Thursday, October 9, 2014

EQUINE PIROPLASMOSIS
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Equine piroplasmosis (EP) is a tick-borne disease of horses caused by the intraerytrocytic protozoan parasites Babesia caballi and Theileria equi of the Order Piroplasmida. Theileria equi was previously designated as Babesia equi but compelling evolutionary, morphologic, biochemical, and genetic evidence supports its reclassification as a Theileria.
Resistance to physical and chemical action
This agent does not survive outside its hosts and can only be transmitted through a tick vector, therefore, parameters associated with resistance to physical and chemical actions (i.e. temperature, chemical/disinfectants, and environmental survival) are not meaningful. Efficacy of medicines and biologics are described under “Prevention and control.”
EPIDEMIOLOGY
This disease is a tick-transmitted disease of equids and its presence requires competent arthropod vectors. Infected animals may remain carriers of these blood parasites for long periods and act as sources of infection for other ticks. The introduction of carrier animals into areas where competent tick vectors are prevalent can lead to an epizootic spread of the disease.
Hosts
• Horses, mules, donkeys and zebra
Life Cycle and Transmission
• Babesia sporozoites invade red blood cells (RBCs) and transform into trophozoites which grow and divide into two round, oval or pear-shaped merozoites which, in turn, are capable of infecting new RBCs and the division process is then repeated • Theileria equi sporozoites inoculated into horses via a tick bite invade the lymphocytes and these intralymphocytic forms undergo development and eventually form Theileria-like schizonts; merozoites released from these schizonts invade red blood cells (RBCs) and transform into trophozoites which grow and divide into pear-shaped tetrad (‘Maltese cross’) merozoites  • Twelve species of Ixodid ticks in the genera Dermacentor, Rhipicephalus and Hyalomma have been identified as transstadial vectors of B. caballi and T. equi, while eight of these species were also able to transmit B. caballi infections transovarially o Babesia spp. can be found in various organs of tick vectors and do transmit transovarially from egg to larva o Theileria equi develop in salivary glands of tick vector and not found in other tick organs; not transmitted transovarially from egg to larva • Transmission is also possible through mechanical vectors contaminated by infected blood (e.g. contaminated needles)
Sources of infection
• Blood infected with causative parasites of piroplasmosis and associated vectors (i.e. ticks and mechanical vectors) • Infected animals may remain carriers of these blood parasites for long periods and act as sources of infection for tick vectors
Occurrence
The parasites occur in southern Europe, Asia, countries of the Commonwealth of Independent States, Africa, Cuba, South and Central America, and certain parts of the southern United States of America.
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Theileria equi has also been reported from Australia (but, apparently never established itself in this region), and is now believed to have a wider general distribution than B. caballi.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.
DIAGNOSIS
Incubation period of equine piroplasmosis associated with T. equi is 12 to 19 days and approximately 10 to 30 days when caused by B. caballi.
Clinical diagnosis
The clinical signs of equine piroplasmosis are often nonspecific, and the disease can easily be confused with other similar hemolytic conditions presenting fever, anemia and jaundice. Theileria equi tends to cause more severe disease than B. caballi. Piroplasmosis can occur in peracute, acute, subacute and chronic forms. Documented case fatality rates vary from 10–50%. Most animals in endemic areas survive infection.
Peracute form
• Rare form of disease with only clinical observation being moribund or dead animals
Acute form
• Most common form of disease cases • Characterised by fever that usually exceeds 40°C • Reduced appetite and malaise • Elevated respiratory and pulse rates • Congestion of mucous membranes • Production of a dark red urine; faecal balls that are smaller and drier than normal • Affected animals may appear unthrifty; anemic and/or icteric
Subacute form
• Similar to acute form but accompanied by weight loss in affected animals and intermittent fever • Mucous membranes vary from pale pink to pink, or pale yellow to bright yellow; petechiae and/or ecchymoses may also be visible on the mucous membranes • Normal bowel movements may be slightly depressed and the animals may show signs of mild colic
Chronic form
• Chronic cases usually present nonspecific clinical signs such as mild inappetence, poor performance and a drop in body mass
Lesions
• Lesions observed are those most often associated with an intravascular hemolytic condition • Pale or icteric mucous membranes; blood may appear thin and watery • Swollen liver with an orange-brown or paler coloration • Enlarged, dark, friable spleen; palpable on rectal examination • Kidneys may appear paler or darker than normal with possible petechial hemorrhages • Subepicardial and subendocardial hemorrhages may be visible on cardiac tissue • Mild oedematous swelling of the distal part of the limbs sometimes occurs in subacute forms • Secondary infections may lead to various non-specific lesions including oedema, emphysema or pneumonic condition of lungs
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Differential diagnosis
• Surra • Equine infectious aonemia • Dourine • African horse sickness • Purpura haemorrhagica • Plant and chemical toxicities
Laboratory diagnosis
Samples
• Several thick and thin blood smears collected from superficial skin capillaries of live animals during the acute phase of the disease (appearance of fever); organ smears can be acquired at necropsy (cerebral cortex, kidney, liver, lung, bone marrow) o slides with blood or organ smears should be air-dried and then fixed in methanol • Serum samples should also be collected
Procedures
Identification of the agent
• Microscopic examination of blood  o demonstration of parasites in stained blood; using Giemsa staining method o thick blood smear technique also used in instances very low parasitaemia o as co-infections of T. equi and B. caballi occur, accurate identification of the species of parasite is sometimes desirable  • Identification of equine piroplasmosis in carrier animals by means of blood smear examination is difficult, inaccurate and not practical on large-scale; serological methods are preferred • Molecular techniques for the detection of T. equi and B. caballi have been described o based on species-specific polymerase chain reaction
Serological tests
• Indirect fluorescent antibody (IFA) test (a prescribed test for international trade) o IFA test has been successfully applied to the differential diagnosis of T. equi and B. caballi infections o recognition of a strong positive reaction is relatively simple, but any differentiation between weak positive and negative reactions requires considerable experience in interpretation o detailed description of the protocol of the IFA test is available from published sources and an example of an IFA protocol is provided in the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animal. • Enzyme-linked immunosorbent assay (a prescribed test for international trade) o indirect ELISAs using recombinant T. equi and B. caballi proteins have shown high sensitivity and specificity in detecting antibodies in infected horses o a competitive inhibition ELISA (C-ELISA) using recombinant protein and a specific monoclonal antibody (MAb) that defines merozoite surface protein epitope overcomes problems associated with antigen purity; 94% correlation was shown between the C- ELISA and the CF test in detecting antibodies to T. equi • Complement fixation (CF) test o CF test has been used by some countries to qualify horses for importation  o The test may not identify all infected animals, especially those that have been drug- treated or that produce anti-complementary reactions, or because of the inability of IgG(T) (the major immunoglobulin isotype of equids) to fix guinea-pig complement o Therefore, the IFA test and C-ELISA have replaced the CF as the prescribed tests for international trade
For more detailed information regarding laboratory diagnostic methodologies, please refer to Chapter 2.5.8 Equine piroplasmosis the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Diagnostic Techniques”.
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PREVENTION AND CONTROL
Sanitary prophylaxis
• EP is most commonly introduced into an area by means of carrier animals or infected ticks • Thus, movement of equids requires testing (by IFA or ELISA as described above) • Reducing exposure of equids to ticks o repellents, acaricides and regular inspection; animals and premises o control and eradication of the tick vector; including removal of nearby vegetation that could harbour ticks  • Any detected EP-positive animals should be quarantined from surrounding horses and vectors • Special care in possible mechanical infection of horses with contaminated blood
Medical prophylaxis
• No biological products are available currently • Antiprotozoal agents only temporarily clear T. equi from carriers
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the OIE Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
• Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc. • Coetzer J.A.W. & Tustin R.C. Eds. (2004). - Infectious Diseases of Livestock, 2nd Edition. Oxford University Press. • Homer M.J. & et al. (2000) - Clin. Microbiol. Rev., 13 (3): 451. • Kahn C.M., Ed. (2005). - Merck Veterinary Manual. Merck & Co. Inc. and Merial Ltd.  • Spickler A.R. & Roth J.A. Iowa State University, College of Veterinary Medicine - http://www.cfsph.iastate.edu/DiseaseInfo/factsheets.htm.  • World Organisation for Animal Health (2009). - Terrestrial Animal Health Code. OIE, Paris. • World Organisation for Animal Health (2008). - Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris.
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The OIE will periodically update the OIE Technical Disease Cards. Please send relevant new references and proposed modifications to the OIE Scientific and Technical Department (scientific.dept@oie.int). Last updated October 2009
Equine Infectious Anemia
Equine infectious anemia (EIA) is a noncontagious, infectious disease of horses and other Equidae. It is caused by an RNA virus classified in the Lentivirus genus, family Retroviridae. EIA can present as an acute, subacute, or chronic infection. On occasion, the virus can be a cause of significant morbidity and mortality. The most frequently encountered form of the disease is the inapparent, chronically infected carrier.
Transmission and Pathogenesis
Under natural conditions, the most important mode of transmission of EIA is by the transfer of virus- infective blood by blood-feeding insects between horses in close proximity. Virus is found free in the plasma or cell associated, principally in monocytes and macrophages of infected animals. Although infection is considered primarily blood borne, all tissues and body fluids are potentially infectious, especially during episodes of clinical disease when viral burdens are high. Transmission of EIA by biting flies is purely mechanical; the virus does not replicate in the vector. The chance of transmission of EIA is directly proportional to the volume of blood retained on the mouthparts of the insect after feeding. In that respect, horse flies, deer flies, and to a lesser extent, stable flies are likely the most efficient vectors. It is also because they are capable of triggering host defensive behavior that interrupts feeding and results in their seeking a new susceptible host to complete their blood meal.
Additionally, EIA can be readily transmitted iatrogenically through use of blood-contaminated syringes, needles, or surgical equipment, or by transfusion of infective blood or blood products. Infrequently, transplacental transmission can occur in infected mares that experience one or more clinical episodes during pregnancy. There is evidence, although circumstantial, from a significant outbreak of EIA in an equine hospital in Ireland, of probable spread of the virus by direct or indirect transfer between horses in stalls sharing the same barn.
A very close relationship exists between presence and severity of clinical signs of EIA and the amount of virus present in infected animals. Viral burdens are highest during febrile episodes of the disease. Many of the clinical signs associated with the acute form of EIA result from infection of macrophages and the release of pro-inflammatory mediators or cytokines, specifically tumor necrosis factor α, IL- 1, IL-6, and transforming growth factor ß. This response together with suppression of platelet production are believed to be the factors responsible for the thrombocytopenia that is a characteristic feature of EIA. In addition, immune responses play a major role in the pathogenesis of EIA. Platelets from infected horses have significant amounts of bound IgG or IgM, which leads to their immune-mediated destruction, contributing to both splenomegaly and hepatomegaly.
