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trypanosomosis

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trypanosomosis

Summary

  • Last modified
  • 12 October 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • trypanosomosis
  • Overview
  • Trypanosomes are microscopic unicellular protozoan flagellates in the genus Trypansoma. They are obligatory parasites of vertebrates, and infect fish and amphibian species, reptiles, birds and mammals (ltard,...

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Pictures

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PictureTitleCaptionCopyright
Camel affected by Trypanosoma infection in Sudan.
TitleSymptoms: Camel
CaptionCamel affected by Trypanosoma infection in Sudan.
CopyrightCIRAD-EMVT
Camel affected by Trypanosoma infection in Sudan.
Symptoms: CamelCamel affected by Trypanosoma infection in Sudan.CIRAD-EMVT
African trypanosomiasis. Severe emaciation.
TitleExternal symptoms
CaptionAfrican trypanosomiasis. Severe emaciation.
Copyright©USDA-2002/Foreign Animal Diseases Training Set/USDA-Animal and Plant Health Inspection Service (APHIS)
African trypanosomiasis. Severe emaciation.
External symptomsAfrican trypanosomiasis. Severe emaciation.©USDA-2002/Foreign Animal Diseases Training Set/USDA-Animal and Plant Health Inspection Service (APHIS)
Trypanosomiasis or 'sleeping sickness' symptoms in young boy.
TitleSymptoms
CaptionTrypanosomiasis or 'sleeping sickness' symptoms in young boy.
CopyrightDFID Animal Health Programme
Trypanosomiasis or 'sleeping sickness' symptoms in young boy.
SymptomsTrypanosomiasis or 'sleeping sickness' symptoms in young boy.DFID Animal Health Programme
African trypanosomiasis. Parasites in a blood smear.
TitleHistopathology
CaptionAfrican trypanosomiasis. Parasites in a blood smear.
Copyright©USDA-2002/Foreign Animal Diseases Training Set/USDA-Animal and Plant Health Inspection Service (APHIS)
African trypanosomiasis. Parasites in a blood smear.
HistopathologyAfrican trypanosomiasis. Parasites in a blood smear.©USDA-2002/Foreign Animal Diseases Training Set/USDA-Animal and Plant Health Inspection Service (APHIS)

Identity

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Preferred Scientific Name

  • trypanosomosis

International Common Names

  • English: hemolytic anemia due rbc parasitism; nagana; sleeping sickness; surra; surra, trypanosoma evansi, in cattle - exotic; surra, trypanosoma evansi, in pigs - exotic; trypanosomiasis; trypanosomiasis in ruminants

Overview

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Trypanosomes are microscopic unicellular protozoan flagellates in the genus Trypansoma. They are obligatory parasites of vertebrates, and infect fish and amphibian species, reptiles, birds and mammals (ltard, 1989). Trypanosomes were first found in frog’s blood and described by Gruby in 1843. The taxonomic position of the genus Trypanosoma was reported by Hoare (1972).

Based on the nature of their life cycle and transmission, pathogenic trypanosomes may be classified as stercorarian or salivarian (Siegmund, 1979; Losos, 1986). The stercorarian trypanosomes have a cycle of development in an insect vector. The infective form is transmitted to the mammalian host in the faeces of the vector. The salivarian trypanosomes follow a cycle of development in tsetse flies (Glossina spp.), and the infective metacyclic forms are transmitted to the mammalian hosts through the vectors’ saliva during their haematophagous bites (ltard, 1989). The stercorarian trypanosomes are in the subgenera Megatrypanum and Schizotrypanum, while the salivarian trypanosomes are in the subgenera Duttonella, Nannomonas, Trypanozoon and Pycnomonas.

The pathogenic salivarian species and their animal hosts are as follows:

Trypanosoma (Duttonella) vivax - ruminants, horses, camels

T. (D.) uniforme – ruminants

T. (Nannomomas) congolense - ruminants, horses, dogs and cats

T. (N.) simiae - Pigs and camels

T. (Trypanozoon) brucei brucei - all domestic mammals, including ruminants, horses, dogs, cats, pigs

T. (T.) brucei gambiense – Man

T. (T.) brucei rhodesiense – Man

T. (T.) evansi evansi - Camels, horses, pigs, dogs, cats, domestic buffaloes, ruminants

T. (T.) evansi equinum - equine species

T. (T.) equiperdum - horses, donkeys

T. (Pycnomonas) suis - pigs.

The trypanosomes transmitted biologically or cyclically are T. brucei subspecies, T. vivax, T. congolense, T. simiae, T. uniforme and T. suis. Mechanical transmission, as seen with horse flies - Tabanidae and Stomoxys (Hoare, 1947; Mwongela et al., 1981), occurs with T. vivax and T. evansi. Although under laboratory conditions mechanical spread of T. congolense has been noted (Desquesnes and Dia, 2003), due to low parasitaemia levels it is unlikely to be epidemiologically important in the field under natural conditions (Desquesnes et al., 2009). T. equiperdum, transmitted during coitus, is the only trypanosome that does not require an insect vector for transmission. Transplacental transmission of T. vivax and T. evansi has been reported in domestic animals (Ikede and Losos, 1972a; Ogwu et al., 1986) and T. brucei has been reportedly transmitted to dogs and cats by feeding on infected goat meat (Moloo et al., 1973).

The disease caused by the trypanosomes (trypanosomiasis or trypanosomosis) was first studied in South Africa by Sir David Bruce where it was called nagana (Joubert et al., 1993; Cook, 1994). Animal trypanosomosis, now generally termed as nagana (Itard, 1989), is a constraint to livestock production in parts of Africa, where tsetse flies infest a vast region of sub Saharan Africa, including 37 countries totalling approximately 9 million km2 and 50 million cattle at risk (Cecchi and. Mattioli, 2009).

The disease has assumed greater importance in South America, with regard to mechanically transmitted T. vivax infection (Otte et al., 1994; Payne et al., 1994; Vargas-Teran and Arellano-Sota, 1997; Silva et al., 1988 a, b, c).

T. evansi the causative agent of surra is found in countries where camels are present in northern Africa. It spread to the Arabian Peninsula and later to India, Russia, China, Mongolia and South East Asia (Lun et al., 1993; Ling, 1996; Manuel et al., 1988; Abo-Shehada et al., 1999; Cheah et al., 1999; Pholpark et al., 1999). Surra spread to South America in the 15th century and is now present in in many of the countries of South and Central America, including Argentina, Brazil, Bolivia and Mexico (Desquesnes et al. 2013) Within the Americas as well as being spread mechanically by various biting flies, it can also be transmitted by the vampire bat (Desmodus rotundus) (Hoare, 1972). More recently T. evansi has spread to the Canary Islands and has often been observed since 1995, while cases have been recorded in France (Desquesnes et al. 2008) and Spain (Tamarit et al. 2010) in camels imported from the Canary Islands. T. evansi has the widest host range of any of the salivarian trypanosomes. It is highly pathogenic in camels and Equidae and within South East Asia it infects water buffaloes and many wildlife species including deer, elephants, jaguars, tigers and lions (Desquesnes et al. 2013). Within South America, horses are severely affected, although cattle can become infected they rarely show clinical signs. Wildlife including capybara, coatis and lamas have also found to be infected. Dourine, the venereal contagious disease caused by T. equiperdum in equines, exists in North and South Africa, and parts of Asia and South America (Losos, 1986).

The stercorarian trypanosomes that have been reported to cause disease in animals are T. (Megatrypanum) theileri (Monzon et al., 1993), Trypanosoma (Herpetosoma) lewisi and T. (Schizotrypanum) cruzi.

T. theileri, which is usually non-pathogenic, is a parasite of cattle, buffalo and antelope worldwide (Itard, 1989). However, a disease caused by T. theileri resembling nagana has been reported in a calf in Ireland (Doherty et al., 1993), in cattle in Formosa province, Argentina (Monzon et al., 1993) and in a cow in Iran (Seifi, 1995). T. theileri is known to cause disease in cattle that are severely stressed by a concurrent disease and poor nutrition (Radostits et al., 1994).

T. lewisi was described first by Kent in 1880 and then by Laveran and Mesnil in 1901. It is a parasite of rats (Rattus rattus and Rattus norvegicus), that in temperate countries is transmitted by fleas: Nosopsyllus fasciatus, and in inter-tropical countries by Xenopsylla cheopis (Hoare, 1972). T. lewisi can also develop in other species of fleas including the dog flea (Ctenocephalides canis), the mouse flea (Leptopsylla segnis) and the human flea (Pulex irritans). Rats become infected when they ingest T. lewisi either through flea faeces or the fleas themselves. Generally infections are restricted to rats, with infections being non-pathogenic. There have been several cases of human infections, these are discussed by Truc et al. (2013).

T. cruzi, which causes Chagas disease in human beings, has been reported to precipitate natural acute syndromes in dogs in the USA, with various features similar to the human syndrome (Williams et al., 1977; Losos, 1986; Rowland et al., 2010). Experimental infections with culture forms of T. cruzi in pigs, lambs, goats and calves have been reported (Diamond and Rubin, 1958; Fernandes et al., 1994), but such infections are usually not accompanied by clinical signs and lesions in organs.

