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Animal Health and Production Compendium
Selected sections for: rinderpest
Identity Pathogen/s Overview Distribution Distribution Table Hosts/Species Affected Host Animals Systems Affected List of Symptoms/Signs Epidemiology Zoonoses and Food Safety Pathology Diagnosis Disease Course Disease Treatment Prevention and Control References Links to Websites Images
Datasheet Type(s): Animal Disease
Identity
Preferred Scientific Name
rinderpest
International Common Names
English acronym
RP
English
cattle plague, rinderpest in pigs- exotic, rinderpest in ruminants- exotic
French
peste bovine
Arabic
taoun
Local Common Names
East Africa
olodua
Pathogen/s
rinderpest virus
Overview
Rinderpest is an acute to subacute contagious viral disease of ruminants and pigs 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) infecting cattle and other large ruminants, peste des petits ruminants virus (PPRV) infecting sheep and goats, canine distemper virus (CDV) which infects carnivores, human measles virus (MV), and other members in marine mammals. Members of the genus are closely related antigenically and are distinguished from the other paramyxoviruses by their lack of neuraminidase activity.
In terms of economic losses in domestic animals, rinderpest is the most important member of the group. In the world today it is an infection of extensive pastoral cattle herds in two areas of Africa and in some concentrations of inadequately vaccinated cattle and buffalo herds in Pakistan.
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 Handistatus 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.
Distribution
Rinderpest is found in localised regions of Pakistan, Somalia and Sudan. Its presence is uncertain in Afghanistan, Yemen and Kenya.
Distribution Table
Country Distribution Last Reported Origin First Reported Invasive References Notes
ASIA
Afghanistan Disease not reported OIE, 2009
Armenia Disease not reported OIE, 2009
Azerbaijan Disease not reported OIE, 2009
Bahrain Disease not reported OIE, 2009
Bangladesh Disease not reported NULL OIE, 2009; OIE, 2003
Bhutan Disease not reported OIE, 2009
Brunei Darussalam Disease not reported OIE Handistatus, 2005
Cambodia Disease not reported OIE, 2009
China No information available OIE, 2009
-Hong Kong Disease not reported OIE, 2009
Georgia (Republic of) Last reported 1989 OIE Handistatus, 2005
India Disease not reported OIE, 2009
Indonesia Disease not reported OIE, 2009
Iran Disease not reported OIE, 2009
Iraq Disease not reported OIE, 2009
Israel Disease not reported OIE, 2009
Japan Disease not reported OIE, 2009
Jordan Disease not reported OIE, 2009
Kazakhstan Disease not reported OIE, 2009
Korea, DPR Last reported 1948 OIE Handistatus, 2005
Korea, Republic of Disease not reported OIE, 2009
Kuwait Disease not reported OIE, 2009
Kyrgyzstan Disease never reported OIE, 2009
Laos Disease not reported OIE, 2009
Lebanon Disease not reported OIE, 2009
Malaysia Disease not reported OIE, 2009
-Peninsular Malaysia Disease never reported OIE Handistatus, 2005
-Sabah Disease never reported OIE Handistatus, 2005
-Sarawak Disease never reported OIE Handistatus, 2005
Mongolia Disease not reported OIE, 2009
Myanmar Disease not reported OIE, 2009
Nepal Disease not reported OIE, 2009
Oman Disease not reported OIE, 2009
Pakistan Disease not reported OIE, 2009
Philippines Disease not reported OIE, 2009
Qatar Disease not reported OIE, 2009
Saudi Arabia Disease not reported OIE, 2009
Singapore Disease not reported OIE, 2009
Sri Lanka Disease not reported OIE, 2009
Syria Disease not reported OIE, 2009
Taiwan Last reported 1950 OIE Handistatus, 2005
Tajikistan Disease never reported OIE, 2009
Thailand Disease not reported OIE, 2009
Turkey Disease not reported OIE, 2009
Turkmenistan Disease never reported OIE Handistatus, 2005
United Arab Emirates Disease not reported OIE, 2009
Uzbekistan Disease never reported OIE Handistatus, 2005
Vietnam Disease not reported OIE, 2009
Yemen No information available OIE, 2009
AFRICA
Algeria Disease never reported OIE, 2009
Angola Disease not reported OIE, 2009
Benin Disease not reported OIE, 2009
Botswana Disease not reported OIE, 2009
Burkina Faso Disease not reported OIE, 2009
Burundi Disease not reported OIE Handistatus, 2005
Cameroon Last reported 1986 OIE Handistatus, 2005
Cape Verde No information available OIE Handistatus, 2005
Central African Republic Last reported 1983 OIE Handistatus, 2005
Chad Disease not reported OIE, 2009
Congo Disease never reported OIE, 2009
Congo Democratic Republic Disease not reported OIE Handistatus, 2005
Côte d'Ivoire Last reported 1986 OIE Handistatus, 2005
Djibouti No information available OIE, 2009
Egypt Disease not reported OIE, 2009
Eritrea No information available OIE, 2009
Ethiopia Disease not reported OIE, 2009
Gabon Disease never reported OIE, 2009
Gambia Disease not reported OIE, 2009
Ghana Disease not reported OIE, 2009
Guinea Disease not reported OIE, 2009
Guinea-Bissau Disease not reported OIE, 2009
Kenya Disease not reported 2003 OIE, 2009; OIE, 2003
Lesotho Disease not reported OIE, 2009
Libya Last reported 1966 OIE Handistatus, 2005
Madagascar Disease never reported OIE, 2009
Malawi Disease never reported OIE, 2009
Mali No information available OIE, 2009
Mauritius Disease never reported OIE, 2009