Clinical Findings
The clinical findings and course of EIA are variable, depending on the virulence of the virus strain, viral dose, and susceptibility of the horse. After an incubation period of 15–45 days or longer in naturally acquired cases of infection, classic cases of the disease progress through three clinical
phases. An initial or acute episode lasting 1–3 days is characterized by fever, depression, and thrombocytopenia. Because these signs can be mild and transitory, they are often overlooked. Typically, this initial phase is followed by a prolonged period associated with recurring episodes of fever, thrombocytopenia, anemia, petechiation on mucous membranes, dependent edema, muscle weakness, and loss of condition. The interval between episodes can range from days to weeks or months. In most cases, the episodes of clinical disease subside within a year, and infected horses become inapparent carriers and reservoirs of EIA virus. Many of these horses remain clinically normal.
Although the foregoing represents the most commonly described clinical course of the disease, some outbreaks of EIA can be associated with peracute infection in which the primary viral infection is uncontrolled; this can result in a very high fever, severely reduced platelet counts, and acute depression, leading to death. In view of the wide variation in response seen in natural cases of infection, it is not possible to confirm a diagnosis of EIA based solely on clinical grounds.
Lesions
Photographs
Equine infectious anemia, mucous membranes
Equine infectious anemia, mucous membranes
Gross lesions frequently seen in acute cases of EIA include enlargement of the spleen, liver, and abdominal lymph nodes; dependent edema; and mucosal hemorrhages. Chronic cases of infection are characterized by emaciation, pale mucous membranes, petechial hemorrhages, enlargement of the spleen and abdominal lymph nodes, and dependent edema. Histopathologically, there is a nonsuppurative hepatitis and, in some cases, a glomerulonephritis, periventricular leukoencephalitis, meningitis, or encephalitis. Proliferation of reticuloenthelial cells is evident in many organs, especially in the liver, where there is also accumulation of hemosiderin in Küpffer cells. Perivascular accumulation of lymphocytes can be found in various organs.
Diagnosis
A provisional clinical diagnosis of EIA must be confirmed by demonstration of antibodies to the virus in blood. Although the internationally accepted serologic test is the agar gel immunodiffusion or Coggins test, there is increasing acceptance of a variety of ELISA tests, either competitive or synthetic antigen–based, because they can provide rapid results. Because ELISA tests can give a higher rate of false positives, all ELISA positive results must be confirmed by the Coggins test. When used in combination, ELISA and agar gel immunodiffusion tests provide the highest level of sensitivity combined with specificity. The Western blot is a supplemental test that can be resorted to in cases of conflicting results with other diagnostic tests. A problem with available serologic tests is that they can give negative results when testing sera collected within the first 10–14 days of infection. Whereas the vast majority of horses infected with EIA virus will have seroconverted by 45 days, there have been exceptional cases in which the interval has been ≥90 days. Virus detection assays such as the reverse transcription PCR assay are not routinely used to diagnose EIA.
Notwithstanding their sensitivity, they may not detect virus in carrier horses with very low viral loads. Although the animal inoculation test is highly sensitive for detection of EIA virus, for logistical and economic reasons, it is no longer in vogue as a means of diagnosis of EIA.
Treatment and Control
No specific treatment or safe and effective vaccine is available. Because equids infected with EIA virus present the only known source of infection, antibody-positive animals should be kept at a safe distance (~200 m) from other equids. The only recognized exception to this rule is the progeny of seropositive mares, which may possess maternal antibodies to the virus after ingesting colostrum. In most cases, passive antibody against EIA virus wanes and is no longer detectable in the Coggins test by 6–8 mo of age; detectable antibody may persist up to 12 mo, however, if ELISA testing is used.
The risk associated with maintaining infected breeding stock varies. Field studies have indicated excellent success in raising test-negative foals from inapparent carriers of EIA virus. The risks of infection in utero increase dramatically if clinical signs of EIA are seen in the mare before parturition. Unfortunately, it is not possible to accurately determine the risk posed by any equid infected by EIA virus. Inapparent carrier horses maintain low-level viremias that may increase under stressful circumstances. As compared with seronegative healthy horses, inapparent carriers have increased serum globulin concentrations and lymphocyte subset changes that are consistent with immune activation or chronic inflammation. Because EIA virus persists in infected equids for life, most regulatory agencies assume all equids seropositive for EIA virus pose the same high risk.
In the USA, seropositive horses must be placed under quarantine within 24 hr after the positive test results are known. The quarantine area must provide separation of at least 200 yd from all other equids. After a confirmatory test is performed, seropositive horses must be permanently identified using the National Uniform Tag code number assigned by the USDA to the state in which the reactor was tested, followed by the letter “A.” This identification may take the form of a hot brand, chemical brand, freezemark, or lip tattoo, and it must be applied by a USDA representative. Reactor horses must be removed from the herd by euthanasia, slaughter, or quarantine at the premises of origin. They may move interstate only under official permit to a federally inspected slaughter facility or a federally approved diagnostic or research facility, or to return to the premises of origin. After a reactor is detected in a herd, testing for EIA must be performed on all horses on the premises and repeated until all remaining equids on the premises test negative. These horses must be retested at 30- to 60-day intervals until no new cases are found. Quarantine on the premises is released when tests on the entire herd have been negative for at least 60 days after the reactor equids have been removed.
All equids shipped across state lines in the USA must be tested for EIA with a negative result within 12 mo before transport. All equids sold, traded, or donated within a state must have tested negative
for EIA no more than 12 mo before change in ownership and, preferably, no more than 60–90 days. All equids entering horse auctions or sales markets are required to have a negative test before sale, or the horse must be held in quarantine within the state until the test results are known.
It is recommended that horse owners implement an EIA control plan for their premises. All horses should be tested every 12 mo as part of a routine health program. More frequent testing may be indicated in areas that perennially have a high incidence of EIA. Owners of equids entering shows or competitive events should present proof to event officials of a negative EIA test. All new equids introduced to a herd should have a negative EIA test before entry or be isolated while tests are pending. Vector control practices, including application of insecticides and repellents and environmental insect control, should be implemented. Good hygiene and disinfection principles should be maintained to prevent iatrogenic infection of horses with contaminated needles, syringes, or equipment. 
EPIZOOTIC HAEMORRHAGIC DISEASE
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Virus family Reoviridae, genus Orbivirus, 8 or more serotypes
Ibaraki virus is a member of the EHDV serogroup (serotype 2).  EHDV shows immunological cross reactivity with the bluetongue virus group
Resistance to physical and chemical action (adapted from Bluetongue virus)
Temperature:  Extremely unstable at high temperatures. Inactivated by 50°C/3 hours; 60°C/15 minutes or 121°C /15 minutes pH: Sensitive to pH <6.0 and >8.0 Chemicals/Disinfectants: Non-enveloped virus and thus relatively resistant to lipid solvents like ether and chloroform. Readily inactivated by ß-propiolactone, 2% w/v glutaraldehyde, acids, alkalis (2% w/v sodium hydroxide), 2-3% w/v sodium hypochlorite, iodophores and phenolic compounds Survival:  Very stable in blood and tissue specimens at 20°C, 4°C, and –70°C, but not at –20°C. Resistant to ultraviolet and gamma irradiation due to its double-stranded RNA genome
EPIDEMIOLOGY
• EHD can infect most wild and domestic ruminants • Historically EHD is a disease of wild ruminants, particularly white-tailed deer in North America, and rarely a clinical disease of cattle  • A notable exception is Ibaraki virus, which caused an extensive outbreak of disease in cattle in Japan in 1959, and continues to cause cattle disease in the Far East • Recently EHD has become an emerging disease in cattle, and was added to OIE list of notifiable diseases in May 2008, following outbreaks in 4 Mediterranean countries  • Morbidity and mortality may be as high as 90% in white tailed deer; however severity varies between years and geographic locations
Hosts
• White-tailed deer mainly, with mule deer and pronghorn affected to a lesser extent • Other wild ruminants, like black-tailed deer, red deer, wapiti, fallow deer, roe deer, elk, moose, and bighorn sheep may seroconvert • Until recently, only rare outbreaks were reported in cattle, although infection is common and they may serve as temporary reservoir hosts. True persistent infection of ruminants does not occur • Ibaraki disease is seen in cattle • Sheep can be infected experimentally but rarely develop clinical signs, and goats do not seem to be susceptible to infection
Transmission
• Virus is transmitted by biological vectors, usually biting midges of the genus Culicoides, after an external extrinsic period of 10–14 days • In temperate regions infection is most common in the late summer and autumn during peak vector population, while infection occurs throughout the year in tropical regions
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• As in bluetongue infection, viraemia can be prolonged beyond 50 days, despite the presence of neutralising antibody, due to an intimate association between virus and erythrocytes. Infected deer can be viremic for up to 2 months
Sources of virus
• Blood of viraemic animals • Infection in ruminants is not contagious – biological vectors (Culicoides sp.) are required • As the virus infects endothelium, all tissues of the body may be affected
Occurrence
EHDV in cattle has been isolated throughout the world in North America, Australia, Africa, Asia, and the Mediterranean. Ibaraki disease has been reported from Japan, Korea, and Taiwan.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.
DIAGNOSIS
Incubation period is 2–10 days
Clinical diagnosis
The clinical signs of EHD manifest as haemorrhagic disease in deer, but domestic ruminants may be subclinically indected.
