live-virus vaccines contrasts with the short-lived immunity induced by inactivated vaccines, possibly indicating that T cell-mediated immunity is more important for protection than antibody-mediated.
The massive destruction of lymphocytes causes immunosuppression, probably involving both cell-mediated and humoral immune responses.
Rinderpest is a virus disease and there is no specific therapeutic treatment. Symptomatic treatment for diarrhoea and supportive antibiotic and fluid replacement therapy might conceivably be useful in preventing the death or aiding recovery of important individual animals. However, in practice few animals are treated.
Morbilliviruses are extremely fragile; they are sensitive to sunlight, high temperature, low and high pH and chemicals which can destroy their outer lipid-containing envelope. Outbreaks of these viruses are therefore easily controlled by proper quarantine and hygienic measures. Rinderpest was successfully controlled and eliminated from Europe by these means without vaccines. There is only one serotype of each virus, and there is no evidence for a persistent or carrier state in recovered animals. After recovery from infection, an animal is immune for life and, consequently, vaccination was a very effective means of controlling this disease. Vaccination has been used extensively to control measles virus (MV) in the developed world, but many logistical and financial problems are associated with delivering a heat-labile vaccine in developing countries. These hinder the success of vaccination campaigns in developing countries where approximately 1-2 million children die each year as a result of MV infection. Similar problems were associated with delivering the RPV vaccine, though it was nevertheless used with success to control RPV in many parts of Asia and Africa.
The development of live attenuated vaccines against morbillivirus diseases was the key to achieving effective vaccination, because the immunity they generate is long lived and involves a cell-mediated immune response. Studies using immune-stimulating complexes (ISCOM) vaccines containing purified H or F proteins of canine distemper virus, and with poxvirus recombinants expressing either the H or F protein of rinderpest virus have shown that either antigen can confer immunity to clinical disease in the short term. In contrast, purified H or F antigens alone are not protective even though they generate a strong humoral immune response.
During the 1930s, attenuated rinderpest vaccines were developed by passage of the virus in non-natural hosts: for example, rabbit and embryonated eggs (lapinised/avianised) or goats (caprinised). A lapinised/avianised vaccine was developed in Japan that was used extensively to control the disease in Asia. In India and Africa the caprinised virus was used. However, the latter virus was not completely attenuated and caused some clinical reactions. In the early 1960s a cell-culture-attenuated vaccine was introduced which was completely safe and relatively easy to produce and induced no clinical signs following inoculation into domestic animals. In addition, the virus does not replicate at epithelial surfaces and cannot be transmitted by contact. Immunity following vaccination is complete and lifelong. The vaccine is, however, heat labile and establishment of an effective cold-chain and subsequent seromonitoring to determine the level of herd immunity are essential prerequisites for a successful vaccination campaign. Improvements in freeze-drying techniques have greatly increased the stability of the vaccine in the dry form but it is still very labile when reconstituted and, like MV vaccine, must be used within a very short period.
In the 1960s an internationally funded rinderpest eradication campaign (Joint Programme 15 or JP 15) was carried out in Africa using the cell-culture-attenuated vaccine and almost succeeded in clearing the disease from Africa. However, political instability, lack of funds to continue vaccination and disease surveillance, and the existence of persisting foci of mild infection resulted in devastating outbreaks of rinderpest throughout Africa in the early 1980s. Since 1985, internationally funded control campaigns have succeeded in reducing the prevalence and distribution of the disease on the continent. The Global Rinderpest Eradication Programme (GREP), a combination of vaccination campaigns with targeted surveillance, followed by continued seromonitoring after cessation of vaccination, took place under several different headings in Africa (Pan African Rinderpest Campaign or PARC), West Asia (WAREC) and South Asia (SAREC) in an attempt to eradicate the disease globally by the year 2010. This programme was successful as stated above (see Overview, Distribution), and the disease is now eradicated.
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
Anderson J, Barrett T, Scott GR, 1996. Manual on the diagnosis of rinderpest. Second edition. FAO Animal Health Manual, No. 1:143 pp.; 60 ref.
Banyard AC, Baron MD, Barrett T, 2005. A role for virus promoters in determining the pathogenesis of Rinderpest virus in cattle. Journal of General Virology, 86(4):1083-1092.
Baron MD, 2005. Wild-type Rinderpest virus uses SLAM (CD150) as its receptor. Journal of General Virology, 86(6):1753-1757.
