Peste des petit ruminants [PPR] is an acute or sub-acute febrile, highly contagious and often fatal disease of sheep, goats and wild small ruminants. The disease is characterized by fever, erosive stomatitis, conjunctivitis, gastroenteritis, pneumonia and causes serious economic losses in small ruminant’s production. Peste des Petits Ruminants is endemic in Sub-saharan Africa extending to the Arabian Peninsula, the Middle Eastern countries and India. Peste des petit ruminants virus is transmitted by close contact between infected and non-infected susceptible animals, which is likely to occur in common grazing and watering points. Infected animals shed PPRV in exhaled air, in secretions, and excretions. In the field, a presumptive diagnosis of PPR can be made on the basis of clinical, pathological, and epizootiological findings. However laboratory confirmation of PPR may be performed through virus isolation, detection of viral antigens, nucleic acid isolation and sequencing; and detection of specific antibody in the serum. The Food and Agricultural organization and the Office International des Epizooties have developed a global eradication strategy aimed at control and eradication of the disease by the year 2030

•    Peste des petits ruminants
•    Sheep
•    Epidemiology
•    Control

Gitao CG, Kihu S, Maina SM (2016) Review of Peste Des Petits Ruminants in Sheep. J Vet Med Res 3(5): 1060.

PPRV: Peste Des Petits Ruminantsvirus; USD: United States Dollars, C-ELISA:Competitive Enzyme-Linked Immuno-sorbent Assay; F: Fusion Protein, RT-PCR: Reverse Transcription Polymerase Chain Reaction; PCV: Packed Cell Volume; EDTA: Ethylene Diamine Tetracetic Acid; ? C: Degrees Centigrade

Peste des petits ruminants [PPR] severely affects small ruminants in almost 70 countries in Africa, the Middle East and parts of Asia. It is a highly contagious disease that causes USD 1.5 to 2 billion in losses each year in regions that are home to over 80% of the world’s sheep and goats and to more than 330 million of the world’s poorest people, many of whom depend on them for their livelihoods. The disease threatens food security and the livelihoods of smallholders and prevents animal husbandry sectors from achieving their economic potential.

Historical background

Peste des petit ruminants [PPR] is an acute or sub-acute febrile, highly contagious and often fatal disease of sheep, goats and wild small ruminants [1]. The disease is characterized by fever, erosive stomatitis, conjunctivitis, gastroenteritis, pneumonia and causes serious economic losses in small ruminant’s production [2,3]. The disease was first described in Cote d’ Ivoire, West Africa [4]. Peste des petit ruminants is also known as goat plague, pest of sheep and goats, Kata, stomatitis-pneumoenteritis syndrome, contagious pustular stomatitis, and pneumoenteritis complex [5]. The reference to the disease as a “plague” is indicative of the highly contagious nature and economic impacts that result from this disease. It was only in the late 1970s that PPR was determined to be a distinct virus from rinderpest virus through serology, biochemical and cross-protection experiments [6-8]. The disease was initially thought to be confined to the countries of West Africa; however PPR has now been confirmed present in several African, Middle East, Central and South Asia countries, as well as China [9,10].

Etiology

Peste des Petits Puminants [PPR], is caused by Paramyxovirus of the Morbillivirus genus. It was first described in 1942 in Côte d’Ivoire, West Africa and is closely related to rinderpest virus, canine distemper and human measles virus. The Peste des petits ruminants virus has an envelope derived from the host-cell plasma membrane, containing two transmembrane glycoproteins surrounding a nucleocapsid. The presence of the envelope renders virions sensitive to heat, lipid solvents or detergents, non-ionic detergents, formaldehyde and oxidizing agents. Peste des petits ruminants virus is also very sensitive to ultraviolet radiation and desiccation. Like all enveloped viruses PPRV is very sensitive to heat. The half-life of the virus at 37° C was estimated at 2 hours, and at 50° C infectivity was destroyed in 30 minutes [11]. Other studies have confirmed and clarified the thermal sensitivity of PPRV [12,13]. Peste des petits ruminants virus has been shown to survive in lymph nodes for 8 days at 4° C [11].

The PPR virus is also sensitive to low pH, being destroyed after death of the animal by the low pH which accompanies rigor mortis. Peste des petits ruminants virus is stable at pH between 5.8 and 9.5 but rapidly loses activity at pH below 4 or above 11 at room temperature [14]. The optimum pH for survival of PPRV is between 7 and 8 [15].

There are four lineages of PPR viruses have been identified; lineage 1 and 2 viruses in West Africa, lineage 3 in East Africa, Arabian and Southern India and lineage 4 in the Middle East and Asia subcontinent [16].

Epidemiology

Occurrence: Peste des Petits Ruminants is endemic in Subsaharan Africa extending to the Arabian Peninsula. It is also present in Middle Eastern countries and India. Historically, the disease was primarily associated with West Africa, but it extends in a belt across Africa immediately South of the Sahara, extending into Arabian Peninsula [17-19].