• Acute EHD in deer: Fever, weakness, inappetance, excessive salivation, facial oedema, hyperaemia of the conjunctivae and mucous membranes of the oral cavity, coronitis stomatitis, and excessive salivation • In prolonged cases, oral ulcers on the dental pad, hard palate, and tongue may occur. Excessive bleeding occurs in fulminant disease: bloody diarrhoea, haematuria, dehydration, and death • Acute outbreaks in cattle (similar to bluetongue): fever, anorexia, reduced milk, swollen conjunctivae, redness and scaling of the nose and lips, nasal and ocular discharge, stomatitis, salivation, lameness, swelling of the tongue, oral/nasal erosions, and dyspnoea • Ibaraki disease in cattle is characterised by fever, anorexia and difficulty swallowing • Oedema, haemorrhages, erosions, and ulcerations may be seen in the mouth, on the lips, and around the coronets; the animals may be stiff and lame • Abortions and stillbirths have also been reported in some epidemics. Some affected cattle die (up to 10%)
Lesions
EHD in deer:
• Peracute form: Severe oedema of the head, neck, tongue, conjunctiva, and lungs • Acute form: widespread haemorrhages and oedema in the mucous membranes, skin and viscera, especially heart and gastrointestinal tract (GIT) • Erosions may be found in the mouth, rumen and omasum, and necrosis in the hard palate, tongue, dental pads, oesophagus, larynx, rumen and abomasum • Chronic form: growth rings on the hooves or sloughing of the hoof wall, and erosions, ulcers or scars in the rumen
Ibaraki disease in cattle:
• Degeneration of the striated muscles in the oesophagus, larynx, pharynx, tongue, and skeletal muscles with secondary aspiration pneumonia, dehydration and emaciation
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• Marked oedema and haemorrhages in the mouth, lips, abomasum, and coronets may be observed. Erosions or ulcerations may also be present
Differential diagnosis
• Deer: indistinguishable from bluetongue, foot and mouth disease • Cattle: bluetongue, bovine virus diarrhoea, foot and mouth disease, infectious bovine rhinotracheitis, vesicular stomatitis,  malignant catarrhal fever, and bovine ephemeral fever
Laboratory diagnosis
Samples
Virus isolation and detection • Whole blood in EDTA and/or heparin • Spleen  • Lungs • Lymph nodes • Liver
Serological tests • Paired serum samples (3–5 ml each)
Procedures
Identification of the agent • Virus isolation: inoculation of a variety of cell cultures, especially Aedes albopictus cells, also CPAE (cattle pulmonary artery endothelial cells) and BHK-21 (baby hamster kidney cells). Unlike BTV, embryonated chicken eggs are not sensitive for EHDV isolation • Viral detection: reverse transcriptase polymerase chain reaction (RT-PCR): positive test results may be difficult to interpret – EHDV nucleic acid may persist in blood of infected ruminants much longer than infectious virus • Other molecular tests: dot blot and in-situ hybridisation  •  Viral RNA may be found in deer tissues for up to 160 days post infection
Serological tests • Agar gel immunodiffusion test (group specific) • Competitive ELISA (group specific) • Serum neutralisation assay (serotype specific)
PREVENTION AND CONTROL
• Other than Ibaraki in cattle, treatment and control is limited for EHDV
Sanitary prophylaxis
• Control Culicoides vectors with insecticides/larvicides, insect repellents on susceptible ruminants, and management of Culicoides breeding areas
Medical prophylaxis
• No vaccine is commercially available for EHD viruses, however, a live attenuated Ibaraki disease vaccine is used in Japan
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• EHDV vaccines could be produced but would need to be multivalent as humoral immunity is serotype specific
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
• Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc. • Coetzer J.A.W. & Tustin R.C. Eds. (2004). - Infectious Diseases of Livestock, 2nd Edition. Oxford University Press. • Fauquet C., Fauquet M. & Mayo M.A. (2005). - Virus Taxonomy: VIII Report of the International Committee on Taxonomy of Viruses. Academic Press. • Kahn C.M., Ed. (2005). - Merck Veterinary Manual. Merck & Co. Inc. and Merial Ltd.  • Spickler A.R. & Roth J.A. Iowa State University, College of Veterinary Medicine - http://www.cfsph.iastate.edu/DiseaseInfo/factsheets.htm • World Organisation for Animal Health (2009). - Terrestrial Animal Health Code. OIE, Paris. • World Organisation for Animal Health (2008). - Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris.
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The OIE will periodically update the OIE Technical Disease Cards. Please send relevant new references and proposed modifications to the OIE Scientific and Technical Department (scientific.dept@oie.int). Last updated October 2009

DOURINE
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Dourine is a parasitic venereal disease of equines caused by the flagellate protozoan Trypanosoma equiperdum of the order Trypanosomatida. Variable strain pathogenicity has been described but, illustrating the challenges to isolate T. equiperdum, no parasite strain widely accepted to be T. equiperdum has been isolated in any country of the world since 1982 and most of the strains currently available in national veterinary diagnostic laboratories are related to Trypanosoma evansi. One hypothesis asserts that T. equiperdum does not exist as a separate species and the disease condition “dourine” is actually a host- specific immune response to either Trypanosoma brucei equiperdum or T. evansi infection. A more recent study of kinetoplast DNA proposes that T. equiperdum, along with Trypanosoma evansi, are subspecies of Trypanosoma brucei. Definitive categorisation of Dourine is pending.
Resistance to physical and chemical action
This agent does not survive very long outside its hosts and is not transmitted by fomites, therefore, parameters associated with resistance to physical and chemical actions (i.e. temperature, chemical/disinfectants, and environmental survival) are not meaningful.
EPIDEMIOLOGY
Dourine is the only trypanosomosis that is not transmitted by an invertebrate vector and also differs from other trypanosomes in that it is primarily a tissue parasite that rarely invades the blood. Dourine is transmitted during breeding or infected mares may occasionally pass infection to foals. Average mortality associated with acute disease approaches 50% (especially in stallions).
Hosts
 Horses, mules and donkeys  No known natural reservoir of the parasite other than infected equids   Rats, mice, rabbits and dogs can be infected experimentally; rodents are used to maintain strains of the parasite indefinitely and to prepare antigen for diagnostic tests
Transmission
 Natural transmission occurs directly from animal to animal during coitus o mainly from stallion to mare, but may also be transmitted from mare to stallion o infection is not always transmitted by an infected animal at every copulation  Transmission of T. equiperdum by tsetse fly or other vector has not been reported  Rarely, foals may be infected via the mucosa (conjunctiva), during parturition or by drinking milk from an infected dam o foals may then transmit disease when they are sexually mature
Sources of infection
 T. equiperdum may be found in vaginal secretions of infected mares, and seminal fluid, mucous exudate of the penis, and sheath of stallions
Occurrence
The disease occurs in most of Asia, northern and southern Africa, Russia, parts of the Middle East, South America, and southeastern Europe.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.

DIAGNOSIS
Incubation period is very variable and could be from one week to a few months or longer. For the purposes of the OIE Terrestrial Animal Health Code, the incubation period for dourine is 6 months
Clinical diagnosis
Severity and duration of disease vary considerably. Though the disease is often fatal, spontaneous recoveries do occur but may result in latent carriers. Diagnosis is most commonly based on clinical evidence supported by serology.
 Clinical manifestations include:  o fever o local oedema of the genitalia and mammary glands o oedematous coetaneous eruptions o knuckling of the joints, incoordination and unilateral facial and lip paralysis o ocular lesions o anaemia o progressive weight loss and emaciation o nervous form may set in after emaciation and oedema and lead to weakness, lameness mostly of the hinds legs resulting in a „staggering movement‟, and gait abnormalities  Clinical signs are marked by periodic exacerbation and relapse, ending in death, sometimes after paraplegia or, possibly, recovery; acute disease lasts only 1–2 months or, exceptionally, 1 week  A chronic, usually mild, form of the disease may persist for several years  Subclinical infections occur; donkeys and mules are more resistant than horses o may remain inapparent carriers   In fatal cases, the disease is usually slow and progressive, with increasing anaemia and emaciation, although the appetite remains good almost throughout
Lesions
 Raised, oedematous or coetaneous urticarial skin plaques (“Silver dollar plaques”), 5–8 cm in diameter and 1 cm thick, are pathognomonic, but have not been observed in recent cases o plaques usually appear over the ribs, although they may occur anywhere on the body, and usually persist for between 3 and 7 days o not a constant feature and when present not easy to identify  Oedema disappears and returns at irregular intervals causing a thickening and induration of affected tissue; gelatinous exudates are present under the skin  In mare,  o vulva, vaginal mucosa, uterus, bladder, and mammary glands may be thickened with gelatinous infiltration o vaginal mucosa may show raised and thickened semitransparent patches o folds of swollen membrane may protrude through the vulva  In the stallion,  o the scrotum, sheath, and testicular tunica are thickened and infiltrated o testes may be embedded in a tough mass of sclerotic tissue and may be unrecognisable  Depigmentation of the genital area, perineum, and udder may occur  Lymph nodes, particularly in the abdominal cavity, are hypertrophied, softened and, in some cases, haemorrhagic  Spinal cord of animals with paraplegia is often soft, pulpy and discoloured, particularly in the lumbar and sacral regions
Differential diagnosis
 Coital exanthema  Contagious equine metritis   Surra  Nagana  Anthrax  Equine viral arteritis   Equine infectious anaemia  Purpura haemorrhagica  Other conditions leading to weight loss and emaciation: malnutrition, verminosis, dental pathology, chronic infections

Laboratory diagnosis
Samples
 Trypanosomes are present, in low numbers only, in lymph and oedematous fluids of the external genitalia, in the vaginal mucus, and exudates of plaques and mammary gland exudates; may be found in the urethral or vaginal mucus collected from preputial or vaginal washings or scrapings 4–5 days after infection  Aspirate from plaque: skin of the area over the plaque should be washed, shaved and dried, and the fluid contents aspirated by syringe; blood vessels should be avoided  Several thick blood smears o Often necessary to centrifuge collected blood and examine recentrifuged plasma o thick films are made by placing a small drop (approximately 50 μl) of blood/plasma on to a clean glass slide, droplet is air-dried, heat-fixed at 80°C for 5 minutes, and stained in 10% Giemsa for 15–20 minutes o unstained blood smears should not be stored with formalin solutions as it may affect staining quality  Whole blood (in EDTA) and serum   Trypanosoma equiperdum strains are best stored in liquid nitrogen
Procedures
Identification of the agent  A definitive diagnosis depends on the recognition of the clinical signs and the demonstration of the parasite  Rarely possible due to: o although the clinical signs and gross lesions in the developed disease may be pathognomonic, they cannot always be identified with certainty, especially in the early stages or in latent cases  they can be confused with other conditions, such as coital exanthema (moreover, in some countries [e.g. in South America], T. evansi infections give rise to similar clinical signs);  o the trypanosomes are only sparsely present and are extremely difficult to find, even in oedematous areas; and  o the trypanosomes are only fleetingly present in the blood, and in small numbers that defy detection  Microscopic examination of fresh aspirate o motile trypanosomes are present for a few days only, so that lesions should be examined at intervals  As the parasite is rarely found in thick blood films, the use of concentration techniques is recommended, such as capillary tube centrifugation, and mini anion exchange centrifugation technique o dourine is the only trypanosome to affect horses in areas free from Nagana or Surra diseases, observation of agent thick blood films is sufficient for a positive diagnosis o differentiated on the basis of morphological criteria in countries with other members of the subgenus Trypanozoon, is difficult
Serological tests Humoral antibodies are present in infected animals, whether they display clinical signs or not, however, diagnosis of dourine must include history, clinical, and pathological findings as well as serology.