Morocco Disease never reported OIE, 2009
Mozambique Disease not reported OIE, 2009
Namibia Disease not reported OIE, 2009
Niger Last reported 1985 OIE Handistatus, 2005
Nigeria Disease not reported OIE, 2009
Réunion Last reported 1902 OIE Handistatus, 2005
Rwanda Disease not reported OIE, 2009
Sao Tome and Principe Disease not reported OIE Handistatus, 2005
Senegal Disease not reported OIE, 2009
Seychelles Disease not reported OIE Handistatus, 2005
Somalia No information available OIE Handistatus, 2005
South Africa Disease not reported OIE, 2009
Sudan Disease not reported OIE, 2009
Swaziland Disease not reported OIE, 2009
Tanzania Disease not reported OIE, 2009
Togo Disease not reported OIE, 2009
Tunisia Disease never reported OIE, 2009
Uganda Disease not reported OIE, 2009
Zambia Disease not reported OIE, 2009
Zimbabwe Disease not reported OIE, 2009
NORTH AMERICA
Bermuda Disease not reported OIE Handistatus, 2005
Canada Disease never reported OIE, 2009
Greenland Disease never reported OIE, 2009
Mexico Disease not reported OIE, 2009
USA Disease never reported OIE, 2009
-Georgia Disease not reported OIE, 2009
CENTRAL AMERICA
Barbados Disease never reported OIE Handistatus, 2005
Belize Disease never reported OIE, 2009
British Virgin Islands Disease never reported OIE Handistatus, 2005
Cayman Islands Disease never reported OIE Handistatus, 2005
Costa Rica Disease never reported OIE, 2009
Cuba Disease never reported OIE, 2009
Curaçao Disease not reported OIE Handistatus, 2005
Dominica Disease not reported OIE Handistatus, 2005
Dominican Republic Disease never reported OIE, 2009
El Salvador Disease never reported OIE, 2009
Guadeloupe Disease never reported OIE, 2009
Guatemala Disease never reported OIE, 2009
Haiti Disease never reported OIE, 2009
Honduras Disease never reported OIE, 2009
Jamaica No information available OIE, 2009
Martinique Disease never reported OIE, 2009
Nicaragua Disease never reported OIE, 2009
Panama Disease never reported OIE, 2009
Saint Kitts and Nevis Disease never reported OIE Handistatus, 2005
Saint Vincent and the Grenadines Disease never reported OIE Handistatus, 2005
Trinidad and Tobago Disease never reported OIE Handistatus, 2005
SOUTH AMERICA
Argentina Disease never reported OIE, 2009
Bolivia Disease never reported OIE, 2009
Brazil Disease not reported OIE, 2009
Chile Disease never reported OIE, 2009
Colombia Disease never reported OIE, 2009
Ecuador Disease never reported OIE, 2009
Falkland Islands Disease never reported OIE Handistatus, 2005
French Guiana Disease never reported OIE, 2009
Guyana Disease never reported OIE Handistatus, 2005
Paraguay Disease never reported OIE Handistatus, 2005
Peru Disease never reported OIE, 2009
Uruguay Disease never reported OIE, 2009
Venezuela Disease never reported OIE, 2009
EUROPE
Albania Disease not reported OIE, 2009
Andorra Disease never reported OIE Handistatus, 2005
Austria Disease not reported OIE, 2009
Belarus Disease never reported OIE, 2009
Belgium Disease not reported OIE, 2009
Bosnia-Hercegovina Last reported 1883 OIE Handistatus, 2005
Bulgaria Disease not reported OIE, 2009
Croatia Disease not reported OIE, 2009
Cyprus Disease never reported OIE, 2009
Czech Republic Disease not reported OIE, 2009
Denmark Disease not reported OIE, 2009
Estonia Disease never reported OIE, 2009
Finland Disease not reported OIE, 2009
France Disease not reported OIE, 2009
Germany Disease not reported OIE, 2009
Greece Disease not reported OIE, 2009
Hungary Disease not reported OIE, 2009
Iceland Disease never reported OIE, 2009
Ireland Disease not reported OIE, 2009
Isle of Man (UK) Disease never reported OIE Handistatus, 2005
Italy Disease not reported OIE, 2009
Jersey Disease never reported OIE Handistatus, 2005
Latvia Disease not reported OIE, 2009
Liechtenstein Disease not reported OIE, 2009
Lithuania Disease never reported OIE, 2009
Luxembourg Disease never reported OIE, 2009
Macedonia Disease never reported OIE, 2009
Malta Disease never reported OIE, 2009
Moldova Disease never reported OIE Handistatus, 2005
Montenegro Disease not reported OIE, 2009
Netherlands Disease not reported OIE, 2009
Norway Disease never reported OIE, 2009
Poland Disease not reported OIE, 2009
Portugal Disease never reported OIE, 2009
Romania Disease not reported OIE, 2009
Russian Federation Disease not reported OIE, 2009
Serbia Disease not reported OIE, 2009
Slovakia Disease not reported OIE, 2009
Slovenia Disease not reported OIE, 2009
Spain Disease never reported OIE, 2009
Sweden Disease not reported OIE, 2009
Switzerland Disease not reported OIE, 2009
Ukraine Disease never reported OIE, 2009
United Kingdom
-Northern Ireland Last reported 1900 OIE Handistatus, 2005
United Kingdom Disease not reported OIE, 2009
Yugoslavia (former) Disease never reported OIE Handistatus, 2005
Yugoslavia (Serbia and Montenegro) Last reported 1883 OIE Handistatus, 2005
OCEANIA
Australia Disease not reported OIE, 2009
French Polynesia Disease never reported OIE, 2009
New Caledonia Disease never reported OIE, 2009
New Zealand Disease never reported OIE, 2009
Samoa Disease never reported OIE Handistatus, 2005
Vanuatu Disease never reported OIE Handistatus, 2005
Wallis and Futuna Islands No information available OIE Handistatus, 2005
Hosts/Species Affected
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. 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.