The infections cause anaemia, leucopenia, thrombocytopenia and inflammatory and degenerative lesions in the heart, liver, lymph nodes, testes, brain, conjunctiva, cornea, spleen, kidney and some endocrine glands (Losos, 1986; Anosa, 1988; Murray and Dexter, 1988; Itard, 1989; Ogwu et al., 1992; Saseendranath et al, 1995). Infection of dams causes abortion (Edeghere et al., 1992; Le et al., 1994; Rowlands et al., 1994, 1995; Elhassan et al., 1995; Kalu, 1996; Okech et al., 1996a, b; Goossens et al., 1997a), premature birth (Okech et al., 1996a), stillbirth (Okech et al., 1996c) and delivery of calves and lambs which died shortly after birth (Okech et al., 1996a; Elhassan et al., 1995). Infected animals have lower pregnancy rates and longer intervals between breeding (Okech et al., 1996a; Osaer et al., 1998b) compared with uninfected animals. Infertility in infected male animals is due to orchitis (Anosa and Isoun, 1980; Saseendranath et al., 1995), decreased semen output and poor semen quality (Sekoni, 1992, 1993; Boly et al., 1993, 1996; Stefano et al., 1999). Other adverse effects of infection in animals include deaths (Radostits et al., 1994), decreased milk production (Agyemang et al., 1990; Joshi et al., 1993; Pholpark et al., 1999), reduction of growth rate, delay of puberty, delayed age at first lambing and poor feed conversion efficiency (Ilemobade and Balogun, 1981; Payne et al., 1994; Otte et al., 1994; Osaer et al., 1998a, 1999). There is also a depressed immune response for protection to common diseases such as helminthiasis (Kaufmann et al., 1992; Dwinger et al., 1994; Sharma et al., 2000) and pasteurellosis (Phan et al., 1996). Infected animals are unproductive in terms of use for draught (Radostits et al., 1994; Bennison et al., 1998; Pearson et al., 1999).

The importance of animal trypanosomosis in endemic areas is hinged on the effect of the disease on production indices and the cost of controlling the disease to allow profitable animal production. The prevalence of the disease is less than 5% in endemic areas, except where a vector control programme has been mounted (Bauer et al., 1999; Hargrove et al., 2000), along with the use of chemoprophylaxis, chemotherapy and trypanotolerant livestock (Holmes et al., 1997).

This disease is on the list of diseases notifiable to the World Organisation for Animal Health (OIE). The distribution section contains data from OIE's WAHID database on disease occurrence. Please see the AHPC library for further information on this disease from OIE, including the International Animal Health Code and the Manual of Standards for Diagnostic Tests and Vaccines. Also see the website: www.oie.int

Hosts/Species Affected

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The most important hosts of the African trypanosomes are domestic livestock including cattle and small ruminants. Camels can be affected by T. evansi. Many wildlife species also carry trypanosomes and play important roles as reservoirs of infection for domestic livestock (see Anderson et al. (2011) and references therein for information on wildlife that can be infected by trypanosomes).

T. cruzi has a broad host range and has been reported to infect more than 150 mammal species including bats and it is thought that all mammalian species are susceptible. Birds and reptiles, despite being fed on by the vectors, do not appear to be susceptible to infection.

Trypanosomes of Australian mammals have been reviewed by Thompson et al. (2014) while Paparini et al. (2011) discuss trypanosomes found in marsupials.

Trypanosomes of fish are split into two types; these are designated Type A and Type B and are transmitted by leeches (Overath et al. 1999).

Trypanotolerance

Trypanotolerance is defined as the relative capacity of an animal to control the multiplication of the trypanosome in their blood and resist the development of pathological effects (D’leteren et al., 1998). Most wild animals and some domestic animals are trypanotolerant because they are able to remain clinically normal with trypanosome infections for long periods. These animals can tolerate challenge from the tsetse flies. The trypanotolerant cattle breeds of West Africa include the long-horned taurine N’dama breed, short-horned taurine Baoulé, Lagune, Somba, Lobi, Pabli, Namji (Namchi) breeds and taurine crosses with zebu such as Borgou and Djakoré cattle (Itard, 1989; Maillard et al., 1992; Verhulst et al., 1993; Achukwi et al., 1997; Doko et al., 1997a). The trypanosome tolerance of cattle is discussed by Naessens (2006). The trypanotolerant sheep and goats are the Djallonké, West African Dwarf, Guinea and Casamance breeds (Itard, 1989; Adah et al., 1993; Osaer et al., 1994). The trypanotolerance of small ruminants has been reviewed by Geerts et al. (2009). Kirdi ponies show some degree of trypanotolerance (Itard, 1989). The trait of trypanotolerance is attributed to mechanisms that relate to genetic, immunological, cellular, physiopathological and ecological factors (Itard, 1989).

Trypanotolerant breeds develop premunition when they are in constant contact with infected tsetse flies and have the ability to stimulate more rapid and stronger immune responses than susceptible breeds (Authie et al., 1993; Sileghem et al., 1993; Williams et al., 1996). The premunition to infection may be lost when the animals are exposed to new trypanosome species and strains (Williams et al., 1992), intestinal parasitism (Goossens et al., 1997b), poor nutrition and excessive fatigue (Itard, 1989).

The ability of N’dama cattle to resist the development of anaemia in trypanosomosis may be associated with their possession of erythrocyte surface sialic acids in amounts seven times greater than the Zebu breeds (Esievo et al., 1986). The erythrocytes of both Zebu and N’dama breeds had N-acetylneuraminic acid, but N’dama erythrocytes had an additional unidentified type of sialic acid (Esievo et al., 1990), which was considered responsible for the higher erythrocyte surface sialic acid concentrations in N’dama cattle. Shugaba et al. (1994) showed that the N’dama erythrocyte sialic acids had more O-acetyl than glycolyl groups, which may be relevant to trypanotolerance, since O-acetyl groups in sialic acids might hinder rapid hydrolysis. Also, their ability to control anaemia and parasitaemia in trypanosome infection was attributable to their greater efficiency in increasing progenitors from the erythroid and myeloid lineages in the bone marrow compared to the susceptible Boran cattle (Andrianarivo et al., 1994).

Trypanotolerance has been associated with significantly higher erythrocyte levels of zinc and manganese in Keteku cattle, compared to White Fulani cattle (Awolaja et al., 1997).

The high ability of the African buffalo to control trypanosome infections is due to the presence of a trypanocidal factor in the blood (Reduth et al., 1994; Muranjan et al., 1997). This has been identified as xanthine oxidase, which inhibits trypanosome glycolysis (Muranjan et al., 1997).

Differences in trypanosusceptibility

Masai Zebu and Orma Boran cattle are less susceptible than Galana Boran cattle to trypanosome infection (Mwangi et al., 1998). Zebu cattle are more resistant to anaemia caused by trypanosome infection, than Holstein cattle (Doko et al., 1997b). Small East African goats are more resistant than Kigezi and Mubende goats to T. congolense infection (Katunguka-Rwakishaya et al., 1997c). Scottish Blackface lambs develop a more severe disease than Finn Dorset lambs when, infected with T. congolense (Katunguka-Rwakishaya et al., 1997d).

Age of animal

Calves that are less than a year old are more resistant than adults to the effects of trypanosomiasis (Fiennes, 1970; Maxie and Valli, 1979; Wellde et al., 1981; Murray et al., 1982). This is because the calves develop a lower parasitaemia (Wellde et al., 1981) or have superior erythropoietic responses (Murray and Dexter, 1988) compared to the adults. However, lambs and kids were more susceptible to T. congolense infection than the adults (Griffin and Allonby, 1981). Kalu (1995) reported the highest prevalence of trypanosomosis among older cattle aged 6-9 years.

Nutritional Status

Poor levels of nutrition can result in more severe trypanosome infections, even among trypanotolerant breeds (Chandler, 1952; Stephen, 1966; Reynolds and Ekwuruke, 1988; Malik et al., 2000; Osaer et al., 2000). Low-energy intake causes a more severe anaemia in trypanosome-infected animals than in those infected animals with a high-energy intake (Fagbemi et al., 1990; Katunguka-Rwakishaya et al., 1995). Low protein diet did not aggravate the disease, but erythropoietic activity was greater in animals on a high protein diet (Katunguka-Rwakishaya et al., 1993; Katunguka-Rwakishaya, 1996). Therefore, adequate energy and protein nutrition enhances the ability of infected animals to withstand the adverse effects of the infection (Katunguka-Rwakishaya et al., 1993; Little et al., 1994; Katunguka-Rwakishaya et al., 1995, 1997a, b). During the dry season, when food is of an inadequate quality and quantity, clinical disease may become more severe and relapses of infection after treatment may be more frequent (Itard, 1989).

Vectoral and environmental factors

The role of tsetse flies in the epidemiology of trypanosomosis has been reviewed (Rotureau and Van Den Abbeele, 2013). The capacity of the different tsetse flies species to become infected and transmit the trypanosome after its development (vector capacity) varies. There are differences in the percentage of flies infected within a population (infection rate). For example, riverine tsetse flies are more commonly infected with T. vivax than T. congolense. The more complex the lifecycle of the trypanosome species, the lower the vector infection rate. Therefore, the infection rate is highest for T. vivax, followed by T. congolense and T. brucei. The savannah species of tsetse flies (G. morsitans) usually have higher infection rates than riverine forest species (G. palpalis, G. fuscipes and G. tachinoides) and are more efficient vectors.