 Complement fixation test (the prescribed test for international trade) o used to confirm clinical evidence and to detect latent infections o uninfected equids, particularly donkeys and mules, often give inconsistent or nonspecific reactions because of the anticomplementary effects of their sera  Indirect fluorescent antibody (IFA) test o In the case of anticomplementary sera, the IFA test is of advantage o no internationally adopted protocol o cross-reactions are possible due to the presence in some countries of other trypanosomes  Enzyme-linked immunosorbent assay o methodology described in OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animal o competitive ELISA has also been described for detecting antibody against Trypanosoma equiperdum  Other serological tests

o include radioimmunoassay, counter immunoelectrophoresis and agar gel immunodiffusion (AGID) tests  AGID has been used to confirm positive tests and to test anticomplementary sera o immunoblotting method has been published for simultaneous diagnoses of equine piroplasmosis, glanders and dourine  o card agglutination test has been developed that compares favourably with the CF test
There is an OIE Reference Laboratory for Dourine (consult the OIE Web site for the most up-to-date list: http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/).
For more detailed information regarding laboratory diagnostic methodologies, please refer to Chapter 2.5.3 Dourine in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Diagnostic Techniques”.
PREVENTION AND CONTROL
Sanitary prophylaxis
 Control of the disease depends on compulsory notification and slaughter of infected animals  Movement control enforced by legislation in most countries  Good hygiene at assisted matings is also essential o fences may help control the spread, although stallions have been reported to serve mares over fences
Medical prophylaxis
 No vaccines are available for this disease  Pharmaceutical therapy is not recommended because animals may improve clinically but remain carriers of the parasite
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the OIE Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
 Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc.  Claes F., Agbo E.C., Radwanska M., te Pas M.F.W., Baltz T., De Waal D.T., Goddeeris B.M., Claassen E., Büscher P. (2003) - How does Trypanosoma equiperdum fit into the Trypanozoon group? A cluster analysis by RAPD and Multiplex-endonuclease genotyping approach. Parasitology, 126:425-431 
CONTAGIOUS CAPRINE PLEUROPNEUMONIA
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Family Mycoplasmataceae, Mycoplasma capricolum subsp. capripneumoniae (Mccp)
• Closely related to M. capricolum subsp. capricolum and more distantly related to other members of the “Mycoplasma mycoides cluster” such as M. mycoides subsp. capri or M. leachii. • Unlike contagious caprine pleuropneumoniae (CCPP), which is confined to the thoracic cavity, the disease caused by other mycoplasmas of the mycoides cluster is accompanied by prominent lesions in other organs and/or parts of the body besides the thoracic cavity • Formerly known as Mycoplasma sp. type F-38 (5) • Genetic studies have grouped Mccp isolates into major clusters (2 or 3 depending on the study) that correlate with geographic regions. (4, 7)
Resistance to physical and chemical action (based on M. mycoides mycoides SC)
Temperature: Inactivated within 60 minutes at 56°C and within 2 minutes at 60°C, but can survive more than 10 years in frozen, infected pleural fluid.  pH:  No information. Chemicals: Inactivated by formaldehyde (0.05%/30 seconds) and a mercuric chloride (0.01%/1 minute) Disinfectants: Many of the routinely used disinfectants will effectively inactivate the organism, e.g. phenol (1%/3 minutes). Survival: Very fragile and not able to exist long in the external environment. On average only survives outside the host for up to 3 days in tropical areas and up to 2 weeks in temperate zones. Cultures can be inactivated by ultraviolet radiation within a few minutes.
EPIDEMIOLOGY
• CCPP is one of the most severe diseases of goats (9) • Affects the respiratory tract, and is extremely contagious and frequently fatal • In naive flocks, the morbidity rate may reach 100% and the mortality rate can be as high as 80% • CCPP causes major economic losses in East Africa and the Middle East, where it is endemic • During the only confirmed outbreak in wild ruminants, the morbidity rate was 100% in wild goats and 83% in Nubian ibex. The mortality rates in these two species were 82% and 58%, respectively
Hosts
• Goats are the primary hosts. • Sheep may be affected In CCPP outbreaks affecting mixed goat and sheep herds. Mccp has also been isolated from healthy sheep, and their role as a possible reservoir must be considered. • Recently CCPP was confirmed in wild ruminants kept in a wildlife preserve in Qatar. The disease affected wild goats (Capra aegagrus), Nubian Ibex (Capra ibex nubiana), Laristan mouflon (Ovis orientalise laristanica) and Gerenuk (Litocranius walleri) with significant morbidity and mortality in these species. (1) • Disease indistinguishable from naturally occurring CCPP has been experimentally reproduced with Mccp by several groups of workers.
Transmission
• Contagious caprine pleuropneumoniae is contagious.  • Disease is transmitted during close contact by the inhalation of respiratory droplets.
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• Chronic carriers may exist, but this remains unproven. Some outbreaks have occurred in endemic areas when apparently healthy goats were introduced into flocks. • Outbreaks of the disease often occur after heavy rains (e.g. after the monsoons in India), after cold spells or after transportation over long distances. This may be because recovered carrier animals shed the infectious agent after the stress of sudden climatic or environmental changes.
Sources of agent
• Infectious aerosols.  • A carrier state is likely but not proven.
Occurrence
• Occurs in many countries in Africa and the Middle East. Exact distribution is not well known and could well include Asian countries. • Mycoplasma capricolum subsp. capripneumoniae (Mccp), originally known as the F38 biotype, was first isolated in Kenya and subsequently isolated in the Sudan, Tunisia, Oman, Turkey, Chad, Uganda, Ethiopia, Niger, Tanzania, Eritrea and the United Arab Emirates • First reported in mainland Europe in 2004, when outbreaks were confirmed in Thrace, Turkey, with losses of up to 25% in some herds (6) • The exact distribution of Mccp is unknown as CCPP is often confused with other respiratory infections (Pasteurellosis) and isolation of the causative organism is difficult.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.
DIAGNOSIS
The incubation period under natural conditions is commonly six to 10 days, but may be prolonged (3– 4 weeks). Some experimentally infected goats develop fever as soon as three days after inoculation and respiratory signs as early as five days, but others become ill up to 41 days after exposure.
CCPP should be suspected in the field when a highly contagious disease occurs in goats characterised by pyrexia of 41°C or greater, severe respiratory distress, high morbidity and mortality, and post-mortem lesions of fibrinous pleuropneumoniae with pronounced hepatisation and pleural adhesions.
Clinical diagnosis
Post-mortem examination reveals fibrinous pleuropneumoniae with massive lung hepatisation and pleurisy, accompanied by accumulation of straw-coloured pleural fluid.
• CCPP is strictly a respiratory disease. Peracute, acute and chronic forms occur in endemic areas.  • Peracute: affected goats may die within 1–3 days with minimal clinical signs. • Acute: initial signs are high fever (41–43°C), lethargy and anorexia, followed within 2–3 days by coughing and laboured respiration. The cough is frequent, violent and productive. In the final stages of disease, the goat may not be able to move and stands with its front legs wide apart and its neck stiff and extended. Saliva can drip continuously from the mouth, and the animal may grunt or bleat in pain. Frothy nasal discharge and stringy saliva may be seen terminally. Pregnant goats can abort. Acutely affected goats generally die within seven to 10 days. • Chronic: there is chronic cough, nasal discharge and debilitation. • Peracute, acute and chronic disease, resembling the clinical signs in goats, were reported in captive wild goats, Nubian ibex, Laristan mouflon and gerenuk.
Lesions
• Lesions of CCPP are limited to the respiratory system.  • Acute disease is characterised by unilateral pneumonia and serofibrinous pleuritis with straw- coloured fluid in the thorax. On cut surface, the lung is granular with copious straw-coloured
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exudates. Pea-sized, yellow nodules may be found in the lungs; these nodules are surrounded by areas of congestion. Varying degrees of lung consolidation or necrosis can be seen, and the regional (bronchial) lymph nodes are enlarged. Some long-term survivors have chronic pleuropneumoniae or chronic pleuritis, with encapsulation of acute lesions and numerous adhesions to the chest wall. The interlobular septa are not thickened in domesticated goats. • Wild ruminants with CCPP have similar lesions; however, thickening of the interlobular septa has been reported in some animals.
Differential diagnosis
The diagnosis of outbreaks of respiratory disease in goats, and of CCPP in particular, is complicated, especially where it is endemic. Mccp is readily contagious and fatal to susceptible goats of all ages and both sexes, rarely affects sheep, and does not affect cattle. • Peste des petits ruminants, to which sheep are also susceptible;  • Pasteurellosis, which can be differentiated on the basis of distribution of gross lung lesions;  • Contagious agalactia syndrome, also known as Mastitis, arthritis, keratitis, pneumonia and septicaemia syndrome (MAKEPS). As the latter name implies, the pneumonia is accompanied by prominent lesions in other organs, and is caused by other mycoplasmal organisms.
Laboratory diagnosis
• M. capripneumoniae and other members of the M. mycoides cluster cross-react in serological tests and share biochemical and genetic similarities. Classical tests such as biochemical tests and growth inhibition tests are time consuming and not specific. • Definitive diagnosis can be made by isolating M. capripneumoniae from lung tissue and/or pleural fluid at necropsy • This organism has a branching, filamentous morphology in exudates, impression smears or tissue sections examined under the microscope. Other caprine mycoplasmas usually appear as short filamentous organisms or coccobacilli • Biochemical, immunological and molecular tests can be used for identification of the culture  • Polymerase chain reaction (PCR) is the preferred assay to identify M. capripneumonia cultures, and to directly identify the organism in tissue samples. There are two specific PCR assays as well as a recently developed quantitative PCR assay. (2, 3, 11) • Immunohistochemistry can identify M. capripneumoniae antigens in tissue samples, but it is not routinely used in diagnostic laboratories
Samples
• At necropsy, samples from active lung lesions should be collected for culture and histopathology. These samples should be taken from the interface between consolidated and unconsolidated areas. Samples of pleural fluid, exudates from lung lesions, and regional lymph nodes should also be collected. Tissue samples for virus isolation should be collected aseptically, placed in a transport medium, kept cold, and shipped to the laboratory on ice packs. Samples should be frozen if they will not reach the laboratory within a few days; if necessary, samples can be stored at –20°C for months with little apparent loss of mycoplasmal viability. • Paired serum samples should be collected 3–8 weeks apart.
Procedures
Identification of the agent
Definitive diagnosis requires culture of the causative organism from lung tissue samples and/or pleural fluid taken at post-mortem. After cloning and purification, isolates can be identified by several biochemical, immunological and molecular tests. Isolating the causative agent is a difficult task. Recently polymerase chain reaction based tests have been described and shown to be specific and sensitive, and can be applied directly to clinical material, such as lung and pleural fluid.
• Microscopy of lung exudates, impression smears or sections: a branching filamentous organism may be observed by dark-field microscopy or by light microscopy when stained by May–Grünwald–Giesma method.