Transmission: Adverse climatic factors, whether seasonal or not, affect the availability of pasture and water, which leads to increased movements of small stock in search of better nutrition and shelter which then aids spread of the PPRV to susceptible groups [20]. It has been reported that in Maghreb countries of North Africa, traditional sacrifices of sheep during major Islamic festivals provide a major opportunity of seasonal clustering of small ruminants of multiple sources whose health status is often unknown, thus creating a favorable environment for the transmission and dissemination of the PPR virus [15]. The seasonal epidemiologic patterns of the PPR disease differ in different ecological systems, geographical areas and are dependent on culture and livelihood patterns of small stock owners [21]. In Pakistan, seasonal outbreaks of PPR were alluded to [22,23], suggesting that seasonal grazing patterns among nomadic livestock keepers during winter encourage diseases transmission [24]. Similar observations were made by [25] who associated PPR outbreaks in Bangladesh to winter grazing. PPR outbreaks among sheep and goats in India are described to occur any time of the year, but are most frequent during the wet [April to September / October] or cold dry [January and February] seasons [20,26].

Peste des petits ruminants virus is transmitted by close contact between infected and non-infected susceptible animals, which is likely to occur in common grazing areas. Infected animals shed PPRV in exhaled air, in secretions, and excretions approximately 10 days after the onset of fever. Sneezed or coughed out droplets by infected animals contain large amounts of virus which can spread infection. Transmission between animals in the vicinity can occur through inhalation over a distance of about 10 meters. Infected formites can act as source of infection although it is unlikely considering the rapid inactivation of the PPRV in external dry conditions. Peste des petits ruminants can be transmitted to the offspring by feeding them milk from infected dam. The virus is thought to be present in milk from 1-2 days before the signs appear and last until 45 days after complete recovery [27].

Risk factors: The spread of the PPR outbreaks has for a long time been associated with social, cultural and economic activities such as conflicts, disasters, livestock trade, cultural festivals, and change of husbandry practices, nomadism and seasonal climatic and environmental changes [28,29].

In a study carried out in Turkana Kenya, the crowding of sheep and goats at watering points during the dry season was found to be a significant risk factor for PPR outbreaks in 2009 while in 2010, sick adult goats and sheep sharing of grazing and water with lambs and kids was found to be significant source of PPR outbreaks [30].

Host range: Peste des petit ruminants is a disease of sheep and goats. In general goats are more susceptible than sheep; with sheep undergoing a milder form of the disease [27]. Other domestic animals such as camels, cattle and pigs are known to undergo subclinical infection of PPR [31]. The disease has been reported in wild small ruminants in a zoo [1] and those living in the wild [32-34]

Host determinants of PPR: Host determinant factors of PPR spread have been reported in various studies, highlighting age, sex, breed and animal species [9]. Young animals are less likely to have developed protective antibody titers and therefore are more susceptible to PPRV [35]. This high susceptibility in the young has been reported in Ethiopia, Kenya, Pakistan, India and Turkey [20,36-38]. In Oman, the disease is reported to maintain itself in susceptible yearling population, with an increase in incidence being a reflection of increased number of susceptible young goats/sheep recruited [39].

Sex has also been reported as a risk factor for susceptibility/ resistance to the disease [25,36,40,41]. Since the off-take of males, in a farm, is higher and at an early age compared to females, which end up staying in the herds for longer periods [20], females are more likely to demonstrate higher antibody titers than the males. The recruited young males, having been in the herds for a shorter period, are less likely to have been in contact with virus. Indeed, studies in Bangladesh have shown that male goats are significantly more prone to PPR than females [25]. However, studies from Pakistan have shown no significant difference between males and females, with respect to susceptibility [24].

The influences of breeds of the small ruminants on susceptibility to the disease have also been studied [24], with results showing that there are insignificant differences between goat breeds but there are significant differences between sheep breeds. Breed differences to susceptibility to PPR have been reported in other studies [27,42,43]. Goat and sheep species’ differences have been highlighted as major risk factor for PPRV susceptibility [24,36,38]. Though PPR has been described in other species of animals, the camel is emerging as a key risk factor in long distance transmission of the disease [29].

Economic significance: Peste de Petit Ruminants virus has a widespread distribution spanning Africa and Asia [44,45]. These areas encompass much of the developing world that relies heavily on subsistence farming to supply food or goods for trade, and small ruminants provide an excellent supply of both. Unfortunately, in many areas of Asia and Africa, small ruminant production and therefore the livelihoods of poor farmers is threatened by PPR among other trans-boundary animal diseases [TADs]. With its associated high morbidity and mortality, PPRV constitutes one of the major obstacles to subsistence farming [46]. Small stock and mainly sheep and goats are the main farm animals owned by the poor in most developing countries. Goats, and sheep “considered as mobile banks”, are reared as sources of not only milk and meat for family consumption, but also of income that can easily be mobilized for paying household expenditures, particularly in lean times. In addition to this economic role, sheep and goats have significant socio cultural roles. They are used as gifts or emblems for traditional rituals and religious purposes [28].