Host Animals
Animal name Context
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
Sus scrofa (pigs) Domesticated host
Systems Affected
Digestive - Large Ruminants
Digestive - Pigs
Digestive - Small Ruminants
Multisystem - Large Ruminants
Multisystem - Pigs
Multisystem - Small Ruminants
List of Symptoms/Signs
Sign Life Stages Type
Digestive Signs
Mucous, mucoid stools, faeces Sign [C]
Bloody stools, faeces, haematochezia Sign [C]
Grinding teeth, bruxism, odontoprisis Sign [C]
Rumen hypomotility or atony, decreased rate, motility, strength Sign [C]
Decreased amount of stools, absent faeces, constipation Sign [C]
Pharyngeal ulcers, vesicles, erosion, papules, sores pharynx C1 ( All Stages ) Diagnosis
Diarrhoea C1 ( All Stages ) Diagnosis [C]
Unusual or foul odor, stools, faeces C1 ( All Stages ) Diagnosis [C]
Congestion oral mucous membranes, erythema, redness oral mucosa C1 ( All Stages ) Diagnosis [C]
Oral mucosal ulcers, vesicles, plaques, pustules, erosions, tears C1 ( All Stages ) Diagnosis [C]
Tongue ulcers, vesicles, erosions, sores, blisters, cuts, tears C1 ( All Stages ) Diagnosis [C]
Excessive salivation, frothing at the mouth, ptyalism Sign [C]
Anorexia, loss or decreased appetite, not nursing, off feed C1 ( All Stages ) Sign [C]
General Signs
Fever, pyrexia, hyperthermia C1 ( All Stages ) Sign [C]
Trembling, shivering, fasciculations, chilling Sign [C]
Trembling, shivering, fasciculations, chilling Sign [C]
Petechiae or ecchymoses, bruises, ecchymosis Sign [C]
Generalized weakness, paresis, paralysis Sign [C]
Inability to stand, downer, prostration Sign [C]
Tenesmus, straining, dyschezia Sign [C]
Hypothermia, low temperature Sign [C]
Dehydration Sign [C]
Polydipsia, excessive fluid consumption, excessive thirst Sign [C]
Nervous Signs
Tremor Sign [C]
Dullness, depression, lethargy, depressed, lethargic, listless Sign [C]
Ophthalmology Signs
Conjunctival, scleral, injection, abnormal vasculature Sign [C]
Conjunctival, scleral, redness C1 ( All Stages ) Diagnosis [C]
Lacrimation, tearing, serous ocular discharge, watery eyes C1 ( All Stages ) Diagnosis [C]
Blindness L1 ( All Stages ) Diagnosis
Purulent discharge from eye C1 ( All Stages ) Diagnosis [C]
Photophobia Sign [C]
Blepharospasm Sign [C]
Pain/Discomfort Signs
Colic, abdominal pain Sign [C]
Skin pain Sign [C]
Reproductive Signs
Agalactia, decreased, absent milk production Sign [C]
Vulval ulcers, vesicles, erosions, tears, cuts, pustules, papules Sign [C]
Vaginal or cervical ulcers, vesicles, erosions, tears, papules, pustules C3 ( Heifer ), C4 ( Cow ) Diagnosis
Abortion or weak newborns, stillbirth C3 ( Heifer ), C4 ( Cow ) Diagnosis [C]
Respiratory Signs
Nasal mucosal ulcers, vesicles, erosions, cuts, tears, papules, pustules C1 ( All Stages ) Diagnosis [C]
Purulent nasal discharge C1 ( All Stages ) Diagnosis [C]
Mucoid nasal discharge, serous, watery C1 ( All Stages ) Diagnosis [C]
Abnormal breath odor, foul odor mouth C1 ( All Stages ) Diagnosis [C]
Dyspnea, difficult, open mouth breathing, grunt, gasping Sign [C]
Increased respiratory rate, polypnea, tachypnea, hyperpnea Sign [C]
Coughing, coughs Sign [C]
Skin/Integumentary Signs
Alopecia, thinning, shedding, easily epilated, loss of, hair Sign [C]
Rough hair coat, dull, standing on end Sign [C]
Skin erythema, inflammation, redness L1 ( All Stages ) Sign [C]
Skin papules L1 ( All Stages ) Sign [C]
Skin pustules Sign [C]
Skin crusts, scabs Sign [C]
Moist skin, hair or feathers Sign [C]
Urinary Signs
Polyuria, increased urine output Sign [C]
Epidemiology
Rinderpest virus (RV) infects 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 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. 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 RV is effected 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 RV 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, RV 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 is 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 behaves in epidemic fashion with the virus infecting virtually all susceptible individuals and causing severe clinical disease in most age groups. Endemic rinderpest, however, is much milder and is maintained by young animals usually less than 2 years old that have lost their maternal immunity. Intermediate patterns also exist.
Wildlife play 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.
Zoonoses and Food Safety
This disease is not a zoonosis.
Pathology
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 RV 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.
Diagnosis
Clinical Signs
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 are currently circulating in eastern Africa. 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. A presumptive diagnosis of rinderpest can be made on the basis of the clinical signs and gross pathology. However, in countries where the disease is not prevalent, and especially in regions dependent on livestock exports, it is essential to obtain laboratory confirmation of the diagnosis as soon as possible. Countries where rinderpest is either endemic or a high risk should treat any syndrome resembling rinderpest as such until proven otherwise. This will allow immediate steps to be taken to control the disease and restrict losses.
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 RV. The use of anti-proteases increases the survival of RV antigens in tissue suspensions and reduces the degradation of RNA.
Laboratory Diagnosis
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.
The first procedure usually carried out is to detect viral antigen using specific rabbit hyperimmune serum against RV. The most commonly used assay is the agar-gel immunodiffusion test (AGID) which is simple, easy to read, and highly specific. Moreover, 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. Counter-immunoelectrophoresis is quicker and more sensitive than AGID but requires more sophisticated equipment. Immunofluorescence and immunoperoxidase staining are very sensitive but also need more equipment than AGID. Although once widely used, complement fixation and conglutinating complement absorption tests are too complicated in comparison with more recently developed tests. Various haemagglutination assays are sensitive but not yet widely applied though latex bead agglutination tests have given encouraging preliminary results and, if combined with monoclonal antibodies, might prove very sensitive. If classically prepared rabbit hyperimmune sera are unavailable, sera can be prepared using other immunizing techniques in rabbits or in goats or cattle. A positive test result in any of the tests confirms rinderpest.
Where cell cultures are unavailable, specimens can 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. Vero cells are also suitable, whereas culture systems such as microplates can also be used but may be less sensitive. After 12-24 hours adsorption the tubes are washed, re-fed with maintenance medium and rolled 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 to RV or by examining fixed monolayers using immunofluorescent or immunoperoxidase techniques.
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 are 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 that use monoclonal antibodies to different viral antigens such as the H or N protein, and may also use purified or recombinant antigens. The ELISA has the advantage that laboratories without cell-culture can test thousands of sera, which is often required in current eradication programmes, and the sensitivity and specificity of these new tests is under validation at present. During the early antibody response, serum contains significant levels of IgM to RV, 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 are capable of detecting minute quantities of RV RNA in tissues and secretions, and are now often a routine choice for confirmation in reference laboratories. The PCR offers the advantage of 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 specific monoclonal antibody based latex bead agglutination is being developed for use with rinderpest.
Differential Diagnosis
All conditions that cause stomatitis and/or enteritis in domestic stock can be clinically confused with rinderpest. In cattle, difficulties may 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 present 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 can rapidly differentiate the two diseases. Immunohistochemical techniques can 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 must 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 RV and is difficult to differentiate with hyperimmune polyclonal sera. Fortunately, contemporary studies have produced monoclonal antibodies and nucleic-acid techniques that clearly distinguish between PPR virus and RV, at least for the limited number of strains tested to date. In African countries that have previously been free of PPR it is unwise to assume that a rinderpest-like syndrome in small ruminants is not PPR.
Immune Response
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.
Disease Course
Very few sequence changes are needed to alter the virulence of rinderpest virus; the genome of the cell-culture 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 means that is now possible to address questions concerning virus attenuation and pathogenicity by directly altering virus genes. Site-specific mutagenesis, swapping of genes between mild and virulent isolates and the insertion/deletion of genes will lead to an understanding of factors which determine the host range, tissue tropism and pathology of the virus. This technology will also help in the control of rinderpest by providing genetically defined vaccines, which will enable vaccinated animals to be distinguished from those that have been naturally infected.