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• Polymerase chain reaction (PCR) assays for the specific identification of Mccp nucleic acids are available. Due to the difficulty in isolating Mccp, PCR is the technique of choice for the diagnosis of CCPP. However, isolation of Mccp remains the confirmatory test.  • Gel precipitin tests to detect antigen released by Mccp in tissue specimens • Isolation of mycoplasmas: necropsy samples of choice are lung lesions, particularly from the interface between consolidated and unconsolidated areas, pleural fluid, and mediastinal lymph nodes. If microbiological examination cannot be performed immediately, samples or whole lungs can be stored at –20°C for considerable periods (months) with little apparent loss of mycoplasma viability. During transport, samples should always be kept as cool as possible, as mycoplasma viability diminishes rapidly with increasing temperature. Lung samples can be dispatched to other laboratories in frozen form. Send swabs in 2–3 ml of Mycoplasma media and 1 g of tissue sample minced in 9 ml of medium. • Identification of Mccp strains by PCR (and sequencing) has now superseded all other techniques because of its rapidity and reliability. Sequencing is used to type the strain at a finer level.
Serological tests
Complement fixation test (the prescribed test in the OIE Terrestrial Manual)
Serology has not been widely applied to identify the cause of outbreaks of pleuropneumoniae in goats and Sheep, due to occurrence of false positive results and that acute cases caused by Mccp rarely show positive titres before death. Such tests are best used on a herd basis rather than for diagnosis in individual animals.
• Complement fixation test (CFT) remains the most widely used serological test for CCPP. • Latex agglutination test is being increasingly used in diagnostic laboratories and as a pen side test. It can used to test whole blood as well as serum.  • Indirect hemagglutination (IHA) is also used.  • Competitive ELISA has been developed, but is not widely available. As with the other serological tests, it does not detect all reactors, but its specificity and suitability for large-scale testing make it an appropriate test for epidemiological investigations. It should be available commercially in the near future. (10) • Seroconversion to the IHA and CFT in experimentally infected animals begins at 7–9 days after the appearance of clinical signs, peaks between days 22 and 30, and declines rapidly thereafter. Serology should be applied on a herd basis, and paired serum samples collected 3–8 weeks apart whenever possible.
For more detailed information regarding laboratory diagnostic methodologies, please refer to Chapter 2.7.6 Contagious caprine pleuropneumonia in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Diagnostic Techniques”.
PREVENTION AND CONTROL
Sanitary prophylaxis
• Contagious caprine pleuropneumoniae is most likely to enter a country in infected animals • It is uncertain whether long-term subclinical carriers exist; however, some outbreaks in endemic areas have occurred when apparently healthy goats were introduced into flocks  • Outbreaks can be eradicated with quarantines, movement controls, slaughter of infected and exposed animals, and cleaning and disinfection of the premises • Some countries have included vaccination in their eradication procedures • In endemic areas, care should be taken when introducing new animals into the flock • Flock testing, slaughter, and on-site quarantine may be helpful in controlling the spread of disease • Vaccines help prevent disease in some countries • Some antibiotics, such as tetracycline or tylosin, can be effective if given early
The outbreak of CCPP in wild goats, ibex, mouflon and gerenuk suggests that this disease could be a threat to some wildlife and/or captive wild animals. Vaccination was helpful in ending this outbreak. In endemic areas, susceptible species should be kept from contact with goats. Mycoplasma screening should also be considered before animals are released into a zoo or other site, but M. capripneumoniae infections are difficult to detect.
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Medical prophylaxis
• The current CCPP vaccine contains inactivated Mccp suspended in saponin, has a shelf life of at least 14 months, and provides protection for over 1 year (8). It is available commercially.
For more detailed information regarding vaccines, please refer to Chapter 2.7.6 Contagious caprine pleuropneumonia in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Requirements for Vaccines and Diagnostic Biologicals”.
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the OIE Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
• Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc. • World Organisation for Animal Health (2009). - Terrestrial Animal Health Code. OIE, Paris. • World Organisation for Animal Health (2008). - Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris.  • OIE Reference experts and laboratories  • OIE 2008 Online World Animal Health Information Database (WAHID) • The Merck Veterinary Manual, 9th Edition, Cynthia M. Kahn (Editor), Scott Line (Associate Editor) • Infectious Diseases of Livestock, Coetzer, JAW and Tustin RC. 2004. Oxford University Press • Emerging and Exotic Diseases of Animals, Spickler AR and Roth JA. 2006. Iowa State University
1. Arif A., Schulz J., Thiaucourt F., Taha A. & Hammer S. (2007). Contagious caprine pleuropneumonia outbreak in captive wild ungulates at Al Wabra Wildlife Preservation, State of Qatar. J. Zoo. Wildl. Med., 38, 93–96. 2. Bascunana C.R., Mattsson J.G., Bolske G. & Johansson K.E. (1994). Characterization of the 16S rRNA genes from Mycoplasma sp. strain F38 and development of an identification system based on PCR. J. Bacteriol., 176, 2577–2586. 3. Lorenzon S., Manso-Silván L. & Thiaucourt F. (2008). Specific real-time PCR assays for the detection and quantification of Mycoplasma mycoides subsp. mycoides SC and Mycoplasma capricolum subsp. capripneumoniae. Molec. Cell. Probes, 22, 324–328. 4. Lorenzon S., Wesonga H., Ygesu L., Tekleghiorgis T., Maikano Y., Angaya M., Hendrikx P. & Thiaucourt P. (2002). Evolution of M. capricolum subsp. capripneumoniae strains and molecular epidemiology of contagious caprine pleuropneumonia. Vet. Microbiol., 85, 111– 123. 5. MacOwan K.J. & Minette J.E. (1976). A mycoplasma from acute contagious caprine pleuropneumonia in Kenya. Trop. Anim. Health Prod., 8, 91–95. 6. Ozdemir U., Ozdemir E., March J.B., Churchward C. & Nicholas R.A. (2005). Contagious caprine pleuropneumonia in the Thrace region of Turkey. Vet. Rec., 156, 286–287. 7. Pettersson B., Bolske G., Thiaucourt F., Uhlen M. & Johansson K.E. (1998). Molecular evolution of Mycoplasma capricolum subsp. capripneumoniae strains, based on polymorphisms in the 16S rRNA genes. J. Bacteriol., 180, 2350–2358. 8. Rurangirwa F.R., McGuire T.C., Kibor A. & Chema S. (1987). An inactivated vaccine for contagious caprine pleuropneumonia. Vet. Rec., 121, 397–400. 9. Thiaucourt F. & Bolske G. (1996). Contagious caprine pleuropneumonia and other pulmonary mycoplasmoses of sheep and goats. Rev. Sci. Tech., 15, 1397–1414. 10. Thiaucourt F., Bolske G., Libeau G., Le Goff C. & Lefevre P.C. (1994). The use of monoclonal antibodies in the diagnosis of contagious caprine pleuropneumonia (CCPP). Vet. Microbiol., 41, 191–203.
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11. Woubit S., Lorenzon S., Peyraud A., Manso-Silvan L. & Thiaucourt F. (2004). A specific PCR for the identification of Mycoplasma capricolum subsp. capripneumoniae, the causative agent of contagious caprine pleuropneumonia (CCPP). Vet. Microbiol., 104, 125–132.
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The OIE will periodically update the OIE Technical Disease Cards. Please send relevant new references and proposed modifications to the OIE Scientific and Technical Department (scientific.dept@oie.int). Last updated October 2009. 
CONTAGIOUS BOVINE PLEUROPNEUMONIA
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Mycoplasma mycoides subsp. mycoides Small Colony - bovine biotype (MmmSC)
The M. mycoides cluster consists of six mycoplasma strains from bovines and goats that share serological and genetic characteristics, creating difficulties for taxonomy and diagnostics by traditional techniques. Specific identification of MmmSC can now be achieved by polymerase chain reaction (PCR) or the use of specific monoclonal antibodies (MAbs). Although MmmSC has been considered to be a very homogeneous biotype, recent molecular techniques have identified differences among strains. Recently described multi-locus sequence analysis distinguishes the three main lineages that correlate with their geographical origins (Europe, Southern Africa, rest of Africa). The strains of European origin can be differentiated from African ones by molecular methods, and are not able to oxidise glycerol, which may account for an apparent lower pathogenicity. African strains seem to be more diverse. The sequence of the complete genome of the reference strain PG1 has been published.
Mycoplasmas lack cell walls and are, therefore, a) pleomorphic and b) resistant to antibiotics of the beta- lactamine group, such as penicillin
Growth of mycoplasma is relatively fastidious and requires special media rich in cholesterol (addition of horse serum).
Resistance to physical and chemical action
Mycoplasma mycoides subsp. mycoides SC does not survive for long in the environment and transmission requires close contact, although, under favourable atmospheric conditions of humidity and wind, aerosols can transport the agent for longer distances.
Temperature:  Inactivated within 60 minutes at 56°C and 2 minutes at 60°C pH: Inactivated by acid and alkaline pH Chemicals/Disinfectants: Inactivated by many of the routinely used disinfectants. Inactivated by mercuric chloride (0.01%/1 minute), phenol (1%/3 minute), and formaldehyde solution (0.5%/30 seconds) Survival:  Survives outside the host for up to 3 days in tropical areas and up to 2 weeks in temperate zones. May survive more than 10 years frozen.
EPIDEMIOLOGY
Hosts
Cattle, both Bos taurus and Bos indicus, are the main hosts. Infections have also been reported from Asian buffalo (Bubalus bubalis), captive bison (Bison bison) and yak (Poephagus grunnien, formerly Bos grunnien). Sheep and goats can also be naturally infected, but with no clear associated pathology. Wild bovids and camels seem to be resistant, and, so far, do not appear to be important in the transmission of CBPP.
Incubation period of the disease is usually 1–4 months, but can be longer. After experimental inoculation into the trachea, clinical signs may appear in 2–3 weeks.
Transmission
• CBPP is spread mainly by inhalation of droplets from infected coughing animals, especially if they are in the acute phase of the disease. • Although close and repeated contact is generally thought to be necessary for transmission, transmission may occur up to 200 metres under favourable climatic conditions
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• Organism also occurs in saliva, urine, fetal membranes and uterine discharges. • Transplacental infection can occur  • Nonclinical bovine carriers with chronic infection are a major source of infection, and may retain viable organisms in encapsulated lung lesions (sequestra) for up to 2 years.  ƒ It is widely believed that recovered animals harbouring infectious organisms within pulmonary sequestra may become active shedders when stressed or immunodepressed. • Cattle movement is an important factor in the spread of the disease • Outbreaks usually begin as the result of movement and contact of an infected animal with a naive herd • There are a few anecdotal reports of transmission on fomites, but Mycoplasmas do not survive for long periods in the environment, and indirect transmission is thought to be unimportant
Sources of infection
MmmSC occur in great numbers in bronchial secretions, nasal discharges, exhaled air and nasal aerosols. Spread of infection through urine droplets was not fully confirmed. Microorganisms have also been isolated from bull semen, but transmission through semen requires further investigation.
Occurrence
CBPP is widespread in sub-Saharan Africa, including countries in the West, South, East, and Central regions of Africa.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.