Pathogenesis: The route of infection is respiratory and is spread by airborne droplets. All secretions and excretions of infected sheep are contagious throughout the course of the disease, but no carrier state exists [47]. During acute phase of the disease, virus is shed in all secretions and excretions.

Peste des petits ruminants virus exhibits lymphoepitheliotropism [48]. Lymphotropic nature, common to all Morbilliviruses, causes a severe leukopenia in infected animals, which promotes the development of secondary infections by bacterial agents or parasitic opportunists who take advantage of the induced immunosuppression and severe clinical pictures [9]. Infection occurs primarily through naso-pharyngeal route via inhalation of virus particles [15]. Then, the virus multiplies in lymphoid organs before regional spread through blood to the epithelial cells of respiratory and gastro-intestinal tract [21].

The Peste des petits ruminants virus causes cytopathic effect that is distinguished from that of other Morbilliviruses, by it characteristic appearance of multinucleated cells capable of forming round mini syncytia and intracytoplasmic inclusion bodies and eosinophilic intra-nuclear inclusion bodies [49]. The fusion between an infected cell and neighboring cells [syncytia] aided by viral fusion protein [F] which is expressed on the surface of infected cells is one way of spreading the virus. This process of spreading the infection from cell to cell through fusion allows the virus to continue the infectious process free of neutralizing antibodies as nucleocapsids migrate from cell to cell without passing through the external environment.

Acute form

The acute form of PPR is rare in sheep. Signs generally appear 3 to 6 days after being in contact with an infected animal [50]. The disease is highly fatal in the young animals accompanied by a sudden rise in body temperature to 40-41.3° C [50]. It is accompanied by dullness sneezing serous discharge from the eyes and nostrils. A day or two day later, discrete lesions develop in the mouth and extend over the entire oral mucosa, forming diaphtheric plaques. There is a profound halitosis and the animal is unable to eat because of a sore mouth and swollen lips. Nasal and ocular discharge becomes mucopurulent and the exudate dries up, matting the eyelids and partially occluding the external narres. Diarrhea develops 3-4 days after the onset of fever. It is a profuse and faeces may be mucoid and blood tinged. Dyspnea and coughing occur later and the respiratory signs are aggravated when there is secondary bacterial pneumonia. Erosion has been described in the vulva and prepuce. Abortion has been reported during outbreak in India and Tanzania [51]. Death usually occurs within one week of the onset of the illness.

Sub-acute form

The sub-acute form is more common in sheep but they also occur in goats. This form has a longer incubation period of about 6 days. Sub-acutely affected animals are not severely affected and lack characteristic clinical signs and therefore mortality is usually very low. Lesions such as oral crusts due to mucosal discharges may appear making the disease to be confused with contagious ecthyma. Infected animals develop low grade pyrexia [39-40o C] and recover in 10-14 days and remain immune protected [52].

Clinical Pathology

Hemorrhages in the digestive system and the liver reduce number of erythrocytes and hematocrit values significantly in animals naturally infected with PPRV. The virus has affinity for lymphoid organs contributing to marked immune-suppression as indicated by leucopenia, monocytes depletion and lymphopenia (Table 1).

Competitive Enzyme-Linked Immunosorbent Assay [C-ELISA]

The C-ELISA is considered suitable for large scale testing due to its simplicity and availability of the recombinant antigen [55]. C-ELISA sensitivity is 99.4 % and specificity 94.5%. A competitive ELISA based on PPRV monoclonal antibodies specific for haemagglutinin [H] protein [56] or nucleoprotein [N] [55] was developed for detection of antibodies to PPRV in serum samples of sheep and goats.

Experimental infection

In contrast to natural PPR infections, experimental infection of susceptible animals with PPRV results in a clinical disease with high morbidity rate and low mortality rate [57,58]. Sheep experimentally infected with PPRV exhibited characteristic lesions at the lip commissure (Figure 1), severe diarrhea (Figure 2), gooseberry-like mesenteric lymph nodes (Figure 3) and intestinal congestion (Figure 4). A severe experimental form of disease has been reproduced in sheep and goats [59].

Necropsy finding

The carcass of a PPR affected animal is usually emaciated, the hindquarters soiled with soft/watery faeces and the eyeballs sunken. The eyes and nose contain dried-up discharges. Lips may be swollen and possibly scabs or nodules in late cases. The nasal cavity is congested [reddened] lining with clear or creamy yellow exudates and erosions. The pathology caused by PPR is dominated by necrotizing and ulcerative lesions in the mouth and the gastro-intestinal tract. Erosion in the oral cavity is a constant feature affecting the gums, soft and hard palates, tongue and cheeks and into the oesophagus. The abomasum is congested with multiple haemorrhages. The rumen reticulum and omasum rarely exhibit lesions. Occasionally, there may be erosions on the pillars of the rumen. The omasum is a common site of regularly outlined erosions often with oozing blood. Lesions in the small intestine are generally moderate, being limited to small streaks of hemorrhages and, occasionally, erosions in the first portions of the duodenum and the terminal ileum. The large intestine is usually more severely affected, with congestion around the ileo cecal valve, at the ceco-colic junction and in the rectum. In the posterior part of the colon and the rectum, discontinuous streaks of congestion “zebra stripes” form on the crests of the mucosal folds. In the respiratory system, small erosion and petechiae may be visible on the nasal mucosa, turbinates, larynx and trachea. Bronchopneumonia may be present, usually confined to the antero-ventral areas, and is characterized by consolidation and atelectasis. The lung is dark red or purple with areas firm to the touch, mainly in the anterior and cardiac lobes show evidence of pneumonia. Lymph nodes associated with the lungs and the intestines are soft and swollen [60].

