Trypanosoma evansi in Horses

Sarah Womack, DVM; Heather L. Tarpley, DVM; Susan E. Little, DVM, PhD; and Kenneth S. Latimer, DVM, PhD

Class of 2006 (Womack), Department of Parasitology (Little), and Department of Pathology (Tarpley, Latimer), College of Veterinary Medicine, University of Georgia, Athens, GA 30602-7388

Introduction

Trypanosma evansi is the causative agent of surra, one of the most common and widespread of the trypansomal diseases. This trypanosome can infect most mammals, although horses and camels are the principal hosts and represent the most significant sources of economic loss. Surra is endemic in many parts of Africa, Asia, and South America where thousands of animals die during disease outbreaks each year. Although not usually considered of zoonotic concern, one case of human infection with T. evansi recently has been documented in India.¹⁰

Life Cycle

T. evansi is a protozoal hemoflagellate that is transmitted mechanically by arthropod vectors. The most prominent arthropod vector varies geographically. For example, horseflies (Tabanus sp.) and stable flies (Stomoxys sp.) are the primary vectors in China and Indonesia, while tse tse flies are more prevalent in endemic areas of Africa. T. evansi has a limited survival time in the mouth parts of a potential vector, so increased time between blood meals decreases the risk of disease transmission. In South and Central America, vampire bats may serve as vectors and reservoirs for T. evansi.

Unlike other trypanosomes, procyclic or developmental stages have not been observed in any vector. In contrast, T. evansi reproduces by binary fission in the mammalian host. In addition to mechanical transmission, T. evansi infection may be spread during nursing, copulation, or following ingestion of infected tissues by carnivores. Equids and camelids are commonly infected with T. evansi, but other mammals such as buffalo, cattle, deer, dogs, and elephants also are susceptible. The severity of illness depends upon the protozoal strain, concurrent infections, stress, and environmental factors.^1,4

Figure 1. Generalized life cycle of a fly-transmitted trypanosome that reproduces by binary fission in the host (Reprinted from Gardiner CH, Fayer R, Dubey JP: An Atlas of Protozoan Parasites in Animal Tissues, Agriculture Handbook #651. Unites States Department of Agriculture, Washington, DC, 1988, p. 3).

Morphology and Biochemistry

T. evansi and T. equiperdum (the etiology of a venereally transmitted disease of horses called dourine), cannot be distinguished morphologically from one another, or from some forms T. brucei brucei.^1,2 The flagellate is long and slender, measuring 14 to 33 µm in length and 1.5 to 2.2 µm in width. The organism is generally monomorphic, although a shorter intermediate form occasionally is observed. This hemoflagellate has a free flagellum and a circular kinetoplast which is sometimes missing in mutated wild strains or strains that have been isolated following drug treatment. Although T. evansi and T. equiperdum are indistinguishable by light or electron microscopy, the clinical presentation of disease indicates which organism is most suspect.¹ Additionally, T. evansi is covered by a thick layer of a single glycoprotein (or variable surface antigen), that is the primary immunogen eliciting antibody formation. Periodically, the organism alters the glycoprotein coating and evades the host’s defensive responses.⁴

Clinical Signs

Clinical signs of disease may vary by individual or between species. Animals with acute T. evansi infection usually have a short clinical course of disease and die within weeks to months of infection. In contrast, some chronic infections of T. evansi may persist for years. In horses, donkeys, and mules, the incubation period of disease ranges from 5 to 60 days. Presenting signs may include weight loss, lethargy, anemia, progressive paresis, enlarged lymph nodes, petechiae of serous membranes, dependent edema, urticaria, and alopecia, exudation, or necrosis of the coronary bands. Recurrent episodes of pyrexia are associated with transient parasitemia.^1,4,6 Abortions and stillbirths may occur frequently. ⁸ Infection with T. evansi may not always cause overt signs of disease. Immunosuppression to vaccine antigens has been demonstrated in sheep and goats with subclinical T. evansi infections, as well as loss of draught capability in diseased horses.³

Diagnosis

Accurate diagnosis of surra is extremely important both in identifying animals for treatment as well as tracking the prevalence of disease. The differential diagnosis for surra includes include African horse sickness, equine viral arteritis, equine infectious anemia, and other forms of chronic parasitism.⁶ Demonstration of parasitemia, parasite antigen, or host antibodies to the parasite is necessary to obtain a definitive diagnosis. While it is not possible to distinguish between the trypansomes responsible for surra and dourine by light or electron microscopy, the diseases are very different clinically and T. equiperdum is rarely found in blood or tissues.¹

Several methods are used to directly demonstrate T. evansi. It is important to note that blood from major or deep vessels is more likely to yield a positive test result than blood obtained from peripheral sites. Once the sample is collected, either a wet blood film or a stained blood smear can be prepared. Lymph node aspirates may also be evaluated. Because the degree of parasitemia correlates with the probability of finding the organism, concentration techniques are useful to diagnose surra in animals with a low-grade parasite burden. Microhematocrit tube centrifugation and mini anion-exchange chromatography are two diagnostic methods. In addition, laboratory rodents may be inoculated with fresh biological samples from suspected cases in order to demonstrate trypanosomal infection.^1,4 The rodent inoculation test is very time consuming and is not very useful for field situations. In contrast, the more traditional diagnostic method of microscopic slide examination is easily performed in the field by may not always be sensitive. Thus, this latter diagnostic technique may be responsible for the underdiagnosis of surra.³

Figure 2. Romanowsky-stained blood smear from a horse with Trypanosoma evansi infection (surra). Notice the numerous, elongate trypanosomes (image courtesy of Dana Ambrose, MS, Athens Diagnostic Laboratory, University of Georgia, College of Veterinary Medicine, Athens, GA).