DIAGNOSIS
Clinical diagnosis
In adults
• Initial signs are usually a depressed, inappetent animal with moderate fever, followed by coughing, thoracic pain and increased respiratory rate. • As pneumonia progresses, there is laboured respiration and dyspnoea, and animals prefer to stand with elbows abducted to decrease thoracic pain and increase chest capacity • Auscultation of the lungs may reveal a wide variety of sounds, depending on how severely the subjacent pulmonary parenchyma is affected. ƒ Reputations, rales, and pleuretic friction rubs are all possible. ƒ At percussion, dull sounds can be noticed in the low areas of the thorax. • CBPP often evolves into a chronic disease, characterised by ill thrift and recurrent low- grade fever that may be difficult to recognise as pneumonia • Forced exercise may precipitate coughing
In calves
• Pulmonary tropism is not the general rule, and infected calves present arthritis with swelling of the joints  o Co-existence of pulmonary signs in adults and arthritis in young animals should alert the clinician to a diagnosis of CBPP
Lesions
• Gross pathologic lesions of the lung are characteristic and often unilateral; the affected pulmonary parenchyma is odourless
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• The predominant gross change is consolidation, or thickening, of individual lobules that become encased in markedly widened interlobular septa, resulting in the characteristic marbled appearance • Interlobular septa become distended first by oedema, then by fibrin, and finally by fibrosis; the organism produces a necrotising toxin, galactan, which allows for this extensive spread through septa • Abundant yellow or turbid exudate in the pleural cavity (up to 30 litres in severe cases) that coagulates to form large fibrinous clots  • Fibrinous pleurisy: thickening and inflammation of the pleura with fibrous deposits  • Interlobular oedema, marbled appearance due to hepatisation and consolidation at different stages of evolution usually confined to one lung  • Sequestrae with fibrous capsule surrounding grey necrotic tissue (coagulative necrosis) in recovered animals • MmmSC can survive within these sequestra for months or longer, facilitating spread
Differential diagnosis
Acute form
• Acute bovine pasteurellosis • Haemorrhagic septicaemia • East Coast fever (theileriosis) • Bovine ephemeral fever • Traumatic pericarditis
Chronic form
• Ecchinococcosis (hydatid cyst)  • Actinobacillosis • Abscesses, tuberculosis, bovine farcy
Laboratory diagnosis
Samples
• Samples from live animals include nasal swabs and/or broncho-alveolar washings, or pleural fluid obtained by puncture; blood and sera should also be collected • Samples to be taken at necropsy are lung lesions, lymph nodes, pleural fluid and synovial fluid from those animals with arthritis • Samples should be shipped cool but may be frozen if transport to the laboratory is delayed
Procedures
Identification of the agent
• Isolation of pathogen from clinical samples and identification by metabolic and growth inhibition tests • The growth of MmmSC takes can take up to 10 days. In specific culture media (agar and broth), growth is visible within 3–10 days as a homogeneous cloudiness with whirls when shaken; on agar, small colonies develop, 1 mm in diameter, with the classical ‘fried-egg’ appearance. • The organism is then identified routinely with immunological tests (growth inhibition, immunofluorescence or dot immunobinding on a membrane filter [MF-dot] test) • Definitive identification is best done by an OIE Reference Laboratory (http://www.oie.int/eng/OIE/organisation/en_listeLR.htm), using biochemical tests combined with immunological assays. • Polymerase chain reaction is now used as a rapid, specific, sensitive and easy to use test
 4
Serological tests
• Modified Campbell & Turner complement fixation (CF) test is suitable for determining existence of disease and is a prescribed test in the OIE Terrestrial Manual. However, it has low sensitivity (70%), and may miss animals in early infection, those with chronic lesions, and those where therapy has been given; for herds, however, it can detect nearly 100% of infected groups. • Competitive ELISA is also an OIE prescribed test for international trade and is described in the OIE Terrestrial Manual. • An immunoblotting test (IBT) is highly specific and sensitive; it should be used at the local level in CBPP eradication programmes as a confirmatory test for positive or doubtful results after screening by the CF test and/or ELISA.
For more detailed information regarding laboratory diagnostic methodologies, please refer to Chapter 2.4.9 Contagious bovine pleuropneumonia in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Diagnostic Techniques”.
PREVENTION AND CONTROL
Effectiveness of treatment has not been adequately studied. Antibiotic treatment is not recommended because it may delay recognition of the disease, create chronic carriers and encourage emergence of resistant MmmSC strains. The methods used for control depend on the epidemiological situation, animal husbandry methods in effect, and the availability and efficacy of veterinary services in a specific country.
Sanitary prophylaxis
• In disease-free areas: quarantine, movement controls, serological screening and slaughtering of all positive and in-contact animals  • Control of cattle movements is the most efficient way of limiting the spread of CBPP
Medical prophylaxis
• In enzootic areas like Africa vaccination is very important in the control of CBPP • The only vaccines commonly used today are produced with attenuated MmmSC strains; their efficacy is directly related to the virulence of the original strain used in production • Attenuated virulent strains stimulate the best immunity, but also induce the most severe and undesirable local and systemic reactions • Two strains are used for preparing CBPP vaccines: strain T1/44, a naturally mild strain isolated in 1951 by Sheriff & Piercy in Tanzania, and strain T1sr; T1sr is completely avirulent but has shorter immunity than T1/44, which may induce an unpredictable number of animals with post-vaccinal reactions requiring treatment with antibiotics two to three weeks after vaccination • In low prevalence or free areas such as Europe, vaccination is not recommended as it can interfere with screening surveillance serological tests
For more detailed information regarding vaccines please refer to Chapter 2.4.9 Contagious bovine pleuropneumonia in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Requirements for Vaccines and Diagnostic Biologicals”.
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
• Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc. • Coetzer J.A.W. & Tustin R.C., Eds. (2004). - Infectious Diseases of Livestock, 2nd Edition. Oxford University Press.
 5
• Recommended standards for epidemiological surveillance systems for Contagious Bovine Pleuropneumonia. 1997. Rev. Sci. Tech. 16 (3): 898-918 • World Organisation for Animal Health (2009). - Terrestrial Animal Health Code. OIE, Paris. http://www.oie.int/eng/info/en_ppcb.htm; http://www.oie.int/ENG/normes/mcode/en_sommaire.htm; http://www.oie.int/ENG/normes/mcode/en_chapitre_1.11.8.htm • World Organisation for Animal Health (2008). - Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris. http://www.oie.int/Eng/Normes/Mmanual/A_summry.htm; http://www.oie.int/Eng/Normes/Mmanual/2008/pdf/2.04.09_CBPP.pdf
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The OIE will periodically update the OIE Technical Disease Cards. Please send relevant new references and proposed modifications to the OIE Scientific and Technical Department (scientific.dept@oie.int). Last updated October 2009. 
CLASSICAL SWINE FEVER (hog cholera)
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Family Flaviviridae, genus Pestivirus, one serotype divided into three major genotypes  and ten subtypes. Closely related to ruminant pestiviruses causing bovine virus diarrhoea and border disease.
Resistance to physical and chemical action
Temperature:  Readily inactivated by cooking: heating meat to 65.5°C for 30 minutes or 71°C for one minute. Survives months in refrigerated meat and years in frozen meat. Some strains are partially resistant to moderate heat (56°C). pH:  Stable at pH 5-10. Rapidly inactivated at pH <3.0 or pH >11.0. Chemicals/Disinfectants:  Susceptible to ether, chloroform, ß-propiolactone (0.4%).  Inactivated by chlorine-based disinfectants, cresol (5%), sodium hydroxide (2%), formalin (1%), sodium carbonate (4% anhydrous or 10% crystalline, with 0.1% detergent), ionic and non-ionic detergents, and strong iodophors (1%) in phosphoric acid. Survival:  Moderately fragile and does not persist in the environment. Sensitive to drying and ultraviolet light. Survives well in pens during cold conditions (up to 4 weeks in winter). Survives 3 days at 50°C and 7-15 days at 37°C. Survives in meat during salt curing and smoking for 17 to >180 days depending on the process used. Virus persists 3–4 days in decomposing organs and 15 days in decomposing blood and bone marrow.
EPIDEMIOLOGY
Virulence of disease is related to strain of virus isolate, age of pig and immune status of herd. Virus is highly contagious. Acute disease is still the prevalent form in younger animals, with subacute and chronic forms often observed in older animals.
Hosts
Pigs and wild boar are the only natural reservoir of classical swine fever virus. All feral and wild pigs, including European wild boar, are susceptible. Collared peccaries  were susceptible in one study, but recovered in 10 days.
Transmission
• Mainly by the oral and oronasal routes, via direct or indirect contact. • Direct contact between animals (secretions, excretions, semen, blood) • Spread by farm visitors, veterinarians, pig traders • Indirect contact through premises, implements, vehicles, clothes, instruments and needles • ‘Neighbourhood effect’ during outbreaks in areas of high pig farm density: airborne transmission over short distances (up to 1 km in one study) • Insufficiently cooked waste food fed to pigs: most common means of entry into free countries • Transplacental infection: may create inapparent carrier piglets or congenital abnormalities • Wild boar populations may harbour virus; domestic pigs in the affected area are at a high risk; and biosecurity is crucial
 2
Sources of virus
• Blood, secretions and excretions (oronasal and lachrymal discharges, urine, faeces and semen) and tissues of sick or dead animals, including meat • Congenitally infected piglets are persistently viraemic and may shed the virus for 6– 12 months before dying • Infection routes: ingestion (most common), contact with the conjunctiva or mucous membranes, skin abrasions, genital transmission, artificial insemination, percutaneous blood transfer
Occurrence
The disease occurs in much of Asia, Central and South America, and parts of Europe and Africa. Many countries are free of the disease.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.
DIAGNOSIS
Incubation period is 2–14 days. Clinical form varies with the strain of virus, the age/susceptibility of pigs and the occurrence of other pathogens in the herd (herd health status).