Histopathology of the disease

Peste des petits ruminants virus causes epithelial necrosis of the mucosa of the alimentary and respiratory tracts marked by the presence of eosinophilic intracytoplasmic and intranuclear inclusion bodies [61]. Multinucleated giant cells [syncytia] can be observed in all affected epithelia as well as in the lymph nodes where there is severe depletion of lymphocytes (Figure 5), [49]. In the lungs multifocal degeneration, ulceration and necrosis, followed by alveolar type II pneumocytes hyperplasia which mostly ends up with syncytial cell formation is a prominent feature [9]. Infiltration of the lymphocytes, plasma cells and histiocytes into the alveolar septae leads to its hypertrophy and desquamation with alveolar casts [9]. Intestinal lesions are characterized by blunted villi, degeneration of surface and crypt epithelial cells; expansion of lamina propria by a primarily mononuclear infiltration with scattered syncytial cells [49].

Diagnosis of the disease

In the field, a presumptive diagnosis of PPR can be made on the basis of clinical, pathological, and epizootiological findings. However laboratory confirmation is an absolute requirement. Diagnosis of PPR may be performed through virus isolation, detection of viral antigens, nucleic acid isolation and sequencing; and detection of specific antibody in the serum [21].

Virus isolation

Detection of the virus is done by isolation of the PPR virus in cultured cells. This method of diagnosis can be very valuable as it provides live virus for biological characterization studies and the isolated viruses are stored for later studies [50]. Samples for virus isolation include heparinized blood, eye and nasal swabs [from live animals], tonsil, mesenteric lymph nodes, spleen, section of colon and lung from necropsied cases. For successful isolation, samples must be collected during the hyperthermic phase [62] and submitted to the testing laboratory in cold ice. The most widely used cell culture systems are primary lamb kidney and ovine skin [63,64] and Vero cells [65].

Table 1: Mean hematological parameters and their standard error for experimentally infected sheep before and after infection. Source: [Maina et al., 2015] [58].

Parameters	Before infection (n=5)	After infection (n=5)	Mean of the  Difference	P-value	t
	X±Se	X±Se	?±Se
WBC (X 103/ML)	19.57±7.70	61.68±25.42	42.11±28.04	0.0305	3.3581*
HGB (mg/dl)	9.46±2.15	10.34±1.57	0.88±1.12	0.1535	1.7569
RBC (X106/ML)	9.46±2.15	12.56±1.54	3.88±1.97	0.0117	4.404*
PCV (%)	21.72±7.23	26.42±3.64	4.68±4.83	0.0949	2.17
PCV (%)	16.42±1.57	21.6±2.49	5.18±2.77	0.0139	4.18
MCH (fl)	10.94±0.58	8.14±0.55	2.8±1.08	0.0044	5.7972*
MCHC (g/dl)	45.04±6.56	38.64±2.04	6.4±5.97	0.0746	2.40
Platelets (103/ml)	838±542.18	209.6±32.62	628.4±515.94	0.0528	2.7235
Neutrophils (%)	46.2±15.51	58.4±15.53	12.2±3.90	0.0022	6.9949*
Lymphocytes (%)	53.2±16.39	36.4±11.55	16.8±5.81	0.0029	6.47*
N/L	1.098±0.951	1.942±1.378	0.8436±0.461	0.0012	4.09*
Subscript *  in the same row indicate there was a significant difference (P<0.05) Key: X - Mean Se - Standard error of mean ? - Mean of the difference Abbreviations: WBC: White Blood Cells; HGB: Hemoglobin Concentration; RBC: Red Blood Cells PCV: Packed Cell Volume; MCV: Mean Corpuscular Volume; MCH: Mean Corpuscular Hemoglobin; MCHC: Mean Corpuscular Hemoglobin  Concentration These observations are predominant particularly during acute phase of disease [53]. The number of eosinophils may remain unaltered because  these cells are primarily associated with parasitic infections [54]. On the other hand; the vaccine virus induces only a transient lymphopenia without  significantly affecting the immune response to nonspecific antigen or to itself. As diarrhea develops there is a progressive hemo-concentration and  low serum sodium and potassium.

