Peste des Petits Ruminants

Also known as: PPR — PPRV — Goat Plague — Pest of Sheep and Goats — Pneumoenteritis Complex — Pseudorinderpest of Small Ruminants — Stomatitis-Pneumoenteritis Syndrome

Introduction

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 south Asia through the Middle East, and from the horn of Africa through to West Africa. 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 condition is caused by a morbillivirus of the family paramoxyviridae. 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). 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 Office International des Epizooties (OIE) as a 'List A' pathogen 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 by 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.

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.

Disease occurs in goats and sheep, and has been recognized in captive wild ungulates from families of Gazellinae (Dorcas gazelle), Caprinae (Nubian ibex and Laristan sheep) and Hippotraginae (gemsbok). Experimentally, severe disease also occurs in white-tailed deer. Cattle undergo mainly subclinical reactions, and pigs develop a viraemia. 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.

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). 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. 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.

Clinical Signs

Signs include sudden onset of pyrexia (40 - 42⁰C), marked depression, lethargy, weight loss or reduced weight gain and a reduced appetite. There will be ulcers, vesicles and erosion on the tongue and oral mucosa and the animal may be smacking its lips, salivating excessively and grinding its teeth in pain. Gentle rubbing of the gum line will reveal a foul-smelling material and shreds of epithelial tissue. Similar changes may also be seen in the mucous membranes of the vagina and vulva. There will also be excessive lacrimation and nasal discharge, both of which may become purulent later on in the condition, following secondary bacterial infection.
The animal will later develop foul-smelling diarrhoea, containing blood and pieces of dead gut tissue. Dyspnoea, tachypnoea and coughing may be present. Pregnant animals may abort.

Pathology

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 lymphadenopathy.
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.

Diagnosis

History, signalment and clinical signs may lead to a presumptive diagnosis of this disease in countries where it is endemic.

Other conditions which need to be eliminated as differentials are (rinderpest in small ruminants), contagious caprine pleuropneumonia, bluetongue, pasteurellosis, contagious ecthyma, foot-and-mouth disease, heartwater, coccidiosis and mineral poisoning.

Rinderpest virus can cause disease in small ruminants, but where RP exists as a risk to small ruminants, disease in cattle would be expected, since many countries have ceased vaccination. Rinderpest has now been eradicated worldwide.

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 susceptible breeds, but without a stomatitis.

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. In live animals samples taken should include gum debris, conjunctival swabs, clotted blood and whole blood and tissues from post mortem. 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.

Detection of virus antigens by the agar gel immunodiffusion test (AGIDT) is a relatively simple, fast and cheap process. It is extremely useful as an initial test, but it does not discriminate between PPR and rinderpest viruses and further tests are needed to do this. Histopathology combined with immunohistochemical staining (e.g. immunoperoxidase) is a useful procedure because it is performed on formalin-fixed material and can discriminate between PPR and rinderpest when performed with specific monoclonal antibodies. Virus antigens can also be detected by immunocapture ELISA.
Antigen-capture ELISA is sensitive and specific, using monoclonal antibodies to give a result with PPR virus and enabling differentiation from rinderpest in small ruminants (Libeau et al., 1994). The availability of the ELISA as a kit has undoubtedly assisted countries in the detection of PPR epidemics. PCR 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. Despite these issues it is now commonly used alongside ELISA.

Antibodies are strongly induced by infection, and become detectable from the diarrhoeic 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, 2000). Since there is cross-neutralisation between antibodies to PPR and RPV, a positive VNT result to PPR virus needs to be compared to the titre obtained with RPV. The OIE considers 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. 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 tests are sensitive and specific, and enable differentiation of antibodies to rinderpest virus, should these occur as a result of use of RP vaccine in small ruminants or exposure to the type 1 lineage of RPV. Haemagglutination inhibition tests for antibody have also been described with good correlation with VNT (Raj et al., 2000). After countries have been declared free of rinderpest infection, it may be sufficient to use more simple antibody detection methods that do not require differentiation of RPV infection of small ruminants.

Treatment and Control

Chloramphenicol, penicillin, streptomycin can each be used and should be given intramuscularly for 5 days. Fluid therapy is also useful as a supportive treatment aswell as electrolyte replacement.

A live, attenuated strain (PSRV 75/1) of the PPR virus has been developed for use as a vaccine and provides protection for over 3 years. Previous trials (usually with RP vaccine) usually reported reduction in mortality, particularly in weaned young stock, and positive benefit-to-cost ratios.

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. 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.

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, 2001b). 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 infected 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.

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References

Diallo A, Barrett T, Barbron M, Subbarao SM, Taylor WP, 1989. Differentiation of rinderpest and peste des petits ruminants viruses using specific cDNA clones. Journal of Virological Methods, 23(2):127-136; 24 ref.
FAO, 1998. Recognising peste des petits ruminants. A field manual. Rome, Italy: Food and Agriculture Organisation (FAO).
Gargadennec L, Lalanne A, 1942. La peste-des-petits-ruminants. Bulletin du Service Zootechnique Epizootique du Afrique Occidentale Francaise, 5:16-21.
Gibbs EPJ, Taylor WP, Lawman MJP, Bryant J, 1979. Classification of peste des petits ruminants virus as the fourth member of the genus Morbillivirus. Intervirology, 11(5):268-274.
Libeau G, Diallo A, Colas F, Guerre L, 1994. Rapid differential diagnosis of rinderpest and peste des petits ruminants using an immunocapture ELISA. Veterinary Record, 134(12):300-304; 24 ref.
OIE, 2000. The Manual of Standards for Diagnostic Tests and Vaccines. Paris, France: Office International Des Epizooties.
OIE, 2001. World Animal Health in 2000. Parts 1 and 2. Paris, France: Office International Des Epizooties.
Raj GD, Nachimuthu K, Nainar AM, 2000. A simplified objective method for quantification of peste des petits ruminants virus or neutralizing antibody. Journal of Virological Methods, 89(1/2):89-95.
Rossiter PB, Taylor WP, 1993. Peste des Petits Ruminants. In: Infectious Diseases of Livestock, with special reference to Southern Africa. Chapter 75.
Rowland AC, Scott GR, Hill HD, 1969. The pathology of an erosive stomatitis and enteritis in West African Dwarf goats. Journal of Pathology, 98:83-87.