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 RV 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 RV, 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.
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 RV 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, implying that recovered animals perhaps may be continually re-stimulated immunologically by RV antigen throughout their lives.
The massive destruction of lymphocytes causes immunosuppression, probably involving both cell-mediated and humoral immune responses.
Disease Treatment
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.
Prevention and Control
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 is 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 are associated with delivering RPV vaccine, though it has been used with success to control RPV in 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. Currently, vaccination campaigns are underway 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. Rinderpest has not been reported from West or Central Africa for 10 years and, as stated above (see Overview, Distribution), the disease is now confined to two most insecure areas of eastern Africa.
References
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.
Barrett T, Rossiter PB, 1999. Rinderpest: the disease and its impact in man and humans. Advances in Virus Research, 53:89-110.
Mack R, 1970. The great African cattle plague epidemic of the 1890s. Tropical Animal Health and Production 2:210-219.
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
Plowright W, 1968. Rinderpest virus. Monographs in Virology, 3:25-110.
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.
Links to Websites
Website URL Comment
Office International des Epizooties http://www.oie.int
FAO -EMPRES http://www.fao.org/ag/AGA/AGAH/EMPRES/index.asp Information about the FAO initiative, Emergency Prevention System against transboundary animal and plant pests and diseases (EMPRES).
Institute for Animal Health http://www.iah.bbsrc.ac.uk
Images
Picture Title Caption Copyright
External symptoms Conjunctivitis and mucopurulent exudate in the early stages of RP infection. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
External symptoms Excessive salivation in the early stage of RP infection. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
External symptoms Oral erosions. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Sloughing of the epithelium over a necrotic Peyer's patch. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Ulcerations in the mucosa of the upper colon. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Hyperemia of the cecum and colon with accentuation of lesions (haemorrhage) at the ceco-colic junction. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Hyperemia and haemorrhages in the longitudinal folds of the colon. Zebra striping. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Haemorrhage in the mucosa of the gall bladder. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Date of report: 05/04/2011
© CAB International 2010
Animal Health and Production Compendium
Selected sections for: rinderpest virus
Identity Taxonomic Tree Disease/s Table Pathogen Characteristics Host Animals References
Datasheet Type(s): Pathogen
Identity
Preferred Scientific Name
rinderpest virus
International Common Names
English acronym
RPV
RV
Taxonomic Tree
Domain: Virus
Group: "Negative sense ssRNA viruses"
Group: "RNA viruses"
Order: Mononegavirales
Family: Paramyxoviridae
Genus: Morbillivirus
Species: rinderpest virus
Disease/s Table
rinderpest
Pathogen Characteristics
The complete genome sequence is available for rinderpest virus (RPV), human measles virus (MV) and canine distemper virus (CDV) and partial sequence data for the other morbilliviruses. They are identical in structure, being pleiomorphic, enveloped viruses of approximately 300-nm diameter with a typical paramyxovirus nucleocapsid structure. The virion is composed of six structural proteins and the genetic material of a single piece of RNA of negative-sense polarity contained in the nucleocapsid coiled within the virus envelope. Once the virus enters the cell, transcription of the negative-sense genome RNA begins and messenger RNAs are produced for each virus protein. Later in infection, replication of the virus RNA begins. This is accomplished by making a full-length positive-sense copy of the genome RNA, instead of the individual mRNAs, and this functions as the template to produce new virion RNA. Newly synthesized template and virion RNAs are surrounded and protected by the nucleocapsid protein (N protein) which, in association with the virus polymerase (L protein) and phosphoprotein (P protein), form ribonucleoprotein complexes called nucleocapsids.
The virus envelope contains two virus-coded glycoproteins, the haemagglutinin (H protein) and fusion proteins (F protein), responsible for attachment to and fusion with the host cell, respectively. The lipid layer of the virus envelope is derived from the host cell during virus budding. A non-glycosylated matrix protein (M protein) interacts both with the internal domains of the envelope glycoproteins and with the nucleocapsids formed within the host cell cytoplasm during genome replication. This brings the internal and external components together and enables the new virus to bud from the host cell. The virus genome RNA is approximately 16 kb in length and consists of a short 3' leader RNA followed by the coding regions for the six structural protein genes, with defined stop-start sequence motifs between each gene, and ends in a short 5' trailer RNA. The organization of the rinderpest genome is 3'Leader: 55; N 1692; P 1658; M 1463; F 2370; H 1961; L 6646; 5'Trailer 37; Total length 15882.
Two virus-encoded non-structural proteins (C and V) are produced in infected cells and are probably concerned with regulation of virus replication. The C non-structural protein is translated from an alternate reading frame in the phosphoprotein mRNA, beginning at the second AUG codon. The V protein is translated from an mRNA which is not an exact copy of the P gene sequence but from an edited mRNA which has an extra G residue inserted at a conserved slippery sequence motif positioned about halfway along the P gene where the virus polymerase 'stutters' and adds the extra non-templated G. This editing (or more properly, alternative transcription) occurs in about 30-50% of the mRNAs transcribed from the P gene in the case of MV, RPV and CDV and there is evidence that it occurs in all other morbilliviruses. Translation of this mRNA produces a chimeric protein consisting of the N-terminus of the P protein with a new C-terminus, rich in cysteine residues, derived from sequences in the third reading frame. The V protein is probably not required for growth of these viruses in tissue culture as mutations in the editing site of DMV did not affect the ability to grow in Vero cells.
Disease(s) associated with this pathogen is/are on the list of diseases notifiable to the World Organisation for Animal Health (OIE). The distribution section contains data from OIE's Handistatus database on disease occurrence. Please see the AHPC library for further information 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.
Host Animals
Animal name Context
Antilope cervicapra
Axis axis (Indian spotted deer)
Bos grunniens (yaks) Domesticated host, Wild host
Bos taurus (cattle) Domesticated host
Bubalus bubalis (buffalo)
Camelus dromedarius (dromedary camel) Domesticated host
Capra hircus (goats)
Capreolus capreolus
Oryctolagus cuniculus (rabbits)
Ovis aries (sheep) Domesticated host
Sus scrofa (pigs) Domesticated host
Syncerus caffer
Tragelaphus imberbis
Tragelaphus imberis
Tragelaphus oryx
References
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, 2005. World Animal Health Publication and Handistatus II (data set for 2004). Paris, France: Office International des Epizooties.