 

Nucleic acid recognition methods

Reverse transcription polymerase chain reaction [RT-PCR] techniques based on the amplification of parts of the N and F protein genes has been developed for the specific diagnosis of PPR [66,67]. This technique is 1000 times more sensitive than classical virus titration on Vero cells [66] with the advantage that results are obtained in 5 hours, including the RNA extraction, instead of 10–12 days for virus isolation.

Specimens required for diagnosis

Tears- cotton buds or swabs of absorbent cotton wool are inserted into the conjunctival sac and swirled around to collect tears. The bud of swab is broken off and placed in Phosphate buffered saline. Gum debris- this material is scraped with a spatula from the gums and placed in Phosphate buffered saline. The tissues to sample include: Lymph nodes found around the lungs [mediastinal] and alimentary tract [mesenteric], portions of the spleen and the lungs. Two sets of each tissues are required; one set of chilled but not frozen and the other is put in 10% formalin solution to preserve the samples. It was found that even after three years of storage in formalin, this sample can still be used to recover RNA for PPR confirmation [68]. Unclotted blood is needed for virus isolation and should be collected in bottles containing anticoagulant [Heparin or Ethylenediaminetetracetic acid [EDTA], serum is needed for antibody detection.

There is no specific treatment against PPR. Control of the disease in previously non-infected countries can be effected through strict quarantine, movement controls, restriction of importation of sheep and goats from affected areas, rapid identification, humane slaughter, disposal of affected animals and burning or burying carcasses and effective cleaning and disinfection of contaminated areas and clothing with lipid solvent solutions of high or low pH. Effective disinfectant agents include alcohol, ether, phenol, sodium hydroxide and common detergents. In areas where PPR is endemic, the commonly employed control mechanism is vaccination [69].

Vaccination is the most effective way to gain control epidemic PPR. In a situation where goats are reared together with sheep, the mixed herd model established that sheep were the main drivers of PPR transmission. Peste des petits ruminants disease in sheep herds was seen to persist longer than was the case in the goats and thus may serve as the reservoir for virus in between outbreaks. A simulation of the model showed that vaccination coverage of 50% of combined sheep and goats herds was enough to curtail the spread of the PPR disease within 254 days in Turkana, Kenya [70].

Although PPRV is classified into four lineages based on F and N genes, they all belong to a single serotype. Therefore vaccination using vaccine prepared from any of the lineage will provide protection against all the lineages. A live attenuated culture vaccine based on Nigeria75/1 strain is widely used for vaccination and immunization in almost all the PPRV endemic areas of the world. This vaccine is safe for pregnant dams and induces immunity in at least 98% of the vaccinated animals in the field [71]. This vaccine protects immunized small ruminants for a period of up to 3 years. The major drawback in using this vaccine is thermo stability especially since PPRV is a disease of tropical countries. Recently, a freeze dried form of this vaccine has been prepared in an excipient containing trehalose to make it thermo stable. This fortified vaccine is resistant to temperature as high as 45 degrees Celsius for 14 days with negligible loss in efficacy. The use of this vaccine to protect small ruminants will lead to effective control of PPR in developing countries [69].

It is expected that the control and eradication of PPR will improve incomes from small ruminant husbandry systems and lead to their improved profitability and productivity. In 2013, the OIE and FAO jointly decided to embark upon the control of PPR on a global scale and develop a ‘PPR Global Control and Eradication Strategy’, [?named ‘Global Strategy’] with a strong willingness to address the animal health problems in a systematic way through approaching horizontal as well as more disease-specific [vertical] issues [72]. The strategy recognizes that situations and contexts can be very different between and even within countries. Consequently, the proposal is to begin by controlling the disease in areas where it is highly endemic and then to consolidate these control efforts by concentrating on areas where a low endemic level has been reached and where eradication is a feasible objective or is already underway. For countries already free of PPR, the Global Strategy is designed to maintain this status. The duration of each stage is variable and will depend on the context [72].

PPR is primarily a problem of sheep and goats in Africa the Middle East and the Indian subcontinent. Historically, the disease was primarily reported in West Africa but has now extended in a belt across Africa and the Arabian Peninsula. It is widely distributed in sheep and goat rearing areas of all the countries. The impact of peste des petits ruminants on livestock rearing communities is very huge and pushes many of them into poverty. In a particular flock, the risk of outbreak is greatly increased when a new stock is introduced or when animals are returned unsold from livestock market. If all countries can embark on comprehensive vaccination targeting over 50 % of their small ruminant population, then the disease can be wiped out following the recommendations of OIE/FAO by the year 2030.

We are grateful to RUFORUM (RU/CFP/CGS/TADS/09/1) for supporting the research work that led to most of the interest in this subject.

1. Furley CW, Taylor WP, Obi TU. An outbreak of PPR in a zoological collection. Vet Rec. 121; 443-447.

2. Merck Sharp, Dohme Corp. The Merck Veterinary Manual. A subsidiary of Merck & Co., Inc. Whitehouse Station NJ, USA. 2016.

