Selected content from the Animal Health and Production Compendium (© CAB International 2013). Distributed under license by African Union – Interafrican Bureau for Animal Resources.
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Identity Pathogen/s Overview Distribution Distribution Map for Africa Distribution Table for Africa Hosts/Species Affected Host Animals Systems Affected Epidemiology Impact: Economic Zoonoses and Food Safety Pathology Diagnosis Disease Course Disease Treatment Table Disease Treatment Vaccines Prevention and Control References Links to Websites OIE Reference Experts and Laboratories Images
Preferred Scientific Name
peste des petits ruminants
International Common Names
goat plague, pest of sheep and goats, pest of small ruminants, pneumoenteritis complex, pseudorinderpest of small ruminants, stomatitis-pneumoenteritis syndrome
Local Common Names
peste des petits ruminants virus
The disease was first described by workers in the Côté d'Ivoire (Gargadennec and Lalanne, 1942), and thereafter in other parts of West Africa in the 1950s and 1960s. It is now recognized to be distributed from Tibet through the Middle East, and through most of Africa, From East to West, and from North Africa (Algeria, Tunisia, Morocco) south as far as Tanzania. The similarity in clinical signs to that of rinderpest in cattle probably accounts for the number of reports of rinderpest in small ruminants from some countries, and delayed the recognition of the disease as a distinct entity in India until the early 1990s. The infective agent was first considered a variant of rinderpest virus adapted to small ruminants, but was later shown to be antigenically (Gibbs et al., 1979) and genetically distinct (Diallo et al., 1989). Although it had been previously proposed that PPR virus (PPRV) emerged from rinderpest virus (RPV) more recently than RPV diverged from measles virus and canine distemper virus, comparison of gene sequences suggest PPRV is no more related to RPV than to non-ruminant morbilliviruses (Das et al., 2000; Baron, 2011). Since PPRV isolates of West African, Middle Eastern and south Asian origin comprise distinct genetic groups, it is likely that the infections have circulated largely independently for long periods in each area.
The disease is recognized by the World Organisation for Animal Health (in French the Office International des Epizooties and still abbreviated as OIE) as a notifiable disease on account of the high mortality and morbidity, and rapidity of spread by contagion. Recognition of PPR as a problem has increased in the 1990s, partly as a result of surveillance activities of the global rinderpest eradication programme (GREP), but also because of the capacity of the infection to invade disease-free countries. The presence of infection restricts international trade in livestock and livestock products from infected countries, and is usually associated with ongoing severe losses where conditions exist that support epidemic spread among susceptible breeds, such as the incursions of infection into 'marginal' zones for persistence of infection such as humid zones of West Africa. Under such conditions there is a high social impact of disease, since small ruminants often represent a readily convertible currency in smallholder agriculture. Control by vaccination is merited in many endemic countries, and the benefit-to-cost ratio is usually high. The disease was also important as a complication in the eradication of rinderpest, since PPRV can infect cattle and the resultant antibodies must be distinguished from those elicited by rinderpest. The use of rinderpest vaccine to protect against PPR is no longer permitted now that rinderpest has been eradicated, to avoid the possibility of RP antibody detection in cattle or small ruminants. An attenuated strain of PPRV for use as a vaccine was developed in the late 1980's (Diallo et al., 1989b) and is in wide use.
This disease is on the list of diseases notifiable to OIE. Please see the AHPC library for further information from OIE on this disease, including the International Animal Health Code and the Manual of Standards for Diagnostic Tests and Vaccines. Also see the website: www.oie.int.
The disease is recognized to be distributed from Tibet through the Middle East, and through most of Africa, From East to West, and from North Africa (Algeria, Tunisia, Morocco) south as far as Tanzania. For current information on disease incidence, see OIE's WAHID database.
The development of tests for the confirmation of the agent and the differentiation from rinderpest was only achieved in the late 1980s, and recent recognition of the infection in many African and Asian countries can be attributed to availability of methods for specific detection of virus and antibodies. However, given the rapidity of spread of epidemics and the lack of a carrier state, it is likely that infection is not maintained in some areas because of inadequate supply of susceptible animals and some incursions of disease appear self-limiting. Presence of infection in a country in one year does not imply endemic infection. However, the risk of trans-boundary spread of this disease is high because sheep and goats are easily transported and trade across borders is difficult to control. Recent extension of infection into China and a number of countries in Africa previously free of the disease (e.g. Kenya, Tanzania and several countries north of the Sahara) highlights the risk to previously free areas.