Infection rates are higher in environments with favourable conditions for tsetse fly survival. Also in areas that have higher numbers of female tsetse flies than males, because females have a longer lifespan and they take bigger bloodmeals than the males. During the rainy season infection rates are high, because humidity and temperature conditions increase the tsetse lifespan and shorten the life cycle of the trypanosomes. Therefore, prevalence of the disease is often higher in the rainy season (Kalu, 1995). In the hot season the lifespan of tsetse flies is too short to allow the trypanosomes to reach the infective stage. The infection rates are lowest in the cold season because the flies are not active and the life cycle of the trypanosomes is longer.

Apparent tsetse fly density in an area is influenced by season, the availability of favourable habitats and the presence of domestic and wild animals to provide bloodmeals. The tsetse density is directly correlated with the prevalence of trypanosomosis (Belot and Leroy, 1998; Kamau et al., 2000). Therefore, the risk of trypanosome infection is increased by transhumance involving the movement of animals through tsetse-infected vegetation and raising livestock near habitats that favour tsetse fly multiplication or in the midst of trypanosome-infected domestic or wild animals (Radostits et al., 1994).

Distribution

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The trypanosome species, which are found only in the tsetse fly belt of Africa are T. brucei subspecies, T. uniforme, T. congolense and T. simiae. There are no reports of such species of trypanosomes in areas where only mechanical transmission is possible because of the need for the tsetse biological cycle. T. vivax and T. evansi are maintained solely by mechanical transmission in some parts of Africa and in other parts of the world.

The trypanosome species that cause animal trypanosomosis (nagana) have been reported in many countries (see Distribution table) including Nigeria (Ahmed et al., 1994; Kalu, 1995; Kalejaiye et al., 1995; Jibike et al., 1995; Anene and Ezekwe, 1995; Nawathe et al., 1995; Basu et al., 1995; Kalu and Lawani, 1996; Ogunsanmi et al., 2000), the Gambia (Leperre and Claxton, 1994), Benin (Pandey et al., 1993), Ghana (Turkson, 1993), Burkina Faso (Bengaly et al., 1995, 1998; Desquesnes et al., 1999), Malawi (Bossche et al., 2000), Namibia (Bossche et al., 1999), Zimbabwe (Bossche and Mudenge, 1997), Uganda, Mali, Senegal, Niger, Chad, Mauritania and Zaire (Itard, 1989).

Nagana caused by T. vivax was introduced into French Guyana and the West Indies (Itard, 1989) by imported Senegalese Zebu cattle in 1830, and has been remained in South America by mechanical transmission through biting flies. Outbreaks of T. vivax infections have been reported in Brazil (Silva et al., 1996, 1998a, 1998c, 1999, Cadiloi et al., 2012), Venezuela (Sandoval et al., 1998), Bolivia (Silva et al., 1998a, b), Colombia (Otte et al., 1994), Costal Guyana (Vokaty et al., 1993), French Guyana (Desquesnes and Gardiner, 1993) and Costa Rica (Oliveria et al., 2009). Genetic analysis of New World T. vivax supports their introduction from West Africa (Cortez et al., 2006; Garcia et al., 2014).

Surra caused by T. evansi is found in African countries such as Morocco, Algeria, Tunisia, Libya, Egypt, Mali, Niger, northern Nigeria, Chad, Sudan, Kenya, Somalia and Ethiopia (Itard, 1989). The disease exists in the Middle East (Arabian Peninsula, Turkey), central and southern Russia, India, Indonesia, Philippines, Southern China, and South and Central America (Itard, 1989). The prevalence of surra has been reported in China (Lun et al., 1993; Ling, 1996), India (Batra et al., 1994; Reddy and Hafeez, 1996; Das et al., 1998; Dhami et al., 1999), Vietnam (Hoang et al., 1996; Thu et al., 1998; My et al., 1998; Pham et al., 1999; Verloo et al., 2000), Indonesia (Payne et al., 1994; Luckins et al., 1998; Davison et al., 2000), Thailand (Tuntasuvan et al., 1998; Pholpark et al., 1999), Venezuela (Aray et al., 1998), Jordan (Abo-Shehada et al., 1999), Malaysia (Arunasalam et al., 1995; Cheah et al., 1999), Philippines (Manuel et al., 1998) and Coastal Guyana (Vokaty et al., 1993).

Dourine caused by T. equiperdum is found in central Italy, northern Africa, sub-Saharan Africa, Turkey, Soviet Asia, Siberia, Iran, Iraq, Arabian Peninsula, Indonesia and South American countries such as Chile, Venezuela and Brazil (Itard, 1989).

For current information on disease incidence, see OIE's WAHID Interface.

During 2011, 21 countries in Africa reported to AU-IBAR outbreaks of trypanosomosis, compared to 23 countries in 2010 and 19 countries in 2009. The 21 countries recorded 1,629 outbreaks involving 195,923 cases and 6,472 deaths (see table below). Countries with the highest number of cases included Benin (83,257), Uganda (80,850) and Congo DRC (19,902).

Country

Outbreaks

Cases

Deaths

Slaughtered

Destroyed

Benin

477

83,257

1,008

993

11

Botswana

7

10

2

0

0

Cameroon

5

121

1

0

 

Congo DRC

32

19,902

3,696

520

78

Côte d'Ivoire

4

998

556

27

229

Egypt

71

277

 

 NS

 NS

Ethiopia

2

33

18

0

 

Gambia

3

204

14

76

0

Ghana

15

132

11

28

0

Kenya

224

367

24

0

0

Mozambique

3

28

0

0

0

Namibia

7

10

0

0

0

Nigeria

1

7

0

0

0

Sierra Leone

5

257

105

0

0

Somalia

476

3,579

321

31

94

South Africa

5

 NS

 NS

 NS

 NS

Tanzania

21

3,155

21

 NS

 NS

Togo

84

906

38

5

8

Uganda

130

80,850

444

584

33

Zambia

39

1732

187

 NS

 NS

Zimbabwe

18

98

26

 NS

 NS

Total

1,629

195,923

6,472

2,264

453

NS=Not specified

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

AfghanistanNo information availableOIE, 2009
ArmeniaDisease not reportedOIE, 2009
AzerbaijanDisease never reportedOIE, 2009
BahrainDisease never reportedOIE, 2009
BangladeshDisease never reportedOIE, 2009
BhutanDisease never reportedOIE, 2009
Brunei DarussalamNo information availableOIE Handistatus, 2005
CambodiaNo information availableOIE, 2009
ChinaRestricted distributionOIE, 2009
-AnhuiPresentLun et al., 1993; Ling YingJi, 1996
-GuangdongPresentLun et al., 1993
-GuangxiPresentLun et al., 1993
-Hong KongNo information availableOIE, 2009
-HubeiPresentLun et al., 1993
-HunanPresentLun et al., 1993
-JiangsuPresentLun et al., 1993
-JiangxiPresentLun et al., 1993
-XinjiangPresentLun et al., 1993
-YunnanPresentLun et al., 1993
-ZhejiangPresentLun et al., 1993
Georgia (Republic of)Disease never reportedOIE Handistatus, 2005
IndiaRestricted distributionOIE, 2009
-Andhra PradeshPresentReddy and Hafeez, 1996; Das et al., 1998
-HaryanaPresentBatra et al., 1994
-Indian PunjabPresentDhami et al., 1999
IndonesiaDisease not reportedOIE, 2009
-JavaPresentPayne et al., 1994; Luckins et al., 1998; Davison et al., 2000
IranDisease never reportedNULLSeifi, 1995; OIE, 2009
IraqDisease not reportedOIE, 2009
IsraelDisease not reportedOIE, 2009
JapanDisease not reportedOIE, 2009
JordanDisease not reportedNULLAbo-Shehada et al., 1999; OIE, 2009
KazakhstanDisease never reportedOIE, 2009
Korea, DPRDisease not reportedOIE Handistatus, 2005
Korea, Republic ofDisease never reportedOIE, 2009
KuwaitDisease not reportedOIE, 2009
KyrgyzstanDisease never reportedOIE, 2009
LaosDisease not reportedOIE, 2009
LebanonDisease not reportedOIE, 2009
MalaysiaDisease not reportedNULLArunasalam et al., 1995; Cheah et al., 1999; OIE, 2009
-Peninsular MalaysiaDisease not reportedOIE Handistatus, 2005
-SabahDisease never reportedOIE Handistatus, 2005
-SarawakDisease never reportedOIE Handistatus, 2005
MongoliaNo information availableOIE, 2009
MyanmarDisease never reportedOIE, 2009
NepalNo information availableOIE, 2009
OmanDisease not reportedOIE, 2009
PakistanPresentOIE, 2009
PhilippinesDisease never reportedNULLManuel, 1998; OIE, 2009
QatarAbsent, reported but not confirmedOIE, 2009
Saudi ArabiaDisease not reportedOIE, 2009
SingaporeDisease never reportedOIE, 2009
Sri LankaDisease never reportedOIE, 2009
SyriaDisease not reportedOIE, 2009
TaiwanDisease not reportedOIE Handistatus, 2005
TajikistanDisease not reportedOIE, 2009
ThailandNo information availableOIE, 2009
TurkeyNo information availableOIE, 2009
TurkmenistanDisease not reportedOIE Handistatus, 2005
United Arab EmiratesDisease never reportedOIE, 2009
UzbekistanDisease not reportedOIE Handistatus, 2005
VietnamPresentOIE, 2009
YemenNo information availableOIE, 2009