Date of report: 05/04/2011
© CAB International 2010
[[Category:To Do - CABI]]
Selected sections for: rinderpest
Identity Pathogen/s Overview Distribution Distribution Table Hosts/Species Affected Host Animals Systems Affected List of Symptoms/Signs Epidemiology Zoonoses and Food Safety Pathology Diagnosis Disease Course Disease Treatment Prevention and Control References Links to Websites Images
Datasheet Type(s): Animal Disease
Identity
Preferred Scientific Name
rinderpest
International Common Names
English acronym
RP
English
cattle plague, rinderpest in pigs- exotic, rinderpest in ruminants- exotic
French
peste bovine
Arabic
taoun
Local Common Names
East Africa
olodua
Pathogen/s
rinderpest virus
Overview
Rinderpest is an acute to subacute contagious viral disease of ruminants and pigs 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) infecting cattle and other large ruminants, peste des petits ruminants virus (PPRV) infecting sheep and goats, canine distemper virus (CDV) which infects carnivores, human measles virus (MV), and other members in marine mammals. Members of the genus are closely related antigenically and are distinguished from the other paramyxoviruses by their lack of neuraminidase activity.
In terms of economic losses in domestic animals, rinderpest is the most important member of the group. In the world today it is an infection of extensive pastoral cattle herds in two areas of Africa and in some concentrations of inadequately vaccinated cattle and buffalo herds in Pakistan.
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 Handistatus 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.
Distribution
Rinderpest is found in localised regions of Pakistan, Somalia and Sudan. Its presence is uncertain in Afghanistan, Yemen and Kenya.
Distribution Table
Country Distribution Last Reported Origin First Reported Invasive References Notes
ASIA
Afghanistan Disease not reported OIE, 2009
Armenia Disease not reported OIE, 2009
Azerbaijan Disease not reported OIE, 2009
Bahrain Disease not reported OIE, 2009
Bangladesh Disease not reported NULL OIE, 2009; OIE, 2003
Bhutan Disease not reported OIE, 2009
Brunei Darussalam Disease not reported OIE Handistatus, 2005
Cambodia Disease not reported OIE, 2009
China No information available OIE, 2009
-Hong Kong Disease not reported OIE, 2009
Georgia (Republic of) Last reported 1989 OIE Handistatus, 2005
India Disease not reported OIE, 2009
Indonesia Disease not reported OIE, 2009
Iran Disease not reported OIE, 2009
Iraq Disease not reported OIE, 2009
Israel Disease not reported OIE, 2009
Japan Disease not reported OIE, 2009
Jordan Disease not reported OIE, 2009
Kazakhstan Disease not reported OIE, 2009
Korea, DPR Last reported 1948 OIE Handistatus, 2005
Korea, Republic of Disease not reported OIE, 2009
Kuwait Disease not reported OIE, 2009
Kyrgyzstan Disease never reported OIE, 2009
Laos Disease not reported OIE, 2009
Lebanon Disease not reported OIE, 2009
Malaysia Disease not reported OIE, 2009
-Peninsular Malaysia Disease never reported OIE Handistatus, 2005
-Sabah Disease never reported OIE Handistatus, 2005
-Sarawak Disease never reported OIE Handistatus, 2005
Mongolia Disease not reported OIE, 2009
Myanmar Disease not reported OIE, 2009
Nepal Disease not reported OIE, 2009
Oman Disease not reported OIE, 2009
Pakistan Disease not reported OIE, 2009
Philippines Disease not reported OIE, 2009
Qatar Disease not reported OIE, 2009
Saudi Arabia Disease not reported OIE, 2009
Singapore Disease not reported OIE, 2009
Sri Lanka Disease not reported OIE, 2009
Syria Disease not reported OIE, 2009
Taiwan Last reported 1950 OIE Handistatus, 2005
Tajikistan Disease never reported OIE, 2009
Thailand Disease not reported OIE, 2009
Turkey Disease not reported OIE, 2009
Turkmenistan Disease never reported OIE Handistatus, 2005
United Arab Emirates Disease not reported OIE, 2009
Uzbekistan Disease never reported OIE Handistatus, 2005
Vietnam Disease not reported OIE, 2009
Yemen No information available OIE, 2009
AFRICA
Algeria Disease never reported OIE, 2009
Angola Disease not reported OIE, 2009
Benin Disease not reported OIE, 2009
Botswana Disease not reported OIE, 2009
Burkina Faso Disease not reported OIE, 2009
Burundi Disease not reported OIE Handistatus, 2005
Cameroon Last reported 1986 OIE Handistatus, 2005
Cape Verde No information available OIE Handistatus, 2005
Central African Republic Last reported 1983 OIE Handistatus, 2005
Chad Disease not reported OIE, 2009
Congo Disease never reported OIE, 2009
Congo Democratic Republic Disease not reported OIE Handistatus, 2005
Côte d'Ivoire Last reported 1986 OIE Handistatus, 2005
Djibouti No information available OIE, 2009
Egypt Disease not reported OIE, 2009
Eritrea No information available OIE, 2009
Ethiopia Disease not reported OIE, 2009
Gabon Disease never reported OIE, 2009
Gambia Disease not reported OIE, 2009
Ghana Disease not reported OIE, 2009
Guinea Disease not reported OIE, 2009
Guinea-Bissau Disease not reported OIE, 2009
Kenya Disease not reported 2003 OIE, 2009; OIE, 2003
Lesotho Disease not reported OIE, 2009
Libya Last reported 1966 OIE Handistatus, 2005
Madagascar Disease never reported OIE, 2009
Malawi Disease never reported OIE, 2009
Mali No information available OIE, 2009
Mauritius Disease never reported OIE, 2009
Morocco Disease never reported OIE, 2009