The number of countries in Africa reporting peste des petits ruminants (PPR) outbreaks to the African Union - Interafrican Bureau for Animal Resources (AU-IBAR; AU-IBAR, 2011) has increased from 19 in 2008 to 20 in 2009, 25 in 2010 and 27 in 2011 as shown in the table below. Although the west, central and eastern Africa regions are regarded largely as endemic foci for PPR, the disease has been showing geographical advances towards the southern and northern regions of Africa; with Tanzania (2008) and Zambia (2010) in the south and Algeria (2011) in the north becoming the most recently affected countries on the continent.
Out of the 27 countries that reported the disease in 2011, the majority also recorded the disease during the past three years. In total, 1185 epidemiological units were affected by PPR in 27 countries causing 101,016 cases and 62,388 deaths with a case fatality rate of 61.8%. The top three countries with highest number of outbreaks in descending order are Benin (285), Ghana (184) and Nigeria (126), all in West Africa.
There appears to be no defined temporal trend for the appearance of the disease as it occurs virtually uniformly throughout the year.
= Present, no further details = Widespread = Localised
= Confined and subject to quarantine = Occasional or few reports
= Evidence of pathogen = Last reported... = Presence unconfirmed
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further information for individual references may be available in the Animal Health and Production Compendium. A table for worldwide distribution can also be found in the Animal Health and Production Compendium.
|Country||Distribution||Last Reported||Origin||First Reported||Invasive||References||Notes|
|Algeria||Last reported||2012||OIE, 2012|
|Angola||Unconfirmed record||OIE, 2012|
|Botswana||Disease never reported||OIE, 2012|
|Burkina Faso||Present||OIE, 2012|
|Burundi||Disease not reported||OIE Handistatus, 2005|
|Cameroon||OIE Handistatus, 2005|
|Cape Verde||Disease never reported||OIE Handistatus, 2005|
|Central African Republic||Reported present or known to be present||OIE Handistatus, 2005|
|Chad||No information available||OIE, 2009|
|Congo Democratic Republic||Reported present or known to be present||OIE Handistatus, 2005|
|Côte d'Ivoire||Reported present or known to be present||Native||OIE, 2004; OIE Handistatus, 2005|
|Djibouti||Disease not reported||OIE, 2009|
|Egypt||Disease not reported||OIE, 2009|
|Gabon||Disease not reported||OIE, 2009|
|Lesotho||Disease never reported||OIE, 2012|
|Libya||Disease never reported||OIE, 2012|
|Madagascar||Disease never reported||OIE, 2012|
|Malawi||Disease never reported||OIE, 2012|
|Mali||Present||AU-IBAR, 2011; OIE, 2001a; OIE, 2009|
|Mauritania||Restricted distribution||OIE, 2012|
|Mauritius||Disease never reported||OIE, 2009|
|Mozambique||Disease never reported||OIE, 2012|
|Namibia||Disease never reported||OIE, 2012|
|Niger||Restricted distribution||OIE, 2012|
|Réunion||Disease never reported||OIE Handistatus, 2005|
|Sao Tome and Principe||Disease not reported||OIE Handistatus, 2005|
|Seychelles||Disease not reported||OIE Handistatus, 2005|
|Sierra Leone||Present||OIE, 2012|
|South Africa||Disease never reported||OIE, 2012|
|Swaziland||Disease never reported||OIE, 2012|
|Zimbabwe||Disease never reported||OIE, 2012|
Disease occurs in goats and sheep, and has been recognized in wild ungulates from families of Gazellinae (Dorcas gazelle), Caprinae (Nubian ibex and Laristan sheep) and Hippotraginae (gemsbok) and other wild sheep/goats (Baron, 2011). Experimentally, severe disease also occurs in white-tailed deer. Cattle undergo mainly subclinical reactions, and pigs develop a viraemia. PPRV has been detected in diseased camels (Roger et al., 2001; Khalafalla et al., 2010; Kwiatek et al., 2011). West African Dwarf goats are more susceptible than European breeds (Scott, 1990), and sheep and goats of the Sahelian zones are more resistant than dwarf types from humid zones to the south (Bourdin, 1983). Seasonality in breeding, in marketing and in crop production all appear related to disease occurrence. In arid and semi-arid zones, surplus animals are sold during the dry season, which may lead to spread of infection into other areas, especially if sale occurs via markets. In humid zones, sale of surplus animals may occur at the start of the rains, with tethering of animals to avoid crop damage. Both sale of animals and close housing/tethering can increase risk of transmission if virus is introduced. The wet season can also predispose to secondary bacterial infections, exacerbating the viral pneumonia.