Africa

AlgeriaDisease not reportedOIE, 2009
AngolaDisease not reportedOIE, 2009
BeninPresentOIE, 2012
BotswanaPresentAU-IBAR, 2011
Burkina FasoPresentOIE, 2012
BurundiPresentOIE, 2012
CameroonPresentAU-IBAR, 2011
Cape VerdeDisease never reportedOIE, 2012
Central African RepublicPresentOIE, 2012
ChadNo information availableOIE, 2009
ComorosDisease never reportedOIE, 2012
CongoAbsent, reported but not confirmedOIE, 2009
Congo Democratic RepublicPresentAU-IBAR, 2011
Côte d'IvoirePresentOIE, 2012
DjiboutiDisease never reportedOIE, 2012
EgyptPresentOIE, 2012
Equatorial GuineaOIE, 2012
EritreaPresentOIE, 2012
EthiopiaPresentOIE, 2012
GabonPresentOIE, 2012
GambiaPresentAU-IBAR, 2011
GhanaPresentOIE, 2012
GuineaNo information availableOIE, 2009
Guinea-BissauPresentOIE, 2012
KenyaAbsent, reported but not confirmedOIE, 2012
LesothoDisease never reportedOIE, 2012
LibyaLast reported2004OIE, 2012
MadagascarDisease never reportedOIE, 2012
MalawiPresentOIE, 2012
MaliNo information availableOIE, 2009
MauritiusDisease not reportedOIE, 2009
MoroccoDisease never reportedOIE, 2012
MozambiquePresentOIE, 2012
NamibiaDisease not reported2005Bossche et al., 1999; OIE, 2009
NigerPresentOIE, 2012
NigeriaPresentAhmed et al., 1994; Jibike et al., 1995; Kalu, 1995; Nawathe et al., 1995; Ogunsanmi et al., 2000; OIE, 2012
RéunionDisease never reportedOIE Handistatus, 2005
RwandaPresentOIE, 2012
Sao Tome and PrincipeDisease not reportedOIE Handistatus, 2005
SenegalDisease not reported200705OIE, 2009
SeychellesDisease not reportedOIE, 2012
Sierra LeonePresentAU-IBAR, 2011
SomaliaNo information availableOIE, 2012
South AfricaPresentOIE, 2012
SudanLast reported2006OIE, 2009
SwazilandDisease not reportedOIE, 2009
TanzaniaPresentOIE, 2012
TogoPresentOIE, 2012
TunisiaLast reported1989OIE, 2012
UgandaPresentOIE, 2012
ZambiaPresentOIE, 2012
ZimbabwePresentBossche and Mudenge, 1997; OIE, 2012

North America

BermudaDisease not reportedOIE Handistatus, 2005
CanadaDisease never reportedOIE, 2009
GreenlandDisease never reportedOIE, 2009
MexicoDisease never reportedOIE, 2009
USADisease never reportedOIE, 2009

Central America and Caribbean

BarbadosDisease never reportedOIE Handistatus, 2005
BelizeDisease never reportedOIE, 2009
British Virgin IslandsDisease never reportedOIE Handistatus, 2005
Cayman IslandsDisease never reportedOIE Handistatus, 2005
Costa RicaDisease not reportedOIE, 2009
CubaDisease never reportedOIE, 2009
CuraçaoDisease not reportedOIE Handistatus, 2005
DominicaDisease not reportedOIE Handistatus, 2005
Dominican RepublicDisease never reportedOIE, 2009
El SalvadorDisease not reportedOIE, 2009
GuadeloupeNo information availableOIE, 2009
GuatemalaNo information availableOIE, 2009
HaitiDisease never reportedOIE, 2009
HondurasDisease never reportedOIE, 2009
JamaicaNo information availableOIE, 2009
MartiniqueNo information availableOIE, 2009
NicaraguaDisease never reportedOIE, 2009
PanamaNo information availableOIE, 2009
Saint Kitts and NevisDisease never reportedOIE Handistatus, 2005
Saint Vincent and the GrenadinesDisease never reportedOIE Handistatus, 2005
Trinidad and TobagoDisease never reportedOIE Handistatus, 2005

South America

ArgentinaDisease never reportedOIE, 2009
BoliviaNo information availableNULLSilva et al., 1998a; Silva et al., 1998b; OIE, 2009
BrazilDisease never reportedOIE, 2009
-Mato GrossoPresentSilva et al., 1996; Silva et al., 1999
-Mato Grosso do SulPresentSilva et al., 1998a; Silva et al., 1998c
ChileDisease never reportedOIE, 2009
ColombiaPresentNULLOtte et al., 1994; OIE, 2009
EcuadorDisease never reportedOIE, 2009
Falkland IslandsDisease never reportedOIE Handistatus, 2005
French GuianaDisease not reported1996Desquesnes and Gardiner, 1993; OIE, 2009
GuyanaReported present or known to be presentVokaty et al., 1993; OIE Handistatus, 2005
ParaguayNo information availableOIE Handistatus, 2005
PeruDisease never reportedOIE, 2009
UruguayDisease never reportedOIE, 2009
VenezuelaPresentNULLSandoval et al., 1998; OIE, 2009

Europe

AlbaniaNo information availableOIE, 2009
AndorraDisease never reportedOIE Handistatus, 2005
AustriaNo information availableOIE, 2009
BelarusDisease never reportedOIE, 2009
BelgiumDisease not reportedOIE, 2009
Bosnia-HercegovinaDisease not reportedOIE Handistatus, 2005
BulgariaDisease never reportedOIE, 2009
CroatiaDisease never reportedOIE, 2009
CyprusDisease never reportedOIE, 2009
Czech RepublicDisease never reportedOIE, 2009
DenmarkDisease never reportedOIE, 2009
EstoniaDisease never reportedOIE, 2009
FinlandDisease never reportedOIE, 2009
FranceDisease never reportedOIE, 2009
GermanyDisease never reportedOIE, 2009
GreeceDisease not reportedOIE, 2009
HungaryDisease never reportedOIE, 2009
IcelandDisease never reportedOIE, 2009
IrelandDisease never reportedNULLDoherty et al., 1993; OIE, 2009
Isle of Man (UK)Disease never reportedOIE Handistatus, 2005
ItalyDisease never reportedNULLOIE, 2009
JerseyDisease never reportedOIE Handistatus, 2005
LatviaDisease never reportedOIE, 2009
LiechtensteinDisease not reportedOIE, 2009
LithuaniaDisease never reportedOIE, 2009
LuxembourgDisease never reportedOIE, 2009
MacedoniaDisease never reportedOIE, 2009
MaltaDisease never reportedOIE, 2009
MoldovaDisease never reportedOIE Handistatus, 2005
MontenegroDisease never reportedOIE, 2009
NetherlandsDisease never reportedOIE, 2009
NorwayDisease never reportedOIE, 2009
PolandDisease never reportedOIE, 2009
PortugalDisease never reportedOIE, 2009
RomaniaNo information availableOIE, 2009
Russian FederationDisease never reportedOIE, 2009
-Central RussiaPresent
-Eastern SiberiaPresent
-Southern RussiaPresent
-Western SiberiaPresent
SerbiaNo information availableOIE, 2009
SlovakiaDisease not reportedOIE, 2009
SloveniaDisease never reportedOIE, 2009
SpainDisease not reportedOIE, 2009
SwedenDisease never reportedOIE, 2009
SwitzerlandDisease never reportedOIE, 2009
UKDisease never reportedOIE, 2009
-Northern IrelandDisease never reportedOIE Handistatus, 2005
UkraineDisease never reportedOIE, 2009
Yugoslavia (former)No information availableOIE Handistatus, 2005
Yugoslavia (Serbia and Montenegro)Disease not reportedOIE Handistatus, 2005

Oceania

AustraliaDisease never reportedOIE, 2009
French PolynesiaDisease never reportedOIE, 2009
New CaledoniaDisease never reportedOIE, 2009
New ZealandDisease never reportedOIE, 2009
SamoaDisease never reportedOIE Handistatus, 2005
VanuatuDisease never reportedOIE Handistatus, 2005
Wallis and Futuna IslandsNo information availableOIE Handistatus, 2005

Pathology

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If there is severe anaemia, the carcass is pale. Where the animal suffered severe loss of weight and emaciation, there may be oedematous swellings in the lower part of the abdomen and genital organs, and serious atrophy of fat. The liver, spleen and lymph nodes are enlarged and the viscera are congested. Petechiae may appear on lymph nodes, pericardium and intestinal mucosa.

The liver is hypertropic and congested, with degeneration and necrosis of the hepatocytes, dilation of blood vessels and infiltration of mononuclear cells into the parenchyma (Itard, 1989; Damayanti, 1993; Ngeranwa et al., 1993; Audu et al., 1999). A non-suppurative myocarditis, sometimes associated with hydropericarditis, has been reported (Tippit, 1978; Itard, 1989; Damayanti, 1993), which is accompanied by degeneration and necrosis of the myocardial tissue. Other lesions of the disease include glomerulonephritis (Nagle et al., 1982), renal tubular necrosis (Audu et al., 1999), non-suppurative meningo-encephalomyelitis (Ikede, 1983), focal poliomalacia and cerebromalacia (Ikede, 1983; Itard, 1989), keratitis, conjunctivitis, ophthalmitis (Losos, 1986), orchitis (Anosa and Isoun, 1980), interstitial pneumonia (Damayanti, 1993), bronchopneumonia (Ngeranwa et al., 1993), degenerative changes in the thyroid gland (Mutayoba et al., 1988), adrenal gland (Mutayoba et al., 1988; Ogwu et al., 1992) and pituitary gland (Ikede and Losos, 1975), and atrophy of the bone marrow in the chronic stage. Hypertrophy of the spleen and lymph nodes with the formation of germinal centres, and proliferation of plasma cells and macrophages occur at the acute stage, but the lymphoid tissues become exhausted and fibrotic in the chronic stage (Ikede and Losos, 1972b).