Clinical diagnosis
Acute form (more virulent virus strains and/or younger pigs)
• Fever (41°C) • Anorexia, lethargy • Severe leucopenia • Multifocal hyperaemia and/or haemorrhagic lesions of the skin • Conjunctivitis  • Enlarged, swollen lymph nodes • Cyanosis of the skin especially of extremities (ears, limbs, tail, snout)  • Transient constipation followed by diarrhoea  • Vomiting (occasional)  • Dyspnoea, coughing  • Ataxia, paresis and convulsion  • Pigs huddle together  • Death occurs 5–25 days after onset of illness  • Mortality in young pigs can approach 100%
Chronic form (less virulent virus strains or partially immune herds)
• Dullness, capricious appetite, pyrexia, diarrhoea for up to 1 month  • Ruffled appearance of pigs  • Growth retardation • Apparent recovery with eventual relapse and death within about 3 months
Congenital form (outcome depends on virulence of virus strain and stage of gestation)
• Fetal death, resorption, mummification, stillbirth  • Abortion • Congenital tremor, weakness  • Runting and poor growth over a period of weeks or months leading to death
 3
• Born clinically normal but persistently viraemic with no antibody response: important intermittent shedders of virus until dying in 6–12 months (late onset form). Mild form (usually older animals; outcome depends on virulence of virus strain):
• Transient pyrexia and inappetence  • Recovery and (lifelong) immunity
Lesions
Acute form: Lesions are usually complicated by secondary infections
• Leucopoenia and thrombocytopenia  • Enlarged haemorrhagic lymph nodes are common • Widespread petechiae and ecchymoses, especially in the skin, lymph nodes, epiglottis, bladder, kidney and rectum • Severe tonsillitis with necrotic foci sometimes occurs • Multifocal infarction of the margin of the spleen is characteristic: nearly pathognomonic but occurs infrequently with currently circulating strains • Lungs may be congested and haemorrhagic • Encephalomyelitis with perivascular cuffing is common
Chronic form: Lesions are usually complicated by secondary infections
• ‘Button’ ulcers in the caecum and large intestine mucosa • Generalised depletion of lymphoid tissue • Transverse striations of unmodelled growth cartilage at costochondral junctions in growing pigs • Haemorrhagic and inflammatory lesions are often absent
Congenital form
• Central dysmyelinogenesis, cerebellar hypoplasia, microencephaly, pulmonary hypoplasia, hydrops and other malformations.
Differential diagnosis Varies with form of the disease
• African swine fever (indistinguishable clinico-pathologically. It is essential to send samples for laboratory confirmation.) • Septicaemias: erysipelas, eperythrozoonosis, salmonellosis, streptococcosis, pasteurellosis, actinobacillosis, and Haemophilus parasuis • Haemorrhage: porcine dermatitis and nephropathy syndrome, haemolytic disease of the newborn, coumarin poisoning, thrombocytopenic purpura • Runting: post weaning multisystemic wasting syndrome, enterotoxicosis, swine dysentery, campylobacteriosis • Abortions: Aujeszky’s disease (pseudorabies virus), encephalomyocarditis virus infection, porcine reproductive and respiratory syndrome, parvovirus • Nervous signs: viral encephalomyelitis, salt poisoning • Congenital infection with ruminant pestiviruses: Bovine virus diarrhea, Border disease
Laboratory diagnosis
Procedures
Samples
Identification of the agent
Method of choice for detecting herds early in infection is to collect whole blood and tissues from multiple febrile or recently dead animals.
 4
• Tonsil  • Lymph nodes (pharyngeal, mesenteric)  • Spleen  • Kidney  • Distal ileum  • Blood in EDTA or Heparin (live cases)
Refrigerate and ship to laboratory as quickly as possible
For details, refer to the OIE Terrestrial Manual
Identification of the agent
• Reverse transcription polymerase chain reaction (RT-PCR) or real time RT-PCR.  • Virus isolation in cell culture, with virus detection by immunofluorescence or immunoperoxidase. Confirmatory identification with monoclonal antibodies. • Direct immunofluorescence test on cryostat sections of organs from affected pigs.
Serological tests
Antibodies develop only during the third week of illness: submit sera from convalescent pigs and from contact herds when >3 weeks have elapsed since suspected contact took place. Serum should also be tested from sows with suspected congenitally infected litters. Antibodies persist for life in recorded pigs. The following may be used for serological diagnosis or surveillance, and are also tests prescribed by the OIE for screening for international trade:
• Neutralisation peroxidase-linked assay  • Fluorescent antibody virus neutralisation • ELISA
For more detailed information regarding laboratory diagnostic methodologies, please refer to Chapter 2.8.3 Classical swine fever in  the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals  under the heading “Diagnostic Techniques” .
PREVENTION AND CONTROL
No treatment is possible. Affected pigs must be slaughtered and the carcases buried or incinerated.
Sanitary prophylaxis
• Effective communication between veterinary authorities, veterinary practitioners and pig farmers  • Effective disease reporting system  • Strict import policy for live pigs, pig semen, and fresh and cured pig meat  • Quarantine of pigs before admission into herd  • Efficient sterilisation (or prohibition) of waste food fed to pigs  • Efficient control of rendering plants  • Structured serological surveillance targeted to breeding sows and boars  • Effective pig identification and recording system • Effective hygiene measures protecting domestic pigs from contact with wild boar
Medical prophylaxis
Vaccination with modified live virus strains is effective in preventing losses in countries where classical swine fever is enzootic, but is unlikely, on its own, to eliminate infection entirely. In countries which are free of disease, or where eradication is in progress, vaccination is normally prohibited.
 5
For more detailed information regarding vaccines, please refer to Chapter 2.8.3 Classical swine fever in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Requirements for Vaccines and Diagnostic Biologicals”.
Response to outbreaks
• Slaughter of all pigs on affected farms  • Safe disposal of carcasses, bedding, etc.  • Thorough disinfection  • Designation of infected zone, with control of pig movements  • Detailed epidemiological investigation, with tracing of possible sources (up-stream) and possible spread (down-stream) of infection  • Surveillance of infected zone, and surrounding area
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the OIE Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
• Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc. • Coetzer J.A.W. & Tustin R.C., Eds. (2004). - Infectious Diseases of Livestock, 2nd Edition. Oxford University Press. • Fauquet C., Fauquet M., & Mayo M.A. (2005). - Virus Taxonomy: VIII Report of the International Committee on Taxonomy of Viruses. Academic Press. • Spickler A.R. & Roth J.A. Iowa State University, College of Veterinary Medicine - http://www.cfsph.iastate.edu/DiseaseInfo/factsheets.htm • World Organisation for Animal Health (2009). - Terrestrial Animal Health Code. OIE, Paris. • World Organisation for Animal Health (2008). - Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris. • Technical Part accompanying the “Diagnostic Manual” of Commission Decision 2002/106/EC • Commission Decision 2002/106/EC approving of a “Diagnostic Manual”, Anonymous (2002)
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The OIE will periodically update the OIE Technical Disease Cards. Please send relevant new references and proposed modifications to the OIE Scientific and Technical Department (scientific.dept@oie.int). Last updated October 2009. 
BOVINE BABESIOSIS
Aetiology Epidemiology Diagnosis Prevention and Control References
AETIOLOGY
Classification of the causative agent
Bovine babesiosis (BB) is a tick-borne disease of cattle caused by the protozoan parasites of the genus Babesia, order Piroplasmida, phylum Apicomplexa. The principal species of Babesia that cause BB are: Babesia bovis, Babesia bigemina and Babesia divergens. Other Babesia that can infect cattle include B. major, B. ovata, B. occultans and B. jakimovi.
Resistance to physical and chemical action
This agent does not survive outside its hosts and can only be transmitted through a tick vector. Therefore, parameters associated with resistance to physical and chemical actions (such as temperature, chemical/disinfectants, and environmental survival) are not meaningful. Susceptibility to medicines and vaccines are described under “Prevention and control”.
EPIDEMIOLOGY
All Babesia are transmitted by ticks with a limited host range. The principal vectors of B. bovis and B. bigemina are Rhipicephalus spp. ticks and these are widespread in tropical and subtropical countries. The major arthropod vector of B. divergens is Ixodes ricinus. BB is principally maintained by subclinically infected cattle that have recovered from disease. Morbidity and mortality vary greatly and are influenced by prevailing treatments employed in an area, previous exposure to a species/strain of parasite, and vaccination status. In endemic areas, cattle become infected at a young age and develop a long-term immunity. However, outbreaks can occur in these endemic areas if exposure to ticks by young animals is interrupted or immuno-naïve cattle are introduced. The introduction of Babesia infected ticks into previously tick-free areas may also lead to outbreaks of disease.
Hosts
 B. bovis and B. bigemina o cattle o water buffalo (Bubalus bubalis) and African buffalo (Syncerus caffer) o reports of disease in white-tailed deer (Odocoileus virginianus) in Mexico  B. divergens o cattle and reindeer (Rangifer tarandus) o Mongolian gerbils (Meriones unguiculatus); other peridomestic rodents are resistant to disease o Splenectomised humans and non-human primates are highly susceptible o Experimental infection with no clinical signs have been documented in splenectomised ungulates including mouflon (Ovis musimon), red deer (Cervus elaphus), roe deer (Capreolus capreolus), and fallow deer (Dama dama)
Life Cycle and Transmission
 BB is principally transmitted by means of ticks o Tick vectors of Babesia bigemina: Rhipicephalus microplus (formerly Boophilus microplus) and Rhipicephalus annulatus (formerly Boophilus annulatus); Rhipicephalus decoloratus, Rhipicephalus geigyi, and Rhipicephalus evertsi are also competent vectors  B. bigemina transmitted by feeding of adult and nymphal stages of one-host Rhipicephalus spp. ticks o Tick vectors of Babesia bovis: Rhipicephalus microplus and Rhipicephalus annulatus; Rhipicephalus geigyi is also a competent vector  B. bovis transmitted by feeding of larval stages of one-host Rhipicephalus spp. ticks
o Tick vectors of Babesia divergens: principal vector is Ixodes ricinus  Ixodes ricinus is a three-host tick with only adult stages feeding on vertebrates (eg. cattle)  Babesia sporozoites are inoculated into the vertebrate host by ticks and invade red blood cells (RBCs) where they transform into trophozoites o These grow and divide into two round, oval or pear-shaped merozoites which, in turn, are capable of infecting new RBCs; the division process is then repeated  Babesia parasites can be transmitted transovarially between tick generations; in the case of Ixodes, surviving up to 4 years without a vertebrate host  Babesia may also be transmitted by fomites and mechanical vectors contaminated by infected blood  Infrequently, calves can become infected in utero
Sources of infection
 Blood infected with Babesia parasites and associated vectors of infected blood (especially ticks, but also by mechanical means)
Occurrence
BB is found in areas where its arthropod vector is distributed, especially tropical and subtropical climates. Babesia bovis and B. bigemina are more widely distributed and of major importance in Africa, Asia, Australia, and Central and South America. Babesia divergens is economically important in some parts of Europe and possibly northern Africa.
For more recent, detailed information on the occurrence of this disease worldwide, see the OIE World Animal Health Information Database (WAHID) Interface [http://www.oie.int/wahis/public.php?page=home] or refer to the latest issues of the World Animal Health and the OIE Bulletin.
DIAGNOSIS
Incubation period is often 2–3 weeks or longer after tick infestation. Shorter incubation periods have however been documented in the field and through experimental inoculation (4–5 days for B. bigemina and 10–12 days for B. bovis).