Baron MD, Banyard AC, Parida S, Barrett T, 2005. The plowright vaccine strain of Rinderpest virus has attenuating mutations in most genes. Journal of General Virology, 86(4):1093-1101.
Barrett T, Pastoret PP, Taylor WP, 2006. Rinderpest and Peste des Petits Ruminants: Virus Plagues of Large and Small Ruminants (Biology of Animal Infections). London, UK: Academic Press, 288 pp.
Barrett T, Rossiter PB, 1999. Rinderpest: the disease and its impact in man and humans. Advances in Virus Research, 53:89-110.
Brüning A, Bellamy K, Talbot D, Anderson J, 1999. A rapid chromatographic strip test for the pen-side diagnosis of rinderpest virus. Journal of Virological Methods, 81(1-2):143-154.
Diop BA, Bastiaensen P, 2005. Achieving full eradication of rinderpest in Africa. Veterinary Record, 157(8):239-240.
Erlenhoefer C, Wurzer WJ, Loffler S, Schneider-Schaulies S, Meulen Vter , Schneider-Schaulies J, 2001. CD150 (SLAM) is a receptor for measles virus but is not involved in viral contact-mediated proliferation inhibition. Journal of Virology, 75:4499-4505.
FAO, OIE, 2011. Joint FAO/OIE Committee on Global Rinderpest Eradication, Final Report, May 2011. Rome, Italy: Food and Agriculture Organisation of the United Nations (FAO), 22 pp.
Hashimoto K, Ono N, Tatsuo H, Minagawa H, Takeda M, Takeuchi K, Yanagi Y, 2002. SLAM (CD150)-independent measles virus entry as revealed by recombinant virus expressing green fluorescent protein. Journal of Virology, 76:6743-6749.
KOCKRA, 2006. Rinderpest and wildlife. In: Rinderpest and Peste des Petits Ruminants, Virus Plagues of Large and Small Ruminants [ed. by Barrett, T. \Pastoret, P. P. \Taylor, W. P.]. Oxford, UK: Academic Press, 143-162 pp.
Mack R, 1970. The great African cattle plague epidemic of the 1890s. Tropical Animal Health and Production 2:210-219.
Normile D, 2008. Rinderpest. Driven to extinction. Science, 319:1606-9.
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, 2003. Rinderpest in Bangladesh. Disease Information, 16, No. 29.
OIE, 2003. Rinderpest in Kenya. Disease Information, 16, No. 48.
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
Plowright W, 1968. Rinderpest virus. Monographs in Virology, 3:25-110.
Rossiter P, Wamwayi H, Ndungu E, 2006. Rinderpest seroprevalence in wildlife in Kenya and Tanzania, 1982-1993. Preventive Veterinary Medicine, 75(1/2):1-7.
Rossiter PB, 1994. Rinderpest. In: Coetzer JAW, Thomson GR, Tustin RC, eds. Infectious Diseases of Livestock in Southern Africa. Cape Town, RSA: Oxford University Press.
Scott GR, 1964. Rinderpest. Advances in Veterinary Science, 9:113-224.
Spinage CA, 2003. Cattle plague: a history [ed. by Spinage, C. A.]. New York, USA: Kluwer Academic/Plenum Publishers, xx + 765 pp.
Taylor WP, Roeder PL, Rweyemamu MM, Melewas JN, Majuva P, Kimaro RT, Mollel JN, Mtei BJ, Wambura P, Anderson J, Rossiter PB, Kock R, Melengeya T, Ende Rvan den, 2002. The control of rinderpest in Tanzania between 1997 and 1998. Tropical Animal Health and Production, 34(6):471-487.
Wamwayi HM, Fleming M, Barrett T, 1995. Characterisation of African isolates of rinderpest virus. Veterinary Microbiology, 44(2/4):151-163.
(http://www.oie.int, accessed 5 June 2013)
Dr Geneviève Libeau
Control of Exotic and Emerging Animal Diseases
Programme Santé Animale
Campus International de Baillarguet TA A-15/G
34398 Montpellier Cedex 5
Tel: +33 (0)4 67 59 37 98 Fax: +33 (0)4 67 59 38 50
Dr Michael Baron
Institute for Animal Health
Ash Road, Pirbright
Woking, Surrey, GU24 0NF
Tel: +44-1483 23.24.41 Fax: +44-1483 23.24.48
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.