Diagnosis

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Due to the absence of pathognomonic signs, clinical examination is of little help in the diagnosis of animal trypanosomosis (Losos, 1986). Definitive diagnosis of the disease is based on the identification of the trypanosomes in the blood samples of animals, using parasitological methods (Itard, 1989). Microscopic examination of fresh blood samples may be carried out on wet-blood films. The parasites are identified by their motility in the wet films, examined using a clear-background or phase-contrast illumination. Preparation and examination of thin and thick blood films stained by Giemsa, May-Grunwald-Giemsa, Stevenel blue or methylene blue staining techniques, have been described (Itard, 1989). Specific diagnosis of trypanosome species is possible on stained blood films using trypanosome morphological characteristics (Itard, 1989).

In cases of low parasitaemia, infection may not be detected by wet, thin and thick film methods. Therefore, the techniques of concentrating trypanosomes by centrifugation of a whole blood sample or separation of the trypanosomes by filtration or erythocyte lysis are adopted to improve the efficiency of diagnosis (Baker, 1970; Itard, 1989). The blood is concentrated in heparinized capillary tubes after centrifugation in the haematocrit centrifugation technique (HCT; Woo, 1971) and the buffy coat method (BCM; Murray et al., 1977). After centrifugation, the capillary tube in HCT is mounted on glass sides with immersion oil covering the buffy coat-plasma junction, which is examined for trypanosomes. The capillary tubes in BCM are cut with a diamond pencil at the buffy coat-plasma junction and the content at the junction is smeared on a glass side and is examined as a wet film.

The miniature anion-exchange centrifugation method (mAEC; Lumsden et al., 1977, 1979) separates the trypanosomes from the erythrocytes on a DEAE column, the elute containing trypanosomes is centrifuged and the sediment examined by wet film method. The silicone centrifugation method (SCM; Ogbunude and Magaji, 1982) separates the trypanosomes after layering of the blood on silicone fluid. The erythrocyte lysis centrifugation method (LCM; Itard, 1989) involves the lysis of erythrocytes by mixing heparinized blood with twice the volume of distilled water, after which the fluid’s osmolarity is balanced with a hypertonic solution, centrifuged and the sediment examined by wet film method.

The wet, thin and thick film methods were considered poor diagnostic tools for animal trypanosomosis (Kalu and Agu, 1984), when compared with the concentration methods (Kalu et al., 1986). The HCT was less sensitive than the BCM (Paris et al., 1982; Kalu et al., 1986). The superiority of BCM over HCT and other concentration methods was enhanced by the phase-contrast facility (Murray et al., 1977; Paris et al., 1982). Also, BCM is a more rapid and practicable supplement to HCT (Kalu et al., 1986). The HCT has been reported to be less sensitive than the mAEC method (Lumsden et al., 1981).

Laboratory animal inoculation (Itard, 1989) using blood, lymph, oedema or tissue fluid, provides an excellent technique for detection of trypanosome species in the subgenus Trypanozoon (Losos, 1986), but is less sensitive with other trypanosome species (Kalu et al., 1986). The sample is inoculated subcutaneously or intraperitoneally into rats or mice for T. congolense, T. evansi and T. brucei, or intrascrotally into rabbits for T.equiperdum (Itard, 1989). The incubation period varies from 5 to 85 days. Mouse inoculation cannot be carried out routinely in the field, but it has been reported to be more sensitive than HCT and has a lower detection limit of three parasites per ml of blood for T. evansi infection (Le et al., 2000).

Seroimmunological techniques such as the mercuric chloride test, formol gel method, IgM titration by precipitation-diffusion, indirect fluorescent antibody test, enzyme-linked immunosorbent assay (ELISA), complement fixation test, agglutination test and passive haemagglutination test have been indicated as being applicable in the diagnosis of trypanosomosis (Itard, 1989). Among these techniques, only ELISA has been used in epidemiological surveys of trypanosomosis (Nantulya et al., 1992; Komoin-Oka et al., 1994; Singh et al., 1995; Delafosse et al., 1996; Desquesnes et al., 1999; Espinoza et al., 1999a, b).

The antigen ELISA was considered unsuitable for reliable detection of trypanosomal antigens (Rebeski et al., 1999a), because the trypanosome species specificity’s were poor (Eisler et al., 1998) and required improvement. The technique is less sensitive than BCM (Doko et al., 1996). Its sensitivity was less than 50% in primary trypanosome infections, but the sensitivity was high in repeated or chronic infections (Mattioli and Faye, 1996), aparasitaemic chronic infections (Kanwe et al., 1992; Trail et al., 1992) and mixed infections (Trail et al., 1992; Anosa et al., 1993). The estimated sensitivity of T. congolense antigen-based ELISA was greater than 96% (Rebeski et al., 2000). A sandwich-ELISA was highly sensitive for the detection of antigen in the blood of T evansi-infected cattle (Swarnkar et al., 1993). Antigen ELISAs are 4-5.5 times more sensitive than BCM in the diagnosis of sub-patent infections (ILRAD, 1992; Masake et al., 1995) and more sensitive and specific than antibody ELISAs in the diagnosis of latent T. evansi infection of buffaloes and horses (Singh et al., 1995). In a survey, an ELISA for trypanosome antibody detection gave a prevalence of 81.7%, whereas BCM was less sensitive with a prevalence of 15% (Desquesnes et al., 1999). An ELISA to detect antibodies against T. evansi appeared to be sensitive and specific (Tuntasuvan et al., 1996). Other antibody detection methods already in use, include direct card agglutination tests(CATT; Wuyt et al., 1995; Verloo et al., 2000) and indirect card agglutination tests (LATEX; Verloo et al., 2000), with specificity’s of 95% and 82% respectively. An indirect latex agglutination test (Suratex) has been developed for detecting circulating trypanosomal antigens in latent infections, which cannot be detected by parasitological techniques (Nantulya et al., 1998). Double immunodiffusion and counter-immunoelectrophoresis tests are not effective for detection of trypanosomal antigens (Swarnkar et al., 1993).

Techniques using polymerase chain reactions (PCR) have been widely used to screen infected animals, however, due to cost they are not used as diagnostic tools in the field. The parasitological (HCT, BCM) and PCR techniques have approximately the same sensitivities for detection of T. vivax infections in cattle (Desquesnes, 1997). However, some workers reported that PCR proved consistently more sensitive than parasitological techniques (Almeida et al., 1997), especially in antigenaemic but aparasitaemic animals (Majiwa et al., 1994). PCR amplifying the internal transcribed spacer (ITS1) region (Njiru et al., 2005) allows several pathogenic trypanosome species to be diagnosed in a single reaction (based on the size of the band produced). PCR tests have also been developed that can discriminate between T. b. brucei and T. b. rhodesiense (Picozzi et al., 2008). The need for improvement in the PCR technique has led to the consideration of dipstick assays and tests (Rebeski et al., 1999b), however, they have not been widely used. Loop-mediated isothermal amplification (LAMP) offers a simple way to amplify DNA using a waterbath and can amplify Trypanozoon (Kuboki et al. 2003; Njiru et al., 2008), T. congolense (Thekisoe et al., 2007), T. vivax (Njiru et al., 2011) and T. cruzi (Thekisoe et al., 2007, 2010) DNA.

Clinical signs

The clinical signs of animal trypanosomosis (nagana and surra) include intermittent fever, anorexia, ocular discharge, photophobia, pale mucous membranes, swelling of visible lymph nodes, oedema of the throat, underline and scrotum, dullness, lethargy, emaciation, reduction in milk yield and capacity to work in draught animals, male and female infertility, abortion, stillbirth, premature births and deaths.

Nervous signs may occur in infections involving T. brucei subspecies and T. evansi, and include ataxia, circling movement, incoordination, aimless running, staggering gait, head pressing and banging, paraplegia, paralysis and prostration (Radostits et al., 1994; Reddy and Hafeez, 1996; Manuel et al., 1998). T. simiae infected pigs may die within a few hours, presenting as the severest and quickest form of trypanosomosis (Losos, 1986), with signs of fever, increased respiration, dullness, stiff and unsteady gait and cutaneous hyperaemia.

The clinical signs of dourine (Radostits et al., 1994) are usually recognized during breeding, although the affected animal may not exhibit signs. In the primary stage, the genital signs of dourine in the male may include swelling of the penis, scrotum, prepuce and the surrounding skin, paraphimosis, mucopurulent urethral discharge and swelling of the inguinal lymph nodes. In the female, there may be sexual excitement, swelling of the vulva, vaginal discharge, hyperaemia and ulceration of the vaginal mucosa, oedema of the perineum, udder and abdominal floor. Cutaneous urticaria-like plaques known as ‘silver dollar spots’, which are pathognomonic, appear in the secondary stage. Anaemia, emaciation and nervous signs such as stiffness and weakness of limbs, inco-ordination, ataxia and paralysis occur in the tertiary stage, sometimes with atrophy of the hindquarters. The disease is mild and recurrent, but death occurs in severe forms.