|Bos indicus (zebu)||Domesticated host|
|Bubalus bubalis (buffalo)|
|Camelus dromedarius (dromedary camel)|
|Capra hircus (goats)||Domesticated host, Wild host|
|Ovis aries (sheep)||Domesticated host, Wild host|
Digestive - Large Ruminants
Digestive - Small Ruminants
Respiratory - Large Ruminants
Respiratory - Small Ruminants
Skin - Large Ruminants
Skin - Small Ruminants
PPRV has a direct life cycle, maintained by infected animal to susceptible animal transmission, without involvement of carrier animals or vectors. The underlying requirement is a regular supply of susceptible hosts plus sufficient animal movement to allow mixing of the population (Rossiter and Taylor, 1993). Risk factors include unconfined husbandry, whether in urban or rural settings. The population size required to maintain infection is not known, but small populations probably cannot maintain infection for long and, therefore, given the rapid population turnover of small ruminants, disease associated with re-introduction can be severe. Severe epidemics probably reflect introduction of virus into areas with mainly susceptible populations and breeds, rather than endemic presence of infection throughout the year (Rossiter and Taylor, 1993). Re-introduction of infection at intervals appear frequently related to livestock movement into areas, for example introduction for slaughter at festivals (Bonniwell, 1980). There is one serotype of virus, and immunity is long-lasting, probably life-long. Because colostral antibody protects young animals, in endemic areas most disease occurs in animals after waning of colostral immunity, from 4 months to 2 years of age, with protection of animals which have had previous exposure. Serological studies indicate that many, if not most, infections are subclinical or insufficiently severe to attract attention, and this may be related to breed resistance. Mortality rate in areas considered endemic may be in the region of 4-5% whereas rates from 20% to 90% in outbreaks have frequently been reported under epidemic conditions (Rossiter and Taylor, 1993). Climate appears to play a role in increasing the severity of disease, via secondary infections exacerbating the infection of the lung, but also affects the migration of pastoralist flocks in response to rainfall and drought and the resultant exposure to infection.
The virus will infect a range of ruminant hosts, but appears to cause disease in only goats, sheep, certain antelope species and occasionally water buffalo (Govindarajan et al, 1997). The virus will infect cattle and pigs, but subclinically; potentially these species might be involved in transmission between small-ruminant populations, although evidence that this is important in the field is lacking. Infection of cattle with PPRV protects against rinderpest virus, and the exposure rates of cattle in West Africa to PPRV may have contributed to a failure to induce antibodies to RPV after vaccination (Anderson and McKay, 1994), and also to the rapidity of eradication efforts against RPV in this region. In the same way, antibodies to RPV can protect sheep/goats from PPRV (Taylor, 1979), and it is possible that the recent rapid spread of PPR may be due to the eradication of RP and the cessation of vaccination against this disease. Antibodies to RPV can be distinguished from those to PPRV in cattle by use of monoclonal antibody based tests. PPR virus isolates differ in their pathogenicity for each host, with some strains resulting in more disease in sheep than goats. However, the outcome of infection is also markedly affected by animal breed, with some breeds from Sahelian countries showing marked resistance in contrast to those of West African humid zones (Bourdin, 1983).
The annual toll of infection in Nigeria in 1976 was US$1.5 million (Hamdy et al., 1976). The disease contributes to loss of the live export trade in sheep and goats because PPR is a list A disease; however, in most countries where it occurs other list A diseases would also require to be eradicated or controlled before PPRV was the limiting constraint to trade. Stem (1993) estimated a US$24 million return on an investment of US$2 million in PPR control in Niger over a 5-year period. Awa et al. (2000) also indicated benefit-to-cost ratios of 2.26 to 4.23 for a PPR vaccination and strategic anthelminthic programme in Cameroon.
PPR virus has not been shown to present any risk to human health. However, the carcasses of animals suffering from PPR are unlikely to provide good-quality meat and should be buried or destroyed by heat.