Clinical diagnosis
Clinical manifestations of disease associated with BB are typical of a haemolytic anaemia disease process but vary according to agent (i.e. species of parasite) and host factors (i.e. age, immune status). BB is predominantly observed in adult cattle with B. bovis generally being more pathogenic than B. bigemina or B. divergens. Infected animals develop a life-long immunity against re-infection with the same species and some cross-protection is evident in B. bigemina-immune animals against subsequent B. bovis infections.
Babesia bovis
 High fever  Ataxia and incoordination  Anorexia  Production of dark red or brown-colored urine  Signs of general circulatory shock  Sometimes nervous signs associated with sequestration of infected erythrocytes in cerebral capillaries  Anaemia and haemoglobinuria may appear later in the course of the disease  In acute cases: maximum parasitaemia (percentage of infected erythrocytes) in circulating blood is often less than 1%
Babesia bigemina
 Fever
 Haemoglobinuria and anaemia  Production of dark red or brown-colored urine  Nervous signs minimal or non-existent as intravascular sequestration of infected erythrocytes does not occur  Parasitaemia often exceeds 10% and may be as high as 30%
Babesia divergens
 Parasitaemia and clinical appearance are similar to B. bigemina infections
Lesions
 Lesions observed are those most often associated with an intravascular haemolytic condition  Pale or icteric mucous membranes; blood may appear thin and watery   Subcutaneous tissues, abdominal fat and omentum may appear icteric  Swollen liver with an orange-brown or paler coloration; enlarged gall bladder containing thick, granular bile  Enlarged, dark, friable spleen   Kidneys appear darker than normal with possible petechial haemorrhages  Bladder may contain dark red or brown-colored urine  Possible oedema of lungs  Petechiae or ecchymoses on surface of heart and brain
Differential diagnosis
 Anaplasmosis  Trypanosomiasis  Theileriosis  Bacillary haemoglobinuria  Leptospirosis   Eperythrozoonosis   Rapeseed poisoning   Chronic copper poisoning
Laboratory diagnosis
Samples
 Several thick and thin blood smears collected from superficial skin capillaries (e.g. tip of the ear or tip of the tail) of live animals during the acute phase of the disease (appearance of fever) o thin blood films should be air-dried, fixed in absolute methanol for 1 minute  and stained with 10% Giemsa stain for 20–30  minutes  blood films should be stained as soon as possible after preparation to ensure proper stain definition o thick films are made by placing a small drop (approximately 50 µl) of blood on to a clean glass slide and spreading this over a small are using a circular motion eith the corner of another slide. The droplet is air-dried, heat-fixed at 80°C for 5 minutes, and stained (without fixing in methanol) in 10% Giemsa for 15 minutes o unstained blood films should not be stored with or near formalin solutions as formalin fumes may affect staining quality; moisture also affects staining quality.  If it is not possible to make fresh films from capillary blood, sterile jugular blood should be collected into an anticoagulant such as lithium heparin or ethylene diamine tetra-acetic acid (EDTA)  o The sample should be kept cool, preferably at 5°C, until delivery to the laboratory B. bovis is sequestered and found in higher numbers in capillary blood, B. bigemina and B. divergens parasites are uniformly distributed through the vasculature  Samples from dead animals should consist of thin blood films, as well as smears from organs  Organ smears acquired at necropsy: cerebral cortex, kidney (freshly dead), spleen (when decomposition is evident), heart muscle, lung and liver o organ smears are made by pressing a clean slide on to a freshly cut surface of the organ or by crushing a small sample of the tissue (particularly cerebral cortex)
between two clean microscope slides drawn lengthwise to leave a film of tissue on each slide o organ smear is then air-dried (assisted by gentle warming in humid climates), fixed for 5 minutes  in absolute methanol, and stained for 20–30  minutes in 10% Giemsa o especially suitable for the diagnosis of B. bovis infections using smears of cerebral cortex but unreliable if sample taken 24 hours or longer after death has occurred, especially in warmer weather  Babesia parasites can sometimes be detected in capillary blood taken from the lower limb region one or more days after death  Serum samples should also be collected
Procedures
Identification of the agent
 Microscopic examination of blood – traditional method of identifying agent in infected animals by microscopic examination of Giemsa-stained thick and thin blood films  o stained films are examined under oil immersion using (as a minimum) a ×8 eyepiece and a ×60 objective lens o morphology of Babesia described in various sources, including OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals o sensitivity of thick films can detect parasitaemias as low as 1 parasite in 106 red blood cells  o Babesia species differentiation is good in thin films but poor in the more sensitive thick films o adequate for detection of acute infections, but not for detection of carriers where parasitaemias are very low o parasite identification and differentiation improved by using a fluorescent dye, such as acridine orange instead of Giemsa  Nucleic acid-based diagnostic assays - very sensitive particularly in detecting B. bovis and B. bigemina in carrier cattle o a PCR- based techniques are reported to be at least 1000 times more sensitive than thin blood smears for detection of B. bovis o a number of PCR techniques have been described that can detect and differentiate species of Babesia in carrier infections o current PCR assays generally do not lend themselves well to large-scale testing; unlikely to supplant serological tests as the method of choice for epidemiological studies o PCR assays are useful as confirmatory tests and in some cases for regulatory testing  In-vitro culture methods o used to demonstrate presence of carrier infections of Babesia spp.; B. bovis has also been cloned in culture o minimum parasitaemia detectable by this method depends on the facilities available and the skills of the operator but could be as low as 10–10, making it a very sensitive method for the demonstration of infection, with 100% specificity  Animal inoculation is not suitable for diagnostic purposes
Serological tests
 Babesia bovis enzyme-linked immunosorbent assay o ELISA for diagnosis of B. bovis infection uses a whole merozoite antigen; undergone extensive evaluation o Competitive ELISAs using recombinant merozoite surface and rhoptry associated antigens of B. bovis have recently been developed o Reduction in specificity of the indirect B. bovis ELISA using recombinant antigens has been noted in some situations  Babesia bigemina enzyme-linked immunosorbent assay o a competitive ELISA developed and validated in Australia and USA are apparently the only ELISAs in routine use. It has been included in the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals o no other well-validated ELISA available for B. bigemina; due in part to the fact that antibodies to B. bigemina crude antigen typically have poor specificity o ELISAs have also been developed for B. divergens using antigen derived from culture, Meriones or cattle, but none has been validated internationally
o An immunochromato-graphic test for simultaneous rapid serodiagnosis of bovine babesiosis caused by B. bovis and B. bigemina was developed recently  Indirect fluorescent antibody (IFA) test o widely used in the past to detect antibodies to Babesia spp., but the B. bigemina test has poor specificity o cross-reactions with antibodies to B. bovis in the B. bigemina IFA test are a particular problem in areas where the two parasites coexist o disadvantages of low sample throughput and subjectivity  Complement fixation o has been used  to detect antibodies against B. bovis and B. bigemina o used to qualify animals for importation into some countries  Other tests: dot ELISA, slide ELISA, latex and card agglutination tests, and an immunochromatographic test o tests show acceptable levels of sensitivity and specificity for B. bovis and, in the case of the dot ELISA, also for B. bigemina o however, none of these tests appears to have been adopted for routine diagnostic use in laboratories other than those in which the original development and validation took place o adaptability of these tests to routine diagnostic laboratories is therefore unknown
For more detailed information regarding laboratory diagnostic methodologies , please refer to Chapter 2.4.2 Bovine babesiosis in  the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals  under the heading “Diagnostic Techniques” .
PREVENTION AND CONTROL
Sanitary prophylaxis
 Eradication of BB has been accomplished by elimination of tick vector and/or intensive chemotherapeutic regimes o in areas where eradication of tick is not feasible or desirable, ticks are controlled by repellents and acaricides  Reducing exposure of cattle to ticks o repellents, acaricides and regular inspection; animals and premises o control and eradication of the tick vector   Cattle develop a durable, long-lasting immunity after a single infection with B. bovis, B. divergens or B. bigemina, a feature that has been exploited in some countries to immunise cattle against babesiosis  Endemic environments should be monitored carefully  o introduction of immuno-naïve animals o introduction of new species or strains of disease agent o interruptions in exposure to ticks and disease due to changes in climate, host factors and management  Special care in possible mechanical infection of horses with contaminated blood
Medical prophylaxis
Vaccine for Babesia:
 Live vaccine: most live vaccines contain specially selected strains of Babesia (mainly B. bovis and B. bigemina) and are produced in calves or in vitro in government-supported production facilities as a service to the livestock industries o caution should be used in their employment as they may be virulent in adult animals, may be contaminated with other disease agents and could lead to hypersensitivity reactions; usually used in younger animals o an experimental B. divergens vaccine prepared from the blood of infected Meriones has also been used successfully  Killed vaccine: prepared from blood of B. divergens-infected calves; little information available on level and duration of the conferred immunity  Other vaccines:  o Despite the worldwide efforts, the prospects for recombinant vaccines against Babesia spp. Remain challenging o To date, no effective subunit vaccine is available commercially
o experimental vaccines containing antigens produced in vitro have been developed but the level and duration of protection against heterologous challenge are unclear
For more detailed information regarding vaccines, please refer to Chapter 2.4.2 Bovine babesiosis in the latest edition of the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals under the heading “Requirements for Vaccines”.
Endemic areas
 Clinically affected animals treated with an antiparasitic drug (diminazene diaceturate, imidocarb, amicarbalide); efficacy depends on timely detection early in disease  o Babesia parasites can be cleared from carrier animals; reduces clinical signs o Imidocarb has been reported to protect animals from disease but allow development of immunity; caution in regard to residues in milk and meat  Consideration can be given to blood transfusions and other supportive therapy, if appropriate
For more detailed information regarding safe international trade in terrestrial animals and their products, please refer to the latest edition of the OIE Terrestrial Animal Health Code.
REFERENCES AND OTHER INFORMATION
 Brown C. & Torres A., Eds. (2008). - USAHA Foreign Animal Diseases, Seventh Edition. Committee of Foreign and Emerging Diseases of the US Animal Health Association. Boca Publications Group, Inc.  Coetzer J.A.W. & Tustin R.C. Eds. (2004). - Infectious Diseases of Livestock, 2nd Edition. Oxford University Press.  Homer M.J. & et al. (2000) - Clin. Microbiol. Rev., 13 (3): 451.  Kahn C.M., Ed. (2005). - Merck Veterinary Manual. Merck & Co. Inc. and Merial Ltd.   Spickler A.R., & Roth J.A. Iowa State University, College of Veterinary Medicine - http://www.cfsph.iastate.edu/DiseaseInfo/factsheets.htm  World Organisation for Animal Health (2012). - Terrestrial Animal Health Code. OIE, Paris.  World Organisation for Animal Health (2012). - Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris.
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The OIE will periodically update the OIE Technical Disease Cards. Please send relevant new references and proposed modifications to the OIE Scientific and Technical Department (scientific.dept@oie.int). Last updated April 2013.