Mozambique Disease not reported OIE, 2009
Namibia Disease not reported OIE, 2009
Niger Last reported 1985 OIE Handistatus, 2005
Nigeria Disease not reported OIE, 2009
Réunion Last reported 1902 OIE Handistatus, 2005
Rwanda Disease not reported OIE, 2009
Sao Tome and Principe Disease not reported OIE Handistatus, 2005
Senegal Disease not reported OIE, 2009
Seychelles Disease not reported OIE Handistatus, 2005
Somalia No information available OIE Handistatus, 2005
South Africa Disease not reported OIE, 2009
Sudan Disease not reported OIE, 2009
Swaziland Disease not reported OIE, 2009
Tanzania Disease not reported OIE, 2009
Togo Disease not reported OIE, 2009
Tunisia Disease never reported OIE, 2009
Uganda Disease not reported OIE, 2009
Zambia Disease not reported OIE, 2009
Zimbabwe Disease not reported OIE, 2009
NORTH AMERICA
Bermuda Disease not reported OIE Handistatus, 2005
Canada Disease never reported OIE, 2009
Greenland Disease never reported OIE, 2009
Mexico Disease not reported OIE, 2009
USA Disease never reported OIE, 2009
-Georgia Disease not reported OIE, 2009
CENTRAL AMERICA
Barbados Disease never reported OIE Handistatus, 2005
Belize Disease never reported OIE, 2009
British Virgin Islands Disease never reported OIE Handistatus, 2005
Cayman Islands Disease never reported OIE Handistatus, 2005
Costa Rica Disease never reported OIE, 2009
Cuba Disease never reported OIE, 2009
Curaçao Disease not reported OIE Handistatus, 2005
Dominica Disease not reported OIE Handistatus, 2005
Dominican Republic Disease never reported OIE, 2009
El Salvador Disease never reported OIE, 2009
Guadeloupe Disease never reported OIE, 2009
Guatemala Disease never reported OIE, 2009
Haiti Disease never reported OIE, 2009
Honduras Disease never reported OIE, 2009
Jamaica No information available OIE, 2009
Martinique Disease never reported OIE, 2009
Nicaragua Disease never reported OIE, 2009
Panama Disease never reported OIE, 2009
Saint Kitts and Nevis Disease never reported OIE Handistatus, 2005
Saint Vincent and the Grenadines Disease never reported OIE Handistatus, 2005
Trinidad and Tobago Disease never reported OIE Handistatus, 2005
SOUTH AMERICA
Argentina Disease never reported OIE, 2009
Bolivia Disease never reported OIE, 2009
Brazil Disease not reported OIE, 2009
Chile Disease never reported OIE, 2009
Colombia Disease never reported OIE, 2009
Ecuador Disease never reported OIE, 2009
Falkland Islands Disease never reported OIE Handistatus, 2005
French Guiana Disease never reported OIE, 2009
Guyana Disease never reported OIE Handistatus, 2005
Paraguay Disease never reported OIE Handistatus, 2005
Peru Disease never reported OIE, 2009
Uruguay Disease never reported OIE, 2009
Venezuela Disease never reported OIE, 2009
EUROPE
Albania Disease not reported OIE, 2009
Andorra Disease never reported OIE Handistatus, 2005
Austria Disease not reported OIE, 2009
Belarus Disease never reported OIE, 2009
Belgium Disease not reported OIE, 2009
Bosnia-Hercegovina Last reported 1883 OIE Handistatus, 2005
Bulgaria Disease not reported OIE, 2009
Croatia Disease not reported OIE, 2009
Cyprus Disease never reported OIE, 2009
Czech Republic Disease not reported OIE, 2009
Denmark Disease not reported OIE, 2009
Estonia Disease never reported OIE, 2009
Finland Disease not reported OIE, 2009
France Disease not reported OIE, 2009
Germany Disease not reported OIE, 2009
Greece Disease not reported OIE, 2009
Hungary Disease not reported OIE, 2009
Iceland Disease never reported OIE, 2009
Ireland Disease not reported OIE, 2009
Isle of Man (UK) Disease never reported OIE Handistatus, 2005
Italy Disease not reported OIE, 2009
Jersey Disease never reported OIE Handistatus, 2005
Latvia Disease not reported OIE, 2009
Liechtenstein Disease not reported OIE, 2009
Lithuania Disease never reported OIE, 2009
Luxembourg Disease never reported OIE, 2009
Macedonia Disease never reported OIE, 2009
Malta Disease never reported OIE, 2009
Moldova Disease never reported OIE Handistatus, 2005
Montenegro Disease not reported OIE, 2009
Netherlands Disease not reported OIE, 2009
Norway Disease never reported OIE, 2009
Poland Disease not reported OIE, 2009
Portugal Disease never reported OIE, 2009
Romania Disease not reported OIE, 2009
Russian Federation Disease not reported OIE, 2009
Serbia Disease not reported OIE, 2009
Slovakia Disease not reported OIE, 2009
Slovenia Disease not reported OIE, 2009
Spain Disease never reported OIE, 2009
Sweden Disease not reported OIE, 2009
Switzerland Disease not reported OIE, 2009
Ukraine Disease never reported OIE, 2009
United Kingdom
-Northern Ireland Last reported 1900 OIE Handistatus, 2005
United Kingdom Disease not reported OIE, 2009
Yugoslavia (former) Disease never reported OIE Handistatus, 2005
Yugoslavia (Serbia and Montenegro) Last reported 1883 OIE Handistatus, 2005
OCEANIA
Australia Disease not reported OIE, 2009
French Polynesia Disease never reported OIE, 2009
New Caledonia Disease never reported OIE, 2009
New Zealand Disease never reported OIE, 2009
Samoa Disease never reported OIE Handistatus, 2005
Vanuatu Disease never reported OIE Handistatus, 2005
Wallis and Futuna Islands No information available OIE Handistatus, 2005
Hosts/Species Affected
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. 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.