Indirect methods have been developed to identify T. evansi and increase the accuracy of disease diagnosis. These biochemical tests include flocculation, formol-gel, mercuric chloride precipitation, and thymol turbidity. Each of these tests relies on an increased serum globulin concentration in infected animals. These diagnostic tests have been used primarily in camels and have not demonstrated high specificity.⁴ Antigen detection tests, such as the latex agglutination test and enzyme-linked immunosorbent assay (ELISA), are available for diagnostic purposes. Detection of T. evansi antibodies during infection can be accomplished by ELISA, card agglutination test (CATT), complement fixation test (CFT), and indirect fluorescent antibody testing (IFAT). The polymerase chain reaction (PCR)also is being developed for diagnostic purposes.^4,7,9 Each of these tests requires further evaluation across species to establish which assay is the most sensitive, specific, and economical diagnostic tool.

Treatment and Control

Surra currently is on the list of notifiable diseases of the OIE (World Organization for Animal Health). Reduction of the global economic impact of this disease on animal health requires dependable diagnostic tests, efficacious treatment, and diligent control programs. Historically, three drugs have been used to animals with T. evansi infections. These drugs include diminazene, suramin, and quinapyramine. Cymelarsan is the newest drug to be introduced in the last decade. The choice of drug, dosage, and route of administration vary by species affected, local preference, and presence or absence trypanosome drug resistance.¹ In one study, diminazene cleared the parasitemia of horses following natural infection. However, parasitemia returned in many animals within 1 week of the first treatment with diminazene or within 24 hours of the second treatment with diminazene. This suggests that diminazene affords little protective effect against repeated T. evansi infection or the drug is ineffective in totally eliminating the initial T. evansi infection.⁸ Relapses have also been observed in horses following treatment with quinapyramine sulfate.⁵ In summary, there is a growing body of evidence that T. evansi is becoming resistant to currently available drugs and my may restrict their use or retard their effectiveness in the future. However, in vitro assays may help predict the effectiveness of drug treatment to given isolates of T. evansi.¹

In summary, the decision to treat an animal usually is based on the presence and severity of clinical signs of disease. Given the occasional low sensitivity of certain diagnostic tests, subclinical infections (carriers) may present a dilemma when attempting to control T. evansi infections in herd outbreaks.³

Other measures (besides trypanocidal drug administration) such as minimizing exposure to fly populations may help control surra. Stables with suitable netting can be used to exclude fly populations. Smudge fires may also be used to repel flies.³ Fortunately, T. evansi cannot survive very long outside of its host. Therefore, animal carcasses from surra fatalities pose little threat of disease spread. Finally, it is always prudent to restrict the movement of infected animals.⁶ Because surra is a foreign animal disease, suspected cases in the United States should be reported immediately to state and federal authorities.

References

1. Brun R, Hecker H, Lun ZR. Trypanosoma evansi and T. equiperdum: Distribution, biology, treatment and phylogenetic relationship (A review). Vet Parasitol 1998; 79:95-107.

2. Gilbert, RO. Dourine. Foreign Animal Diseases. Richmond, VA, United States Animal Health Association, 1998, pp. 182-188.

3. Luckins AG. Epidemiology of non-tsetse-transmitted trypanosomiasis - Trypanosoma evansi in perspective. Newsletter on Integrated Control of Pathogenic Trypanosomes and their Vectors. No.1, September 1999. http://www.icptv.org/Newsletters/Newsletter1/newsletter1.html

4. Luckins AG. Surra. OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. Ch. 2.5.15. http://www.oie.int/eng/normes/mmanual/A_00093.htm

5. Monzon CM, Mancebo OA, Russo AM. Antibody levels by indirect ELISA test in Trypanosoma evansi infected horses following treatment with quinapyramine sulphate. Vet Parasitol 2003;111:59-63.

6. Roth SA, August K, Spickler AR. Surra. Emerging and Exotic Diseases of Animals. http://www.cfsph.iastate.edu/Factsheets/pdfs/surra.pdf

7. Singh N, Pathak KM, Kumar R. A comparative evaluation of parasitological, serological and DNA amplification methods for diagnosis of natural Trypanosoma evansi infection in camels. Vet Parasitol 2004;126:365-373.

8. Tuntasuvan D, Jarabrum W, Viseshakul N, et al. Chemotherapy of surra in horses and mules with diminazene aceturate. Vet Parasitol 2003;110227-233.

9. Wernery U, Zachariah R, Mumford JA, Luckins T. Preliminary evaluation of diagnostic tests using horses experimentally infected with Trypanosoma evansi. Vet J 2001;161:287-300.

10. World Health Organization. A new form of human trypanosomiasis in India. Description of the first human case in the world caused by Trypanosoma evansi. Wkly Epidemiol Rec 2005;80:62-63.

Acknowledgement

Image of "Wild One" watercolor by Susan Weber is from the Artwork Section of Susan Weber Designs website and is used with permission.

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