The carcass is usually dehydrated, and soiled with faeces. The peri-orbital and perinasal areas are usually encrusted with muco-purulent discharges. The erosions and ulcerations in the mouth and throat are usually prominent, as is the presence of the secondary broncho-pneumonia. The underlying primary viral pneumonia may be less obvious but is manifested by areas of level red consolidation (Rowland et al., 1969). 'Zebra striping' in the colon may also be seen, and lympadenopathy.
The most important histopathological indicator of PPRV is the presence of multi-nucleated giant cells containing intra-nuclear and intra-cytoplasmic inclusions. Multi-nucleated giant cells (syncytia) are most readily detected in the lungs, but also occur in bronchial, alveolar and ileal epithelium.
A manual on the diagnosis of PPR has been produced (FAO, 1998). Suspicion of PPR would be raised by signs of stomatitis – or pneumonitis, with enteritis in several animals. A high mortality and morbidity rate would be expected in outbreaks occurring in non-endemic areas, or where outbreaks have not occurred for some time and animals are not vaccinated. The occurrence of both stomatitis, with erosions/ulcerations of epithelial surfaces in the oral cavity, together with diarrhoea, occurs in few other single diseases of sheep and goats. In an outbreak situation, early signs of infection, such as oculo-nasal discharges and fever, would be expected for in-contact animals, if some of the group have reached the late stage of the disease or have died. In most countries PPR is a notifiable disease and the authorities require to be informed if the infection is suspected. Given the infectivity of individual animals, immediate actions or advice to community leaders to contain movement of in-contact animals are important to reduce local spread while investigations are proceeding.
Other conditions to be considered are: contagious caprine pleuropneumonia, bluetongue, pasteurellosis, contagious ecthyma, foot-and-mouth disease, heartwater, coccidiosis, mineral poisoning.
Bluetongue infection occurs in many countries that are endemic or at-risk from PPR incursion. It can give rise to a muco-purulent discharge and high morbidity and mortality rate in susceptible sheep, but usually less so in goats. It does not usually result in a severe enteritis, although loose stools may occur, or erosions/ulcerations of epithelial surfaces. Bluetongue usually gives rise to visible signs of haemorrhage on the coronary band of the foot, in contrast to PPR.
Foot-and-mouth disease affects other stock as well as sheep and goats; cattle in the region would be expected to show more severe signs than sheep or goats. On occasion though, the disease is more severe in the latter, and cattle may be absent. However, the enteritis usually present in PPR is not seen in FMD.
The lesions of orf (contagious ecthyma) and sheep and goat pox differ in distribution to that of PPR, but animals recovering from PPR may develop proliferative growths on the lips resembling orf, and the virus may be involved in the pathogenesis of the condition. Contagious caprine pleuropneumonia occurs in many similar countries to PPR but does not usually give a high mortality in sheep, or have an accompanying severe enteritis. Heartwater can give a high mortality rate in susecptible breeds, but without a stomatitis.
Detection of the virus
PPRV is present at a high concentration in secretions and tissue samples in the early stages of the disease, but rapidly becomes difficult to detect after development of antibody responses. Collection of specimens from animals which have a serous ocular-nasal discharge and fever is preferable compared to later-stage signs of necrotic stomatitis-enteritis. The Manual of Standards of the Office International des Epizooties (OIE, 2008) gives a description of the recommended samples to be collected. In live animals, swabs should be made of the conjunctival discharges from the nasal and buccal mucosae. Whole blood should be collected in anticoagulant for virus isolation, polymerase chain reaction (PCR) and haematology. From the necropsy examination of two to three animals, lymph nodes, especially the mesenteric and bronchial nodes, lungs, spleen and intestinal mucosae should also be collected aseptically, chilled on ice and transported under refrigeration. Fragments of organs collected for histopathology are placed in 10% formalin. At the end of the outbreak, blood can be collected for serological diagnosis. The priority is therefore to collect suitable specimens from early cases, after discussion with the national veterinary laboratory and Government officers of the country concerned as to their capability and preferred system for laboratory confirmation. Post-mortem examination of carcasses can be valuable, particularly in wildlife and in situations where samples cannot be kept in suitable conditions during transport to the laboratory, with a view to collection of specimens for detection of multi-nucleated giant cells by histopathology. Infection can be confirmed by identification of the agent with specific tests. Detection of a rise in titre of antibody could also be used, with paired samples collected 14-21 days apart.