Host Animals
Animal name Context
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
Sus scrofa (pigs) Domesticated host
Systems Affected
Digestive - Large Ruminants
Digestive - Pigs
Digestive - Small Ruminants
Multisystem - Large Ruminants
Multisystem - Pigs
Multisystem - Small Ruminants
List of Symptoms/Signs
Sign Life Stages Type
Digestive Signs
Mucous, mucoid stools, faeces Sign [C]
Bloody stools, faeces, haematochezia Sign [C]
Grinding teeth, bruxism, odontoprisis Sign [C]
Rumen hypomotility or atony, decreased rate, motility, strength Sign [C]
Decreased amount of stools, absent faeces, constipation Sign [C]
Pharyngeal ulcers, vesicles, erosion, papules, sores pharynx C1 ( All Stages ) Diagnosis
Diarrhoea C1 ( All Stages ) Diagnosis [C]
Unusual or foul odor, stools, faeces C1 ( All Stages ) Diagnosis [C]
Congestion oral mucous membranes, erythema, redness oral mucosa C1 ( All Stages ) Diagnosis [C]
Oral mucosal ulcers, vesicles, plaques, pustules, erosions, tears C1 ( All Stages ) Diagnosis [C]
Tongue ulcers, vesicles, erosions, sores, blisters, cuts, tears C1 ( All Stages ) Diagnosis [C]
Excessive salivation, frothing at the mouth, ptyalism Sign [C]
Anorexia, loss or decreased appetite, not nursing, off feed C1 ( All Stages ) Sign [C]
General Signs
Fever, pyrexia, hyperthermia C1 ( All Stages ) Sign [C]
Trembling, shivering, fasciculations, chilling Sign [C]
Trembling, shivering, fasciculations, chilling Sign [C]
Petechiae or ecchymoses, bruises, ecchymosis Sign [C]
Generalized weakness, paresis, paralysis Sign [C]
Inability to stand, downer, prostration Sign [C]
Tenesmus, straining, dyschezia Sign [C]
Hypothermia, low temperature Sign [C]
Dehydration Sign [C]
Polydipsia, excessive fluid consumption, excessive thirst Sign [C]
Nervous Signs
Tremor Sign [C]
Dullness, depression, lethargy, depressed, lethargic, listless Sign [C]
Ophthalmology Signs
Conjunctival, scleral, injection, abnormal vasculature Sign [C]
Conjunctival, scleral, redness C1 ( All Stages ) Diagnosis [C]
Lacrimation, tearing, serous ocular discharge, watery eyes C1 ( All Stages ) Diagnosis [C]
Blindness L1 ( All Stages ) Diagnosis
Purulent discharge from eye C1 ( All Stages ) Diagnosis [C]
Photophobia Sign [C]
Blepharospasm Sign [C]
Pain/Discomfort Signs
Colic, abdominal pain Sign [C]
Skin pain Sign [C]
Reproductive Signs
Agalactia, decreased, absent milk production Sign [C]
Vulval ulcers, vesicles, erosions, tears, cuts, pustules, papules Sign [C]
Vaginal or cervical ulcers, vesicles, erosions, tears, papules, pustules C3 ( Heifer ), C4 ( Cow ) Diagnosis
Abortion or weak newborns, stillbirth C3 ( Heifer ), C4 ( Cow ) Diagnosis [C]
Respiratory Signs
Nasal mucosal ulcers, vesicles, erosions, cuts, tears, papules, pustules C1 ( All Stages ) Diagnosis [C]
Purulent nasal discharge C1 ( All Stages ) Diagnosis [C]
Mucoid nasal discharge, serous, watery C1 ( All Stages ) Diagnosis [C]
Abnormal breath odor, foul odor mouth C1 ( All Stages ) Diagnosis [C]
Dyspnea, difficult, open mouth breathing, grunt, gasping Sign [C]
Increased respiratory rate, polypnea, tachypnea, hyperpnea Sign [C]
Coughing, coughs Sign [C]
Skin/Integumentary Signs
Alopecia, thinning, shedding, easily epilated, loss of, hair Sign [C]
Rough hair coat, dull, standing on end Sign [C]
Skin erythema, inflammation, redness L1 ( All Stages ) Sign [C]
Skin papules L1 ( All Stages ) Sign [C]
Skin pustules Sign [C]
Skin crusts, scabs Sign [C]
Moist skin, hair or feathers Sign [C]
Urinary Signs
Polyuria, increased urine output Sign [C]
Epidemiology
Rinderpest virus (RV) infects 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 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. 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 RV is effected 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 RV 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, RV 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 is 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 behaves in epidemic fashion with the virus infecting virtually all susceptible individuals and causing severe clinical disease in most age groups. Endemic rinderpest, however, is much milder and is maintained by young animals usually less than 2 years old that have lost their maternal immunity. Intermediate patterns also exist.
Wildlife play 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.
Zoonoses and Food Safety
This disease is not a zoonosis.
Pathology
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 RV 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.
Diagnosis
Clinical Signs
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 are currently circulating in eastern Africa. 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. A presumptive diagnosis of rinderpest can be made on the basis of the clinical signs and gross pathology. However, in countries where the disease is not prevalent, and especially in regions dependent on livestock exports, it is essential to obtain laboratory confirmation of the diagnosis as soon as possible. Countries where rinderpest is either endemic or a high risk should treat any syndrome resembling rinderpest as such until proven otherwise. This will allow immediate steps to be taken to control the disease and restrict losses.
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 RV. The use of anti-proteases increases the survival of RV antigens in tissue suspensions and reduces the degradation of RNA.
Laboratory Diagnosis
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.
The first procedure usually carried out is to detect viral antigen using specific rabbit hyperimmune serum against RV. The most commonly used assay is the agar-gel immunodiffusion test (AGID) which is simple, easy to read, and highly specific. Moreover, 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. Counter-immunoelectrophoresis is quicker and more sensitive than AGID but requires more sophisticated equipment. Immunofluorescence and immunoperoxidase staining are very sensitive but also need more equipment than AGID. Although once widely used, complement fixation and conglutinating complement absorption tests are too complicated in comparison with more recently developed tests. Various haemagglutination assays are sensitive but not yet widely applied though latex bead agglutination tests have given encouraging preliminary results and, if combined with monoclonal antibodies, might prove very sensitive. If classically prepared rabbit hyperimmune sera are unavailable, sera can be prepared using other immunizing techniques in rabbits or in goats or cattle. A positive test result in any of the tests confirms rinderpest.
Where cell cultures are unavailable, specimens can 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. Vero cells are also suitable, whereas culture systems such as microplates can also be used but may be less sensitive. After 12-24 hours adsorption the tubes are washed, re-fed with maintenance medium and rolled 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 to RV or by examining fixed monolayers using immunofluorescent or immunoperoxidase techniques.
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 are 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 that use monoclonal antibodies to different viral antigens such as the H or N protein, and may also use purified or recombinant antigens. The ELISA has the advantage that laboratories without cell-culture can test thousands of sera, which is often required in current eradication programmes, and the sensitivity and specificity of these new tests is under validation at present. During the early antibody response, serum contains significant levels of IgM to RV, 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 are capable of detecting minute quantities of RV RNA in tissues and secretions, and are now often a routine choice for confirmation in reference laboratories. The PCR offers the advantage of 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 specific monoclonal antibody based latex bead agglutination is being developed for use with rinderpest.
Differential Diagnosis
All conditions that cause stomatitis and/or enteritis in domestic stock can be clinically confused with rinderpest. In cattle, difficulties may 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 present 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 can rapidly differentiate the two diseases. Immunohistochemical techniques can 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 must 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 RV and is difficult to differentiate with hyperimmune polyclonal sera. Fortunately, contemporary studies have produced monoclonal antibodies and nucleic-acid techniques that clearly distinguish between PPR virus and RV, at least for the limited number of strains tested to date. In African countries that have previously been free of PPR it is unwise to assume that a rinderpest-like syndrome in small ruminants is not PPR.
Immune Response
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.
Disease Course
Very few sequence changes are needed to alter the virulence of rinderpest virus; the genome of the cell-culture 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 means that is now possible to address questions concerning virus attenuation and pathogenicity by directly altering virus genes. Site-specific mutagenesis, swapping of genes between mild and virulent isolates and the insertion/deletion of genes will lead to an understanding of factors which determine the host range, tissue tropism and pathology of the virus. This technology will also help in the control of rinderpest by providing genetically defined vaccines, which will enable vaccinated animals to be distinguished from those that have been naturally infected.