List of Symptoms/Signs

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SignLife StagesType
Cardiovascular Signs / Jugular pulse Sign
Cardiovascular Signs / Peripheral venous distention, jugular distention Sign
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:All Stages,Sheep & Goats:All Stages Sign
Digestive Signs / Bloody stools, faeces, haematochezia Cattle & Buffaloes:All Stages Sign
Digestive Signs / Diarrhoea Cattle & Buffaloes:All Stages Sign
Digestive Signs / Excessive salivation, frothing at the mouth, ptyalism Sign
Digestive Signs / Excessive salivation, frothing at the mouth, ptyalism Sign
Digestive Signs / Excessive salivation, frothing at the mouth, ptyalism Sign
Digestive Signs / Grinding teeth, bruxism, odontoprisis Sign
Digestive Signs / Hepatosplenomegaly, splenomegaly, hepatomegaly Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:All Stages,Sheep & Goats:All Stages Diagnosis
Digestive Signs / Rumen hypomotility or atony, decreased rate, motility, strength Sign
General Signs / Ataxia, incoordination, staggering, falling Sign
General Signs / Ataxia, incoordination, staggering, falling Sign
General Signs / Dehydration Sign
General Signs / Dysmetria, hypermetria, hypometria Sign
General Signs / Exercise intolerance, tires easily Cattle & Buffaloes:Ox Sign
General Signs / Fever, pyrexia, hyperthermia Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:All Stages,Sheep & Goats:All Stages Diagnosis
General Signs / Generalized weakness, paresis, paralysis Sign
General Signs / Generalized weakness, paresis, paralysis Sign
General Signs / Head, face, ears, jaw, nose, nasal, swelling, mass Sign
General Signs / Hypothermia, low temperature Sign
General Signs / Hypothermia, low temperature Sign
General Signs / Icterus, jaundice Sign
General Signs / Inability to stand, downer, prostration Sign
General Signs / Inability to stand, downer, prostration Sign
General Signs / Lack of growth or weight gain, retarded, stunted growth Cattle & Buffaloes:Calf,Other:Juvenile,Pigs:Weaner,Pigs:Growing-finishing pig,Sheep & Goats:Lamb Diagnosis
General Signs / Lymphadenopathy, swelling, mass or enlarged lymph nodes Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:All Stages,Sheep & Goats:All Stages Diagnosis
General Signs / Opisthotonus Sign
General Signs / Opisthotonus Sign
General Signs / Pale mucous membranes or skin, anemia Sign
General Signs / Pale mucous membranes or skin, anemia Sign
General Signs / Paraparesis, weakness, paralysis both hind limbs Sign
General Signs / Paraparesis, weakness, paralysis both hind limbs Sign
General Signs / Petechiae or ecchymoses, bruises, ecchymosis Sign
General Signs / Reluctant to move, refusal to move Sign
General Signs / Sudden death, found dead Pigs:All Stages Diagnosis
General Signs / Sweating excessively, hyperhidrosis Sign
General Signs / Swelling mass penis, prepuce, testes, scrotum Other:Adult Female,Other:Adult Male Diagnosis
General Signs / Tetraparesis, weakness, paralysis all four limbs Sign
General Signs / Torticollis, twisted neck Sign
General Signs / Trembling, shivering, fasciculations, chilling Sign
General Signs / Trembling, shivering, fasciculations, chilling Sign
General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift Sign
General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift Sign
General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift Sign
General Signs / Weight loss Other:Adult Female,Other:Adult Male Diagnosis
Nervous Signs / Abnormal behavior, aggression, changing habits Sign
Nervous Signs / Circling Other:All Stages Diagnosis
Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:All Stages,Sheep & Goats:All Stages Diagnosis
Nervous Signs / Excitement, delirium, mania Cattle & Buffaloes:All Stages,Other:All Stages,Sheep & Goats:All Stages Sign
Nervous Signs / Head pressing Other:All Stages Diagnosis
Nervous Signs / Propulsion, aimless wandering Other:All Stages Diagnosis
Nervous Signs / Seizures or syncope, convulsions, fits, collapse Sign
Nervous Signs / Tremor Sign
Ophthalmology Signs / Blindness Sign
Ophthalmology Signs / Chemosis, conjunctival, scleral edema, swelling Sign
Ophthalmology Signs / Chemosis, conjunctival, scleral edema, swelling Sign
Ophthalmology Signs / Conjunctival, scleral, injection, abnormal vasculature Sign
Ophthalmology Signs / Conjunctival, scleral, redness Sign
Ophthalmology Signs / Corneal edema, opacity Sign
Ophthalmology Signs / Lacrimation, tearing, serous ocular discharge, watery eyes Sign
Ophthalmology Signs / Miosis, meiosis, constricted pupil Sign
Ophthalmology Signs / Photophobia Sign
Ophthalmology Signs / Strabismus Sign
Reproductive Signs / Abnormal length of estrus period, heat Sign
Reproductive Signs / Abnormal size testes / scrotum Sign
Reproductive Signs / Abortion or weak newborns, stillbirth Cattle & Buffaloes:Cow,Other:Adult Female,Pigs:Sow,Sheep & Goats:Mature female Diagnosis
Reproductive Signs / Agalactia, decreased, absent milk production Sign
Reproductive Signs / Agalactia, decreased, absent milk production Sign
Reproductive Signs / Anestrus, absence of reproductive cycle, no visible estrus Cattle & Buffaloes:Cow,Other:Adult Female,Pigs:Gilt,Pigs:Sow,Sheep & Goats:Gimmer,Sheep & Goats:Mature female Diagnosis
Reproductive Signs / Female infertility, repeat breeder Cattle & Buffaloes:Cow,Other:Adult Female,Pigs:Sow,Sheep & Goats:Mature female Diagnosis
Reproductive Signs / Foul smelling discharge, vulvar, vaginal Other:Adult Female Sign
Reproductive Signs / Male infertility Cattle & Buffaloes:Bull,Other:Adult Male,Pigs:Boar,Sheep & Goats:Breeding male Diagnosis
Reproductive Signs / Mucous discharge, vulvar, vaginal Other:Adult Female Sign
Reproductive Signs / Paraphimosis or priapism, inability to retract penis Other:Adult Male Sign
Reproductive Signs / Purulent discharge, vulvar, vaginal Other:Adult Female Sign
Respiratory Signs / Coughing, coughs Sign
Respiratory Signs / Coughing, coughs Sign
Respiratory Signs / Dyspnea, difficult, open mouth breathing, grunt, gasping Sign
Respiratory Signs / Dyspnea, difficult, open mouth breathing, grunt, gasping Sign
Respiratory Signs / Dyspnea, difficult, open mouth breathing, grunt, gasping Sign
Respiratory Signs / Epistaxis, nosebleed, nasal haemorrhage, bleeding Sign
Respiratory Signs / Increased respiratory rate, polypnea, tachypnea, hyperpnea Pigs:All Stages Sign
Respiratory Signs / Mucoid nasal discharge, serous, watery Sign
Respiratory Signs / Mucoid nasal discharge, serous, watery Sign
Respiratory Signs / Mucoid nasal discharge, serous, watery Sign
Respiratory Signs / Purulent nasal discharge Sign
Respiratory Signs / Purulent nasal discharge Sign
Respiratory Signs / Purulent nasal discharge Sign
Skin / Integumentary Signs / Alopecia, thinning, shedding, easily epilated, loss of, hair Sign
Skin / Integumentary Signs / Pruritus, itching skin Sign
Skin / Integumentary Signs / Rough hair coat, dull, standing on end Sign
Skin / Integumentary Signs / Rough hair coat, dull, standing on end Sign
Skin / Integumentary Signs / Skin crusts, scabs Sign
Skin / Integumentary Signs / Skin edema Sign
Skin / Integumentary Signs / Skin edema Sign
Skin / Integumentary Signs / Skin erythema, inflammation, redness Pigs:All Stages Sign
Skin / Integumentary Signs / Skin necrosis, sloughing, gangrene Sign
Skin / Integumentary Signs / Skin plaque Sign
Skin / Integumentary Signs / Skin scales, flakes, peeling Sign
Skin / Integumentary Signs / Skin wheal, welt Sign
Skin / Integumentary Signs / Warm skin, hot, heat Pigs:All Stages Sign
Urinary Signs / Proteinuria, protein in urine Cattle & Buffaloes:All Stages,Other:All Stages,Pigs:All Stages,Sheep & Goats:All Stages Sign

Disease Course

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Trypanosomosis has been divided into acute and chronic phases (Anosa, 1988; Murray and Dexter, 1988). After deposition of the infective forms of trypanosomes in the subcutaneous tissues of the host by the vector, the trypanosomes multiply at the site of infection to produce a chancre and invade the blood stream either directly or through the lymphatic system (Losos, 1986). The acute phase begins with a rise in parasitaemia and a decline in packed cell volume. The peaks of the parasitaemic waves are usually high within this acute period and the degree of anaemia is closely correlated with the intensity and duration of the parasitaemia (Dargie et al., 1979a; Murray and Dexter, 1988). The animals that progress into the chronic phase usually have lower parasitaemia and less severe anaemia than those that are in the acute phase (Anosa, 1988). The chronic phase is characterized by low transient parasitaemia or aparasitaemia and the infected animals maintain low packed cell volume. Death may occur at any phase of the disease or the animal may recover naturally (Uzoigwe, 1986).

Pathogenesis

Trypanosome infections initiate pathological mechanisms that cause cellular injury resulting in anaemia, leucopenia, thrombocytopenia, tissue inflammation and tissue damage. Many factors and mechanisms act in unison and it is difficult to identify the primary and secondary factors and also the sequence of their roles in the pathogenesis of the disease (Igbokwe, 1994).