Several methods for detection of virus are recognized by the Office International des Epizooties (OIE, 2008). Of these, agar gel immuno-diffusion (AGID) and counter-immuno-electrophoresis (CIEP) are relatively simple and suitable for small-scale field centres and simple laboratories. The latter gives a faster result than the former, enabling detection in less than 2 hours. Antigen-capture ELISA using monoclonal antibodies is sensitive and specific (Libeau et al., 1994). The availability of the ELISA as a kit has undoubtedly assisted countries in the detection of PPR epidemics, and provides a standardised test. Molecular tests using gene probes have been superseded by PCR-based tests (Forsyth and Barrett, 1995), and these in turn have been superseded by real-time or quantitative PCR-based tests (Bao et al., 2008; Kwiatek et al., 2010; Batten et al., 2011). PCR is extremely sensitive and specific, and may be of advantage in testing tissues where virus cannot be detected by other means, including specimens for histopathology. However, the time taken to extract RNA, and undertake the RT-PCR is usually longer than that needed for CIEP, and higher technical standards are required to avoid false-positive reactions. Other tests have been described, including simple staining methods for detecting syncitia in conjunctival cells collected onto glass slides which are suitable for simple field use (Sumption et al., 1998), and monoclonal-based staining of such cells, or infected cells in histopathological sections (Saliki et al., 1994).
The OIE Manual (OIE, 2008) recommends efforts are made to isolate the virus from outbreaks of disease, and this is particularly relevant, although more difficult, in countries where it is suspected for the first time. The FAO (Food and Agriculture Organization of the United Nations) Reference Laboratories for PPR are given on the OIE website, and are currently at CIRAD-EMVT, Montpellier, France or the Institute for Animal Health, Pirbright, Surrey, UK; either laboratory can advise on the use of various techniques for virus identification in field samples.
Antibodies are strongly induced by infection, and become detectable from the diarrheic stage onwards. The prescribed test for international trade (that which is accepted as a basis for the veterinary certification of animals as having evidence of presence or absence of antibodies) is virus neutralisation (VNT; OIE, 2008). Since there is cross-neutralisation between antibodies to PPR and RPV, a positive VNT result to PPR virus used to need to be compared to the titre obtained with RPV. The OIE considered that a serum is considered to be positive for PPR when the neutralisation titre is at least two-fold higher for PPR than for rinderpest. Since the eradication of RPV, this comparison is not required and indeed continued use of RPV in laboratories is strongly discouraged. Virus neutralisation tests involve use of live virus and cell cultures, and therefore require well-equipped laboratories and biosecurity to prevent escape of virus. VNT is therefore mainly restricted to laboratories with sufficient expertise and through-put of samples to justify the investment involved. The development of competition ELISA (C-ELISA) tests using monoclonal antibodies to either H (Anderson and McKay, 1994) or N (Libeau et al., 1995) antigens has extended the access to serological tests, and proved valuable in investigation of PPR epidemiology in the field. The tests are sensitive and specific. Kits are available from international organizations (IAEA, Vienna) and from the World Reference Laboratories for PPR in France, and the Institute for Animal Health in Pirbright, UK. Haemagglutination inhibition tests for antibody have also been described with good correlation with VNT (Raj et al., 2000).
The pathogenesis has been little studied but what we do know shows it to be essentially similar to rinderpest virus, with infection occurring via aerosols, or ingestion by nuzzling or licking, with entry through the oropharynx and subsequent multiplication in the draining lymph nodes, and thereafter in lymph nodes through the body. Virus is then released which enters the circulation and is transported to the epithelium, where it multiplies in susceptible cells, resulting in the development of lesions and disease signs associated with damage to these sites (Rossiter and Taylor, 1993).
The course of infection is swift, with an incubation period of 2 to 6 days, with death usually occurring within 14 days of infection, 5 to 7 days after onset of pyrexia. The course of infection is development of fever, followed by ocular and nasal discharges and later erosions of the epithelial surfaces of the mouth and/or gums and onset of diarrhoea. Death occurs by dehydration, complicated by pneumonia.