3. Elsawalhy A, Mariner JC, Chibeu D, Wamwayi H, Wakhusama S, OlahoMukani W, et al. Pan African strategy for the progressive control of Peste des petits ruminants [Pan African PPR strategy] Bulletin of Animal Health and Production in Africa. 2010; 58: 185-193.

4. Gargadennec L, Lalanne, A. La peste des petits ruminants. Bulletin des Services Zoo Technique et des. Epizootie de l’Afrique Occidentale Française. 1942; 5: 16-21.

5. Braide, VB : Peste des petits ruminants. World animal. Review. 1981; 39: 25-28.

6. Hamdy FM, Dardiri AH, Nduaka O, Breese SS Jr, Ihemelandu EC. Etiology of the stomatitis pneumoenteritis complex in Nigerian dwarf goats. Can J Comp Med. 1976; 40: 276-84.

7. Taylor WP. Protection of goats against Peste des Petits Ruminants with attenuated rinderpest virus. Research Res Vet Sci. 1979; 27: 321-324.

8. Taylor WP. Serological studies with the virus of peste des petits ruminants in Nigeria. Res Vet Sci. 1979; 26: 236-42.

9. Munir M, Zohari S, Berg M. Molecular Biology and Pathogenesis of Peste des Petits Ruminants Virus. 2013; 151.

10. Libeau G, Diallo A, Parida S. Evolutionary genetics underlying the spread of peste des petits ruminants virus Animal Frontiers. 2014 :14- 20.

11. Lefevre PC. Peste des Petits Ruminants et infection bovipestique des ovins et des caprins, Etudes et synthèses de l’IEMVT. 1987.

12. Rossiter PB, Taylor WP. Peste des petits ruminants. In infectious diseases of livestock. Eds. Coezter JAW. 1994; 2: 758.

13. Diallo A. Peste des petits ruminants. In: OIE Manual of Standards for Diagnostic Tests and Vaccines. Office international des Epizooties. Paris. 2000; 114-122.

14. Diallo A. Morbillivirus group: genome organisation and proteins. Vet Microbiol. 1990; 23: 155-163.

15. Dufour L. The plague of small ruminants: Moroccan outbreak of 2008, a danger to Europe? PhD thesis. The Faculty of Medicine, Creteil, National Veterinary School of Alfort. 2010.

16. Dhar BP, Sreenivasa BP, Barrett T, Corteyn M, Sing RP, Bandyopadhyay SK. Recent epidemiology of Peste de petits ruminants virus. Vet Microbiol. 2002; 88: 153-159.

17. Kwiatek O, Minet C, Grillet C, Hurard C, Carlsson E, Karimov B, et al. Peste des petits ruminants [PPR] outbreak in Tajikistan. J Comp Pathol. 2007; 136: 111-119.

18. Wang Z, Bao J, Wu X, Liu Y, Li L, Liu C, et al. Peste des petits ruminants virus in Tibet, China. Emerg Infect Dis. 2009; 15: 299-301.

19. Anees M, Shabbir MZ, Muhammad K, Nazir J, Shabbir MA, Wensman JJ, et al. Genetic analysis of Peste des petits ruminants virus from Pakistan. BMC Vet Res. 2013; 9: 60.

20. Singh RP, Saravanan P, Sreenivasa BP, Singh RK, Bandyopadhyay SK. Prevalence and distribution of peste des petits ruminants virus infection in small ruminants in India. Rev Sci Tech. 2004; 23: 807-819.

21. Gopilo A. Epidemiology of Peste des Petits Ruminants virus in Ethiopia and molecular studies on virulence. PhD thesis. Institut National Polytechnique de Toulouse. 2005.

22. Abubakar M, Khan HA, Arshed MJ, Hussain M, Ali Q. Peste des petits ruminants [PPR]: Disease appraisal with global and Pakistan perspective. Small Ruminant Research. 2011; 96: 1-10.

23. Abubakar M, Jamal MS, Arshed MJ, Hussain M, Ali Q. Peste des petits ruminants virus [PPRV] infection: Its association with species, seasonal variations and geography. Trop Anim Health Prod. 2009; 41: 1197-1202.

24. Munir M, Siddique M, Shehzad A, Zohari S, Stahl K. Sero-prevalence of antibodies to Peste des Petits ruminants at various governmental livestock farms of Punjab, Pakistan. Asian Journal of Epidemiology. 2008; 1: 82-90.

25. Sarker S, Islam H. Prevalence and Risk Factor Assessment of Peste des petits ruminants in Goats in Rajshahi, Bangladesh. Veterinary World. 2011; 4: 546-549.

26. Balamurugan V, Saravanan P, Sen A, Rajak KK, Venkatesan G, Krishnamoorthy P, et al. Prevalence of Peste des petits ruminants among sheep and goats in India. J Vet Sci. 2012; 13: 279-285.

27. Lefèvre PC, Diallo A. Peste des petites ruminants. Rev Sci Tech. 1990; 9: 935-981.

28. FAO Transboundary Animal Disease. Bulletin 33. 2009.

29. Libeau G, Kwiatek O, Lancelot R, Albina E. Peste des petits ruminants, growing incidence worldwide. The OIE and its partners. Animal Health Information Department-OIE. 2011; 52-54.