The anaemia in the disease is mainly caused by extravascular haemolysis through erythrophagocytosis in the mononuclear phagocytic systems of the spleen, liver, lungs and haemal nodes (Anosa and Kaneko, 1983; Anosa, 1988). Some minimal degree of intravascular haemolysis occurs in the early acute phase (Esievo et al., 1984). Erythropoietic response to the anaemia is inadequate (Igbokwe and Anosa, 1989; Igbokwe and Mohammed, 1991) in the acute and chronic phases. This is due to phagocytosis of erythroid cells (Anosa and Kaneko 1983c 1984; Anosa et al., 1992) and depressed erthropoietin stimulation (Igbokwe and Anosa, 1989; Igbokwe, 1989; Igbokwe 1997; Suliman et al., 1999). In the chronic phase there is probably also decreased haemoglobin synthesis (Igbokwe, 1989). The contributions of haemodilution and haemorrhages in the cause of the anaemia observed are uncertain (Murray and Dexter, 1988).

The leucopenia and thrombocytopenia are caused by mechanisms, which predispose the leucocytes and platelets to phagocytosis (Anosa et al., 1992). The leucocytes and platelets may be wasted at inflammatory sites and through microangiopathy (Igbokwe, 1994). Leucopoiesis seems to be impaired by phagocytosis of the myeloid cells (Anosa et al., 1992) and depression of the process by some unknown factors in the plasma (Kaaya et al., 1979, 1980).

Immunological mechanisms in the pathogenesis lead to extensive proliferation of activated macrophages, which engulf and destroy erythrocytes, leucocytes, platelets and haematopoietic cells (Anosa et al., 1992).

Some immunological reactions in the disease cause the release of vasoactive and chemotactic substances from the activation of complement systems (Lambert and Houba, 1974; Nagle et al., 1974), and the Hageman factor (Boreham, 1978) from the aggregation and degranulation of platelets (Slot et al., 1977; Zwart and Veenedaal, 1978). These chemical mediators elicit tremendous inflammatory reactions, and large numbers of mononuclear cells and a few polymorphonuclear leucocytes infiltrate the infected tissues (Anosa, 1983a, b) in T. brucei infections. However, no neutrophils are present with the mononuclear cell infiltrate in T. vivax infections (Anosa and Isoun, 1980, 1983).

Living and dead trypanosomes produce a number of biologically active substances that cause cellular injury and damage in various ways (Igbokwe, 1994). These substances are proteases (ILRAD, 1986; Knowles et al., 1987), neuraminidase (Esievo, 1979, 1983), phospholipases (Tizard et al., 1978), pyruvate (Coleman, and Von Brand, 1957) and aromatic by-products (Seed and Hall, 1985). Trypanosomes may cause direct mechanical injury to erythrocytes and other cells by the lashing action of their powerful flagella and microtubule-reinforced bodies (Vickerman and Tetley, 1978), after adhering to the tissue (Bungener and Muller, 1976; Banks, 1979).

Fever and microangiopathy may lead to the fragmentation of erythrocytes (Murray and Dexter, 1988) when the erythrocytes, with temperature-induced reduction in plasticity, pass through the fibrin meshwork of microthrombi.

Metabolic changes in trypanosome-infected animals result in energy deficits and the catabolism of protein and fat (Igbokwe, 1995). In acute T. brucei infection of rats, glucose tolerance is progressively impaired as the duration of infection increases, implying that cellular glucose uptake is reduced (Igbokwe et al., 1998). The pathogenic implication of this finding is vital, since low energy intake predisposes T. brucei infected pigs to more severe disease (Fagbemi et al., 1990).

Erythrocytes of T. brucei-infected animals seem to be more susceptible to free-radical-mediated damage due to exposure to hydrogen peroxide (Ameh, 1984; Igbokwe et al., 1994). This observation suggested that the erythrocytes of the infected animals had a reduced antioxidant capacity. Administration of vitamins C and E reduced the severity of anaemia and liver damage in T.brucei-infected animals (Umar et al., 1999, 2000), providing indirect evidence that oxidative injury may contribute in the pathogenesis.

Epidemiology

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Trypanosome related facts

The level of parasitaemia in an infection is related to the virulence of the trypanosome, which is a factor controlled by the capacity of the parasite to multiply and its ability to induce an immune response (Roelants and Pinder, 1987; Murray and Dexter, 1988). The virulence of trypanosomes depends on the species and strain of the trypanosome and the susceptibility of the host involved. Ruminants and equines are susceptible to T. vivax, T. congolense, T. brucei and T. evansi. T. simiae and T. suis are very pathogenic in pigs, but are mildly pathogenic in other animals. Although carnivores are susceptible to T. brucei and T. congolense, they are resistant to T. vivax (Itard, 1989). The strains of T. vivax in West Africa are more pathogenic than the ones in East Africa (Radostits et al., 1994), except for the strain of T. vivax in East Africa that causes acute haemorrhagic disease, which is very pathogenic (Williams et al., 1992). Osaer et al. (1994), showed that a West African strain of T. congolense was more pathogenic than an East African strain. The trypanotolerant N’dama cattle suffered more adverse effects from T. congolense than T. vivax infection (Mattioli et al., 1999). Infections in animals caused by more than one species of trypanosomes at a time have been reported in some studies (Turkson, 1993; Omeke, 1994; Bossche and Mudenge, 1997), and this often lead to a more severe disease (Radostits et al., 1994). Although drug-resistant trypanosomes may have altered virulence, which is not confirmed (Geerts and Holmes, 1998), their presence affects the epidemiology of the disease in areas where control is based on drug treatment (Rowlands et al., 1993). Since the life cycle of T. vivax is short, it is more readily transmitted than other species and mechanical transmission of T. vivax allows it to spread outside the tsetse belt (Radostits et al., 1994).

Impact: Economic

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The total cost of an outbreak of T. vivax infections in the Brazilian Pantanal wetland, on affected ranches, in 1995 was estimated to be US $32,631 after treatment, but the estimated losses would have exceeded US $140,000 if the animals had not been treated (Seidl et al., 1999). In West Africa, the resumption of milk production after trypanosomosis control in dairy farms led to the sale of dairy products which generated an income of $3 per day for the Fulani women (Bauer et al., 1999). An estimated annual loss of US $3 billion in meat and milk production in Africa has been reported, as a result of infections (Budd, 1999). The potential benefits of improved trypanosomosis control in terms of meat and milk productivity alone, is US $700 million per year in Africa; and an estimated current cost of the disease to livestock farmers and consumers is put at US $1340 million annually (Kristjanson et al., 1999). To control animal trypanosomosis through the use of trypanocidal drug treatment, the estimated cost of the approximately 35 million doses in use each year has been estimated to be in excess of US $30 million (Hursey, 2001). There is no available financial estimate of the cost of antivectorial control in farming communities. Kristjanson et al., (1999) has estimated the present value of vaccine research for trypanosomosis control to be at least US $288 million with a benefit/cost ratio of 34:1.

Disease Treatment

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The common drugs used to treat trypanosome infections in animals are diminazene aceturate (Berenil), homidium bromide (ethidium) and chloride (novidium), isometamidium (Samorin or Trypamidium), pyrithidium bromide (prothridium), quinapyramine dimethylsulphate (Antrycide sulphate) and sulphate (Antrycide), combined quinapyramine sulphate and chloride (antrycide prosalt), suramin (naganol) and antrycide-suramin complex (Loso, 1986; Itard 1989; Radostits et al., 1994; Onyeyili and Egwu, 1995).

Diminazene aceturate has remarkable curative properties and is the commonest trypanocide currently used against T. congolense and T.vivax at 3-5 mg/kg and against T. brucei at 5-7 mg/kg in ruminants and horses. Doses of 10mg/kg have been used against T. evansi in cattle. Subcutaneous injections (as 7% solution) lead to slight local reactions, but very rapid absorption is possible with intramuscular injections. The drug is not well tolerated in horses; therefore, deep intramuscular injections at several sites followed by massaging, is recommended since local reactions are relatively frequent. The drug is contra-indicated in camels and dogs, because it can cause damage to the blood vessels, brain, liver and kidneys (Leach, 1961; Abdel-Latif, 1963; Losos and Crockett, 1969; Schmidt et al., 1978). Diminazene aceturate is combined with an antipyretic agent, phenyldimethyl pyrazolon (phenazone) and a synergist, procaine hydrochloride, in a drug formulation known as Trypan (Bourdichon et al., 1998). A single injection (15ml) of the preparation can protect cattle against re-infection with trypanosomes for a period of 3 months.

Homidium bromide and chloride are curative against T. congolense and T. vivax in cattle, sheep, goats and horses at 1 mg/kg. It is injected subcutaneously in the dewlap as a 1% or 2.5% solution to avoid irritation. Intramuscular injections, even if they are deep, should be avoided because of irritation. Its prophylactic effect lasts up to 6 weeks.

Isometamidium can be used as a curative and prophylactic drug in ruminants, horses, donkeys and dogs. It is administered by deep intramuscular injection at a dose of 0.25-1 mg/kg against T. vivax, T. congolense, T. brucei and T. evansi. The dose should be divided over several sites to avoid local reactions. Subcutaneous injection should be avoided because of violent reactions. For prophylaxis, higher doses may be used and protection may last for several months depending on the infection risk. Doses of 12.5-35 mg/kg have been used against T. simiae in pigs with a risk of acute cardiovascular collapse.