The first signs are of fever, with a serous nasal discharge, followed within hours by depression. Pale areas of necrosis become visible on the gums, which develop into erosions develop in the mucous membranes lining the upper alimentary, upper respiratory and uro-genital tracts 1 to 2 days after onset of fever. Salivation becomes profuse, and the nasal discharge becomes muco-purulent and may block the nostrils, and muco-purulent ocular secretions may mat the eyelids together. Ulceration of the lesions in the alimentary tract contributes to a debilitating diarrhoea, and rapid loss of condition. Pneumonia commonly occurs, with frequent secondary infections with Pasteurella, and other latent infections including Mycoplasma capri, adenoviruses, orf virus and dermatophilosis. The condition is rapidly debilitating and death may rapidly occur after development of the diarrhoea/pneumonia. Affected animals have an exceptionally miserable appearance, and recovery is slow, often accompanied by growths on the lips, attributed to recrudescence of orf virus and/or exacerbation of Dermatophilus congolense infection (Scott, 1990).
|Drug||Dosage, administration and withdrawal times||Life stages||Adverse affects||Drug resistance||Type|
|attenuated strain of PPR virus||According to manufacturer's instructions. Always seek veterinary advice before administering treatment.||All Stages||No||Vaccine|
|norfloxacin||For treatment of secondary bacterial pneumonia - intramuscular at 2.5 mg/kg body weight together with oral and intravenous administration of electrolytes. Always seek veterinary advice before administering treatment.||All Stages||No||Drug|
There is no treatment for PPRV infection itself, but antibiotics may be given to prevent secondary infections and other treatments given to alleviate the clinical signs.
Chloramphenicol 10 ml/kg body weight, penicillin 10,000 IU/kg, streptomycin 10 mg/kg, each given intramuscularly for 5 days), intestinal sedatives (Entero Sediv, 20 ml/kg for 4 days orally) and fluid therapy (Pedialyte, 30 ml/kg, for 4 days subcutaneously) were used to treat pneumonia, diarrhoea and restore body fluid ionic balance (Wosu, 1989).
|Vaccine||Dosage, Administration and Withdrawal Times||Life Stages||Adverse Affects|
|attenuated strain of PPR virus||According to manufacturer's instructions. Always seek veterinary advice before administering treatment.|
Immunization and Vaccines
A live, attenuated strain (PPRV Nigeria 75/1) of the PPR virus has been developed for use as a vaccine and provides protection for over 3 years (Diallo et al., 1989b). Other attenuated strains of PPRV have been developed in India (for review, see Sen et al., 2010). These vaccines have superseded the use of an attenuated rinderpest vaccine, whose use is no longer permitted following the eradication of RPV. Previous trials (usually with RP vaccine) reported reduction in mortality, particularly in weaned young stock, and positive benefit-to-cost ratios. A reduction in mortality of 24% was reported in Nigeria compared to controls, for a combined vaccination and dipping programme (Reynolds and Francis, 1988). In Cameroon, a benefit to cost ratio of between 2.26 and 4.23 was reported for goats and sheep through use of PPR vaccination and strategic anthelminthic treatment (Awa et al., 2000). In Niger, Stem (1993) estimated an internal rate of return of 900% for a PPRV vaccination programme over 5 years.
Husbandry methods and good practice
Risk factors for occurrence of disease are the purchase of animals from markets, and free-range husbandry of animals, when PPRV is known to be present in the region. In Oman, animals under controlled grazing (e.g. fenced or paddocked) were at lower risk than free-ranging urban livestock (Taylor et al., 1990).
Movement control, at the level of total standstill of livestock movements and banning of markets may be effective if enforceable and short-lasting in duration, since the incubation period is short. However, an effective quarantine of affected and in-contact animals for one month after the recovery of the last clinically affected case has been recommended (Rossiter and Taylor, 1993). These measures may be accompanied by a slaughter policy of animals on infected and in-contact premises, in addition to the ban on livestock movements, if the aim is rapid eradication. Measures that negatively affect livelihoods will be unpopular and difficult to enforce unless accompanied by incentives, and have rarely been implemented by authorities.
National and International Control Policy
PPR is a list A disease of the OIE, and thus member states are required to inform the OIE of the occurrence of the disease in their territory. The OIE publishes recommendations for zoo-sanitary conditions and certification of trade in animals and livestock products from countries which are not recognized as having freedom from PPR disease (OIE, 2011). The OIE recommends sanitary prophylaxis (movement control, quarantine of infected premises, with slaughter of infected animals and in-contacts) when the disease appears in previously PPR-free countries. The use of a stamping-out policy, involving slaughter of infecteds and in-contact animals on infected premises, can lead to a reduced period of time elapsing after the last case of disease has been reported before the country is internationally recognized as free of PPR.