30. Kihu SM, Gitao CG, Bebora LC, Njenga MJ, Wairire GG, Maingi N, et al. Participatory risk assessment of Peste des petit ruminants: Factor analysis of small ruminants pastoral management practices in Turkana district, Kenya. Research opinions in animal & veterinary sciences. 2012; 2; 503-510.

31. Taylor WP. The distribution and epidemiology of PPR. Preventive Veterinary Medicine. 1984; 2: 157-166.

32. Ogunsanmi AO, Awe EO, Obi TU, Taiwo VO. Peste des petits ruminants [PPR] virus antibodies in african grey duiker [sylvicapra grimmia]. African Journal of Biomedical Research. 2003; 6; 59-61.

33. Sharawi SS, Yousef MR, Al-Hofufy AN, Al-Blowi MH. Isolation, serological and real time PCR diagnosis of Peste Des Petites Ruminants virus in naturally exposed Arabian Gazelle in Saudi Arabia. Veterinary World. 2010; 3: 489-494.

34. Kinne J., Kreutzer R., Kreutzer M., Wernery U. and Wohlsein P. Peste des petits ruminants in Arabian wildlife. Epidemiol Infect. 2010; 138: 1211-1214.

35. Luka PD, Erume J, Mwiine FN, Ayebazibwe C. Seroprevalence of Peste des petits ruminants. Antibodies in Sheep and Goats after Vaccination in Karamoja, Uganda: Implication on Control. International Journal of Animal and Veterinary Advances. 2011; 3: 18-22.

36. Waret-Szkuta A, Roger F, Chavernac D, YigezuL, Libeau G, Pfeiffer DU, et al. Peste des Petits Ruminants [PPR] in Ethiopia: Analysis of a national serological survey. BMC Vet Res. 2008; 4: 34.

37. Abubakar M, Jamal MS, Arshed MJ, Hussain M, Ali Q. Peste des petits ruminantsvirus[PPRV] infection; Its association with species, seasonal variations and geography. Trop Anim Health Prod. 2009; 41: 1197- 1202.

38. Waret-Szkuta A, Roger F, Chavernac D, Yigezu L, Libeau G, Pfeiffer DU, et al. Peste des Petits Ruminants [PPR] in Ethiopia: Analysis of a national serological survey. BMC Vet Res. 2008; 4: 34.

39. Taylor WP, Busaidy SA, Barret T. The epidemiology of Peste des petits ruminants in the sultanate of Oman. Vet Microbiol. 1990; 22: 341-352.

40. Abdalla AS, Majok AA, Malik KHE, Ali AS, et al. Sero-prevalence of Peste des petits ruminants virus [PPRV] in small ruminants in Blue Nile, Gadaref and North Kordofan States of Sudan. J Public Health Epidemiol. 2012; 4: 59-64.

41. Swai ES, Kapaga A, Kivaria F, Tinuga D, Joshua G, Sanka P. Prevalence and distribution of Peste des petits ruminants virus antibodies in various districts of Tanzania. Vet Res Commun. 2009; 33: 927-936.

42. Elhag AB, Taylor WP. Isolation of Peste des petits ruminants virus from the Sudan. Res Vet Sci. 1984; 36: 1-4.

43. Diop M, Sarr J, Libeau G. Evaluation of novel diagnostic tools for Peste des petits ruminants virus in naturally infected goat herds. Epidemiol infect. 2005; 133: 711-717.

44. Nanda YP, Chatterjee A, Purohit AK, Diallo A, Innui, K, Sharma RN, et al. The isolation of Peste des petits ruminants virus from northern India. Vet Microbiol. 1996; 51, 207-216.

45. Shaila MS, Shamaki D, Forsyth MA, Diallo A, Goatley L, Kitching RP, et al. Geographic distribution and epidemiology of peste des petits ruminants virus. Virus Res. 1996; 43: 149-153.

46. Banyard AC, Parida S, Batten C, Oura C, Kwiatek O, Libeau G. Global distribution of Peste des petits ruminants virus and prospects for improved diagnosis and control. J Gen Virol. 2010; 91: 2885-2897.

47. Pugh, D. G. [2002]. Sheep and Goat Medicine. Philadelphia: Saunders.

48. Scott GR. Rinderpest and peste des petits ruminants. In Gibbs EPJ [Ed.]. Virus Diseases of Food Animals. Disease Monographs. 1981; 2: 401-425.

49. Truong T, Boshra H, Embury-Hyatt C, Nfon C, Gerdts V, Tikoo S. Babiuk et al. Peste des Petits Ruminants Virus Tissue Tropism and Pathogenesis in Sheep and Goats following Experimental Infection. PLoS One. 2014; 9: 87145.