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 RV 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 RV, 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.
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 RV 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, implying that recovered animals perhaps may be continually re-stimulated immunologically by RV antigen throughout their lives.
The massive destruction of lymphocytes causes immunosuppression, probably involving both cell-mediated and humoral immune responses.
Disease Treatment
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.
Prevention and Control
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 is 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 are associated with delivering RPV vaccine, though it has been used with success to control RPV in 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. Currently, vaccination campaigns are underway 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. Rinderpest has not been reported from West or Central Africa for 10 years and, as stated above (see Overview, Distribution), the disease is now confined to two most insecure areas of eastern Africa.
References
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.
Barrett T, Rossiter PB, 1999. Rinderpest: the disease and its impact in man and humans. Advances in Virus Research, 53:89-110.
Mack R, 1970. The great African cattle plague epidemic of the 1890s. Tropical Animal Health and Production 2:210-219.
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
Plowright W, 1968. Rinderpest virus. Monographs in Virology, 3:25-110.
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.
Links to Websites
Website URL Comment
Office International des Epizooties http://www.oie.int
FAO -EMPRES http://www.fao.org/ag/AGA/AGAH/EMPRES/index.asp Information about the FAO initiative, Emergency Prevention System against transboundary animal and plant pests and diseases (EMPRES).
Institute for Animal Health http://www.iah.bbsrc.ac.uk
Images
Picture Title Caption Copyright
External symptoms Conjunctivitis and mucopurulent exudate in the early stages of RP infection. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
External symptoms Excessive salivation in the early stage of RP infection. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
External symptoms Oral erosions. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Sloughing of the epithelium over a necrotic Peyer's patch. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Ulcerations in the mucosa of the upper colon. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Hyperemia of the cecum and colon with accentuation of lesions (haemorrhage) at the ceco-colic junction. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Hyperemia and haemorrhages in the longitudinal folds of the colon. Zebra striping. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Pathology Haemorrhage in the mucosa of the gall bladder. USDA, 2002_ Foreign Animal Diseases Training Set_ USDA - Animal _ Plant Health Inspection Service
Date of report: 05/04/2011
© CAB International 2010
Animal Health and Production Compendium
Selected sections for: rinderpest virus
Identity Taxonomic Tree Disease/s Table Pathogen Characteristics Host Animals References
Datasheet Type(s): Pathogen
Identity
Preferred Scientific Name
rinderpest virus
International Common Names
English acronym
RPV
RV
Taxonomic Tree
Domain: Virus
Group: "Negative sense ssRNA viruses"
Group: "RNA viruses"
Order: Mononegavirales
Family: Paramyxoviridae
Genus: Morbillivirus
Species: rinderpest virus
Disease/s Table
rinderpest
Pathogen Characteristics
The complete genome sequence is available for rinderpest virus (RPV), human measles virus (MV) and canine distemper virus (CDV) and partial sequence data for the other morbilliviruses. They are identical in structure, being pleiomorphic, enveloped viruses of approximately 300-nm diameter with a typical paramyxovirus nucleocapsid structure. The virion is composed of six structural proteins and the genetic material of a single piece of RNA of negative-sense polarity contained in the nucleocapsid coiled within the virus envelope. Once the virus enters the cell, transcription of the negative-sense genome RNA begins and messenger RNAs are produced for each virus protein. Later in infection, replication of the virus RNA begins. This is accomplished by making a full-length positive-sense copy of the genome RNA, instead of the individual mRNAs, and this functions as the template to produce new virion RNA. Newly synthesized template and virion RNAs are surrounded and protected by the nucleocapsid protein (N protein) which, in association with the virus polymerase (L protein) and phosphoprotein (P protein), form ribonucleoprotein complexes called nucleocapsids.
The virus envelope contains two virus-coded glycoproteins, the haemagglutinin (H protein) and fusion proteins (F protein), responsible for attachment to and fusion with the host cell, respectively. The lipid layer of the virus envelope is derived from the host cell during virus budding. A non-glycosylated matrix protein (M protein) interacts both with the internal domains of the envelope glycoproteins and with the nucleocapsids formed within the host cell cytoplasm during genome replication. This brings the internal and external components together and enables the new virus to bud from the host cell. The virus genome RNA is approximately 16 kb in length and consists of a short 3' leader RNA followed by the coding regions for the six structural protein genes, with defined stop-start sequence motifs between each gene, and ends in a short 5' trailer RNA. The organization of the rinderpest genome is 3'Leader: 55; N 1692; P 1658; M 1463; F 2370; H 1961; L 6646; 5'Trailer 37; Total length 15882.
Two virus-encoded non-structural proteins (C and V) are produced in infected cells and are probably concerned with regulation of virus replication. The C non-structural protein is translated from an alternate reading frame in the phosphoprotein mRNA, beginning at the second AUG codon. The V protein is translated from an mRNA which is not an exact copy of the P gene sequence but from an edited mRNA which has an extra G residue inserted at a conserved slippery sequence motif positioned about halfway along the P gene where the virus polymerase 'stutters' and adds the extra non-templated G. This editing (or more properly, alternative transcription) occurs in about 30-50% of the mRNAs transcribed from the P gene in the case of MV, RPV and CDV and there is evidence that it occurs in all other morbilliviruses. Translation of this mRNA produces a chimeric protein consisting of the N-terminus of the P protein with a new C-terminus, rich in cysteine residues, derived from sequences in the third reading frame. The V protein is probably not required for growth of these viruses in tissue culture as mutations in the editing site of DMV did not affect the ability to grow in Vero cells.
Disease(s) associated with this pathogen is/are on the list of diseases notifiable to the World Organisation for Animal Health (OIE). The distribution section contains data from OIE's Handistatus database on disease occurrence. Please see the AHPC library for further information 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.
Host Animals
Animal name Context
Antilope cervicapra
Axis axis (Indian spotted deer)
Bos grunniens (yaks) Domesticated host, Wild host
Bos taurus (cattle) Domesticated host
Bubalus bubalis (buffalo)
Camelus dromedarius (dromedary camel) Domesticated host
Capra hircus (goats)
Capreolus capreolus
Oryctolagus cuniculus (rabbits)
Ovis aries (sheep) Domesticated host
Sus scrofa (pigs) Domesticated host
Syncerus caffer
Tragelaphus imberbis
Tragelaphus imberis
Tragelaphus oryx
References
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, 2005. World Animal Health Publication and Handistatus II (data set for 2004). Paris, France: Office International des Epizooties.
Date of report: 05/04/2011
© CAB International 2010
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