The Animal Health Code of the OIE (OIE, 2011) considers:
- The period for quarantine purposes, ('incubation period') for the peste des petits ruminants (PPR) to be 21 days.
- A country may be considered free from PPR when it has been shown that PPR has not been present for at least the past 3 years.
- Or, this period shall be 6 months after the slaughter of the last affected animal for countries in which a stamping-out policy is practised with or without vaccination against PPR.
The Code also recognizes the presence of zones of infection, which shall be considered as infected with PPR until:
- at least 21 days have elapsed after the confirmation of the last case and the completion of a stamping-out policy and disinfection procedures,
- or until 6 months have elapsed after the clinical recovery or death of the last affected animal if a stamping-out policy was not practised.
The risk of entry of PPR into PPR-free countries is dealt with mainly by a strict prohibition of the import of live animals, recognizing that the majority of importations of virus have been via entry of live animals. The Animal Health Code of the OIE recognizes that Veterinary Administrations of PPR-free countries may prohibit importation or transit through their territory, from countries considered infected with PPR:
- of domestic and wild ruminants,
- semen of ruminants,
- embryos/ova of ruminants,
- fresh meat of domestic and wild ruminants,
- meat products of domestic and wild ruminants that have not been processed to ensure the destruction of the PPR virus,
- products of animal origin (from ruminants) intended for use in animal feeding or for agricultural or industrial use which have not been processed to ensure the destruction of the PPR virus,
- products of animal origin (from ruminants) intended for pharmaceutical or surgical use which have not been processed to ensure the destruction of the PPR virus,
- pathological material and biological products (from ruminants) which have not been processed to ensure the destruction of the PPR virus.
For each of the above, the Animal Health Code recommends the use of sanitary measures, including (where required) laboratory test results, backed by international veterinary certificates to ensure that within the boundary of error associated with test or quarantine procedures, the animals and animal products are free of disease and/or infection at the time of importation. For example, for importation of small ruminants from countries not free of PPR, Veterinary Administrations should require the presentation of an international veterinary certificate attesting that the animals:
- showed no clinical sign of PPR on the day of shipment;
- were kept since birth, or for the past 21 days, in an establishment where no case of PPR was officially reported during that period, and that the establishment was not situated in a PPR-infected zone; and/or
- were kept in a quarantine station for the 21 days prior to shipment;
- have not been vaccinated against PPR; or were vaccinated against PPR: not less than 15 days and not more than 4 months prior to shipment in the case of animals for breeding or rearing; or not less than 15 days and not more than 12 months prior to shipment in the case of animals for slaughter.
Importing countries may in addition impose additional conditions on the importation, that is, do not allow importation despite these conditions being fulfilled; they might be challenged in international courts, with the OIE acting as independent assessors of the risk involved in the importation.
There is at present no international programme for PPR control; the situation differs in each country according to the level of state subsidy, if any, for disease control programmes and the degree of emphasis on the private sector to undertake vaccination under a demand and supply system. Positive ratios for benefit-to-cost of national programmes have been reported (Stem, 1993) and increase in small-ruminant numbers in Oman was attributed to control of PPR by vaccination (Rossiter and Taylor, 1993). Additional controls on PPRV may be required if the infection adapts to fill the void left by lack of immunity to RP after eradication of rinderpest, since PPRV can infect cattle and in almost all PPR-endemic countries, RP has been present for at least the twentieth century. Morbilliviruses are highly adaptable, and the potential of PPRV to adapt to transmission via cattle requires monitoring, and continued vigilance for RP-like infections after RP eradication.
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(http://www.oie.int, accessed 5 June 2013)
Dr Geneviève Libeau
Control of Exotic and Emerging Animal Diseases
Programme Santé Animale
Campus International de Baillarguet TA A-15/G
34398 Montpellier Cedex 5
Tel: +33 (0)4 67 59 37 98 Fax: +33 (0)4 67 59 38 50
Dr Michael Baron
Institute for Animal Health
Ash Road, Pirbright
Woking, Surrey, GU24 0NF
Tel: +44-1483 23.24.41 Fax: +44-1483 23.24.48
Date of report: 03/06/2013
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