50. Kahn CM. The Merck Veterinary Manual. 9th ed. White House Station, New Jersey: Merck and Co. 2005; 617.

51. Muse EA, Karimuribo ED, Gitao GC, Misinzo G, Mellau LS, Msoffe PL, et al. Epidemiological investigation into the introduction and factors for spread of Peste des Petits Ruminants, southern Tanzania. Onderstepoort J Vet Res. 2012; 79: 457.

52. iallo A. Control of Peste des petits ruminants and poverty alleviation. J Vet Med. 2006; 53: 11-13.

53. Rajak KK, Sreenivasa BP, Hosamani M, Singh BP, Singh RK. Experimental studies on immunosuppresuve effects of Peste des petits ruminants [PPR] virus in goats. Comp Immuno Microbiol of Infect Disea. 2005; 28: 287-296.

54. Sahinduran, S, Albay MK, Sezer K, Ozmen O, Mamak, N, Haligur M, Karakurum, et al. Coagulation profile, hematological and biochemical changes in kids naturally infected with Peste des petits ruminants. Trop Anim Health Prod. 2012; 44: 453-457.

55. Libeau G, Prehaud C, Lancelot R, Colas F, Guerre L, Bishop DHL. Development of a competitive ELISA for detecting antibodies to the peste des petits ruminants virus using a recombinant nucleoprotein. Res Vet Sci. 1995; 58, 50-55.

56. Anderson J, Mckay JA, Butcher RN. The use of monoclonal antibodies in competitive Elisa for detection of antibodies to rinderpest and peste des petite ruminants viruses. In: Seromonitoring of rinderpest throughout Africa, Phase one .Proceedings of final Research Coordinating Meeting of the FAO/IAEA/ SIDA/OAU/PARC coordinated Research Programme, Bingerville, Cote d’Ivoir. 1991; 43-53.

57. Nussieba A, Osman AS Ali, ME Rahman MA. Fadol. Pathological, Serological and Virological findings in goats experimentally infected with Sudanese Peste des Petits Rumminants virus isolates. J Gen Mol Virol. 2009; 1: 001-006.

58. Maina SM, Gitao CG, Gathumbi PK. Clinico-pathological observations in Sheep and Goats exposed to Lineage III Peste des petits ruminants virus infection in Kenya. J Exp Biol & Agric Sci. 2015; 3: 72-80.

59. Bundza A, Afshar A, Dukes TW, Davis JM, Dullac GC, Susi AW. Experimental PPR [goat plague] in goats and sheep. Can J Vet Res. 1988; 52: 46-52.

60. Roeder PL, Abraham G, Kenfe G, Barret T. PPR in Ethiopian goats. Trop anim Health Prod. 1994; 26: 69-73.

61. Brown CC, Mariner JC, Olander HJ. An immunohistochemical study of the pneumonia caused by PPR virus. Vet Pathol. 1991; 28: 166-170.

62. Lefèvre PC. Peste des petits ruminants et infection bovipestique des ovins et caprins [Synthèse bibliographique], Institute d’Elevage et de Médecine vétérinaire des pays tropicaux, 94704 Maison-Alfort, France. 1987.

63. Taylor WP, A Abegunde. The isolation of Peste des petits ruminants virus from Nigerian sheep and goats. Res Vet Sci. 1979; 26 : 94-96.

64. Adombi CM, Lelenta M, Lamien CE, Shamaki D, Koffi YM, Traoré A, et al. Monkey CV1 cell line expressing the sheep-goat SLAM protein: a highly sensitive cell line for the isolation of Peste des petits ruminants virus from pathological specimens. J Virol Methods. 2011; 173: 306- 13.

65. Hamdy FM, Dardiri AH, Nduaka D, Breese SS, Ihemelandu EC. Etiology of stomatitis pneumocomplex in Nigeria dwarf goats. Can J Comp Med. 1976; 40: 276–284.

66. Couacy-Hymann E, Roger F, Hurard C, Guillou JP, Libeau G, Diallo A. Rapid and sensitive detection of peste des petits ruminants virus by a polymerase chain reaction assay. J Virol Methods. 2002; 100: 17-25.

67. Forsyth MA, Barrett T. Evaluation of polymerase chain reaction for the detection and characterisation of rinderpest and Peste des petits ruminants viruses for epidemiological studies. Virus Res. 1995; 39: 151-163.

68. OIE Workshop on ppr prevention and control, Dar es Salaam (Tanzania). 2016.

69. Kihu SM, Gitao CG, Bebora LC, Njenga JM, Wairire GG, Maingi N. Detection of Peste des petits ruminants virus in formalin fixed tissues. Trop Anim Health Prod. 2015; 47: 247-249.

70. Diallo A, Libeau G, Couacy-Hymann E, Barbron M. Recent development in diagnosis of RP and PPR. Vet Microbiol. 1995; 44: 307-317.

71. Simon Kihu, George Chege Gitao, Lily Caroline Bebora. Peste des petits ruminants disease in Turkana, Kenya. LAP Lambert Academic Publishing a trademark of Omni scriptum GmbH and Co KG. 2015. 72.PPR Eradication strategy. 2015; 88.