Pyrithidium bromide is administered by intramuscular injection as a 2.5% solution at the doses of 2-2.5 mg/kg to ruminants, donkeys and horses as a curative agent and for a minimum prophylaxis of 4 months against T. vivax and T. congolense. It has been used to treat T. evansi infections in dogs and donkeys.

Quinapyramine salts have been used successfully to treat T. congolense, T. vivax, T. brucei, T. evansi and T. equiperdum infections. Quinapyramine sulphate (Antrycide, Trypacide, Tribexin, Noroquin) use has been banned in livestock. It is only used to treat T. brucei infection in horses at 5 mg/kg. Quinapyramine methylsulphate is used against T. evansi and T. equiperdum infections in camels and horses at 3-5 mg/kg subcutaneously and against T. brucei infections in dogs at 5 mg/kg subcutaneously. Quinapyramine prosalt has been used against T. evansi in horses and camels for prophylaxis lasting 2-3 months at 1.1mg/kg subcutaneously. Systemic reactions may occur in horses and dogs involving signs such as salivation, dyspnoea, profuse sweating, colics and collapse.

Suramin is not very effective against T. vivax and T. congolense infections in cattle. Suramin is used against T. evansi and T. brucei infections in camels and horses, at 7-10 mg/kg, administered intramuscularly or intravenously. Systemic reactions may occur after intravenous injection (oedema, urticaria and laminitis in horses) or local inflammatory reaction and necrosis after intramuscular reaction. Prolonged treatment with Suramin causes chronic nephritis. Suramin-antrycide complex is used at 40 mg/kg against T. simiae in pigs.

Prevention and Control

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The elimination of trypanosome infections in animals by the use of trypanocidal drugs is an important control measure (Bealby et al., 1996) for ensuring good returns to the herd owners (Itty et al., 1995). About 35 million doses per year of these trypanocidal drugs, such as isometamidium chloride, homidium salts and diminazene aceturate, are currently used in Africa, but resistance of trypanosomes to these drugs has been reported in at least 14 countries in sub-Saharan Africa (Geerts and Holmes, 1998). The trypanosomes develop drug resistance as a result of underdosing (Boyt, 1986; Peregrine et al., 1997), increased treatment frequency and systematic mass treatment of animals (Geerts and Holmes, 1998). Avoidance of exposure of trypanosomes to sub-therapeutic drug concentrations is achievable through farmer education on correct dosing and the use of improved formulations of drugs, which use controlled release devices to provide more stable drug concentrations (Geerts and Holmes, 1998). The prophylactic effect of isometamidium was enhanced by providing it with a poly (D, L-Lactide)- sustained release device (Diarra et al., 1998). The use of the banned trypanocide, quinapyramine could lead to cross-resistance to the other trypanocidal drugs (Hawking, 1963) and therefore the ban should be continually enforced (Geerts and Holmes, 1998).

To enhance the effective use of trypanocidal drugs, they may be used as ‘sanative pairs’ (isometamidium or ethidium and diminazene), and treatment restricted to individual clinical cases (Geerts and Holmes, 1998). Good nutrition and absence of intercurrent disease help to boost immune response, which may reduce the rate of development of drug resistance and enhance recovery after drug treatment.

Because of the problems of drug resistance and high tsetse challenge in some areas, the control of trypanosomosis should not rely solely on the use of trypanocidal drugs. An integrated approach is adopted using vector control to break the transmission cycle and reduce the need for frequent treatment due to re-infection. Vector control involves ground and aerial spraying, pyrethroid dips, sprays or pour-on, impregnated traps and targets and release of sterile male tsetse flies (Allsopp, 1992; Holmes et al., 1997; Hargrove et al., 2000). Deltamethrin has been used for animal dipping in Tanzania (Fox et al., 1993) and Zambia (Luguru et al., 1993). It has been used as a pour-on in Burkina Faso (Bauer et al., 1995, 1999), Kenya (Baylis and Stevenson, 1998), Zambia (Bossche and Duchateau, 1998) and along the Uganda-Kenya border (Magona et al., 1998, 2000) and in impregnated targets. Cypermethrin has been used as a pour-on in Ethiopia (Leak et al., 1996; Swallow et al., 1995) and Kenya (Kamau et al.,2000) and in impregnated traps in Cote d’lvoire (Coulibaly et al., 1993; Rowlands et al., 1996a, b) and Burkina Faso (Amsler-Delafosse et al., 1995). In these reports, tsetse density and the prevalence of trypanosomosis were reduced by the control campaigns. In tsetse controlled areas, the prevalence of trypanosomosis was zero in some locations in Zambia (Belot and Leroy, 1998), and the health and productivity of animals consequently improved in Ethiopia (Jemal and Hugh-Jones, 1995). The use of restricted application where deltamethrin is applied to the legs and belly of cattle cuts insecticide usage by 40%, improves efficacy by 27% and reduces the impact on non-target organisms and on endemic stability of tickborne diseases (Torr et al., 2007). The sequential aerosol technique was used successfully to eliminate tsetse from the Okavango Delta in Botswana (Kgori et al., 2006).

The use of trypanotolerant livestock is part of a comprehensive programme of trypanosomosis control in parts of Africa (Holmes et al., 1997). In the subequatorial savannahs and equatorial forest-savannah mosaics of Africa, where tsetse infestation is heavy, only trypanotolerant breeds can survive without regular antivectorial measures and drug treatments (Itard, 1989). These tolerant animals are able to maintain apparent good health and productivity even when trypanosome infections exist (Maillard et al., 1992; Osaer et al., 1994; Achukwi et al., 1997; Mattioli et al., 1999), because they control the development of the parasites and limit their pathological effects (D’leteren et al., 1998).

A vaccine prepared from the flagellar pocket antigen derived from T. b. rhodesiense, using bovine serum albumin or ovalbumin as a carrier and alum as the adjuvant, has been used for the immunoprotection of cattle against trypanosomosis (Mkunza et al., 1995). In three field trials, immunized animals were exposed to the risk of trypanosome infections and 0.9-26% were infected while 13-58% of the un-immunized and 43% of samorin-treated controls were infected (Mkunza et al., 1995). This shows a good promise for vaccine development as a control strategy against trypanosomosis despite the limitations of antigenic variation associated with the trypanosomes.

Vector control plays an important role in the control of Chagas disease. The Southern Cone Initiative started in 1991 and initially involved six countries (Argentina, Bolivia, Brazil, Chile, Paraguay and Uruguay). Its goal was to interrupt transmission through treatment of houses with insecticides (Schofield and Dujardin, 1997). This has been successful in many parts of South America and the programme has been reviewed by Dias et al. (2002). A similar programme has also taken place in Central America to disrupt transmission (Hashimoto and Schofield, 2012).

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Links to Websites

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WebsiteURLComment
CFSPH: Animal Disease Informationhttp://www.cfsph.iastate.edu/DiseaseInfo/index.php"Animal Disease Information" provides links to various information sources, including fact sheets and images, on over 150 animal diseases of international significance.
HAT – WHO fact sheethttp://www.who.int/mediacentre/factsheets/fs259/en/
OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animalshttp://www.oie.int/en/international-standard-setting/terrestrial-manual/access-online/The Manual of Diagnostic Tests and Vaccines for Terrestrial Animals (Terrestrial Manual) aims to facilitate international trade in animals and animal products and to contribute to the improvement of animal health services world-wide. The principal target readership is laboratories carrying out veterinary diagnostic tests and surveillance, plus vaccine manufacturers and regulatory authorities in Member Countries. The objective is to provide internationally agreed diagnostic laboratory methods and requirements for the production and control of vaccines and other biological products.
OIE Technical Disease Cardshttp://www.oie.int/animal-health-in-the-world/technical-disease-cards/An updated compilation of 33 technical disease cards, containing summary information, mainly directed to a specialised scientific audience, including 32 OIE-listed priority diseases. USDA-APHIS (USA) are also credited with contributing to the maintenance of the cards.
Tetse and Trypanosomiasis Information 2007http://www.fao.org/docrep/010/i0036e/i0036e00.htm
Trypanosomiasis – CFSPH Fact sheethttp://www.cfsph.iastate.edu/Factsheets/pdfs/trypanosomiasis_african.pdf
Trypanosomiasis - Global Geo-spatial datasets FAOftp://ftp.fao.org/docrep/fao/012/i0809e/i0809e01.pdf
Trypanosomiasis – OIE Manualhttp://www.oie.int/fileadmin/Home/eng/Health_standards/tahm/2.04.18_TRYPANOSOMOSIS.pdf
USAHA: Foreign Animal Diseases. Seventh Editionhttp://www.aphis.usda.gov/emergency_response/downloads/nahems/fad.pdfCopyright © 2008 by United States Animal Health Association ALL RIGHTS RESERVED. Library of Congress Catalogue Number 2008900990 ISBN 978-0-9659583-4-9. Publication with 472pp. aimed at providing information for practitioners within the USA to prevent and or mitigate the incursion of foreign animal diseases into that country. Contains general chapters on surveillance, diagnosis, etc. as well as 48 chapters covering individual diseases, mostly those notifiable to the OIE.

Contributors

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12/03/15 Updated by:

Dr Ewan MacLeod, Division of Infection and Pathway Medicine, College of Medicine and Veterinary Medicine, University of Edinburgh, Chancellor's Building, 49 Little France Crescent, Edinburgh, EH16 4SB, UK

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