Atoxoplasma spp

From Bird
Life cycle of Isospora sp. Atoxoplasma sp. life cycle is similar except infection of mononuclear cells occurs in blood and tissues where the organism proliferates by binary fission (Gardiner CH, Fayer R, Dubey JP: An Atlas of Protozoan Parasites in Animal Tissues. Washington, DC: USDA/ARS, Agriculture Handbook # 651 p.31).
Schematic drawing of Atoxoplasma sp. oocyst containing two sporocysts with four sporozoites each (Gardiner CH, Fayer R, Dubey JP: An Atlas of Protozoan Parasites in Animal Tissues. Washington, DC: USDA/ARS, Agriculture Handbook # 651 p.20).
Atoxoplasma merozoites within macrophges and lymphocytes in a section of liver. H&E stain.
Atoxoplasma merozoites within the cytoplasm lymphocytes in a liver imprint. Scattered free nuclei from lysed hepatocytes also are present. Wright’s stain.
Atoxoplasma merozoites within the cytoplasm of lymphocytes in a blood smear. Wright’s stain

Atoxoplasmosis is a parasitic disease primarily of passerine birds, especially canaries, finches, sparrows, grosbeaks, starlings, and mynahs. It is caused by species of the coccidian protozoan Atoxoplasma, a host-specific parasite. Atoxoplasma serini, the species found in canaries, is not infectious to sparrows; likewise, the species infecting sparrows is non-infectious to canaries[1][2]. The parasite is transmitted via the fecal-oral route and not by mites as was formerly thought when it was known as Lankesterella. Parasitism may cause rapid and fatal disease in fledgling birds. This particular parasite is a special threat to successful species preservation of endangered Bali mynahs.

Much controversy has attended the study and classification of Atoxoplasma spp. This genus as been variously associated with Plasmodium, Haemoproteus, Toxoplasma spp in birds, as well as the amphibian parasite Lankesterella spp over the past hundred years[3]. Due to its morphologic similarity to Isospora spp it was most recently considered a stage of that parasite’s life cycle or a new species of Isospora altogether, called Isospora serini[4]. Although some researchers favor a return to the genus Isospora, Atoxoplasma spp. are currently differentiated from Isospora sp. according to the site of asexual amplification or merogony[5].

Life Cycle

Following ingestion of sporulated oocysts and release of sporocysts, the sporocysts of Isospora and Atoxoplasma invade intestinal epithelial cells. Isospora undergoes merogony in the intestinal epithelium, whereas Atoxoplasma enters the blood stream via the vasculature of the small intestine[6]. Atoxoplasma sporocysts subsequently invade mononuclear leukocytes and undergo asexual division in circulating and tissue lymphocytes, monocytes, and macrophages, as well as in intestinal epithelial cells[7].

The resulting merozoites form microgametes and macrogametes. Gametogony, the sexual stage of the coccidian life cycle, occurs in intestinal epithelial cells. Here the micro- and macrogametes combine to form zygotes, which then undergo multiple fission cycles to produce sporozoites within oocysts. These unsporulated oocysts are passed in the feces beginning nine to ten days post-infection and continue to pass for months, long after any clinical signs in the surviving birds have resolved. The oocysts sporulate in the environment and are then infective. Oocysts may be identified by fecal floatation, but atoxoplasma and isosporan oocysts may be difficult to distinguish. Oocysts of both organisms contain two sporocysts with four sporozoites each.

Clinical signs

Clinical signs of disease in fledglings are nonspecific and include diarrhea, anorexia, depression, ruffled feathers, ataxia, and hepatic enlargement that may be grossly visible through the living bird’s skin as "black spot"[8].

Mortality rates for atoxoplasmosis approach 80% in young passerines and the disease can devastate an aviary. Adult birds that are shedding oocysts often lack clinical signs, making elimination of the parasite from aviaries difficult. Patency of infection lasts for up to eight months due to the long life of macrophages in birds and the resultant long-lived pool of merozoites[9]. Oocysts are very stable in the environment and are not inactivated by most disinfectants. Thus, this parasite is especially virulent due to the damage to intestinal epithelium, liver, spleen, myocardium, and skeletal muscle in young birds. Atoxoplasma also is insidious due to its long life in persistently infected adult birds and its stability in the environment.

Diagnosis

Diagnosis of atoxoplasmosis has traditionally been via postmortem examination and histopathology of fledglings that die acutely or by fecal floatation on persistently infected adult birds. Identification of atoxoplasma oocysts is notoriously difficult because of structural similarity to those of Isospora spp and sporadic shedding of the organism by infected birds. Diagnosis can also be made on the live bird by peripheral blood smear examination and by cytologic examination of liver or spleen imprints or aspirates. Currently, research is underway to further evaluate less invasive, more sensitive, and more specific diagnostic tests for atoxoplasmosis in persistently infected adult birds as well as in sick fledglings[10].

At necropsy, infected birds have hepatomegaly and splenomegaly. Small, white foci are visible grossly on the liver and heart. The intestines may be distended and have translucent walls. Cytologic and histologic specimens reveal granulomatous to lymphohistiocytic inflammation of the heart, spleen, intestine, and liver. Macrophages may contain atoxoplasma merozoites. This form of the organism is round to oval, 3-5 m m diameter, cytoplasmic inclusion that causes indentation of the host cell nucleus, giving infected cells a characteristic appearance. Monocytes and lymphocytes containing merozoites may be seen within blood vessels on histologic sections. In the live patient, parasitized lymphocytes can be found in peripheral blood smears stained with a Romanowsky-type stain such as Wright’s, Giemsa, Leishman, or Diff-Quik stains[11]. Examination of buffy coat smears increases the likelihood of finding organisms. Like macrophages and lymphocytes in tissues, infected circulating cells display pink-staining intracytoplasmic merozoites causing an indented nucleus.

A recently developed polymerase chain reaction (PCR) test performed on feces, blood, or tissues of sick or persistently infected but asymptomatic birds is now available at The University of Georgia by special request.15 This test has proven very sensitive in initial clinical trials and should be useful in identifying asymptomatic, infected birds and determining the efficacy of treatment protocols.

Treatment

Effective prophylaxis of and treatment for atoxoplasmosis have not yet been perfected. Reduction of fecal-oral transmission can be achieved by cleaning cages frequently, using screen or hardware cloth cage bottoms to separate birds from infected droppings, and by frequently changing drinking and bathing water to minimize fecal contamination.

Primaquine is reported to suppress tissue forms of the parasite, and sulfachlor has been recommended to reduce oocyst shedding. Sulfonamides and amprolium, administered to adults before the breeding season and again when chicks are weaned, have also been suggested to reduce chick morbidity. However, none of these treatment protocols have been very effective. The most current recommendations to prevent and treat atoxoplasmosis in Bali mynahs are available on-line via the Species Survival Plan protocol[12].

Toltrazuril (Baycox), diclazuril, and sulfachlorpyrazine are under investigation for treatment of systemic atoxoplasmosis and to evaluate any reduction of oocyst shedding in Bali mynahs. Sulfachlorpyrazine inhibits the intestinal stages of the parasite life cycle and has been shown to reduce or clear oocyst shedding for as long as it is regularly administered. Toltrazuril may reduce mortality from systemic disease, although its efficacy does not appear to be as great as that of sulfachlorpyrazine. However, neither drug completely clears a bird of Atoxoplasma spp infection.

Diclazuril has been used in passerines other than Bali mynahs to treat toxoplasmosis; it may also prove effective against Atoxoplasma spp[13].

The Bali Mynah

Atoxoplasmosis presents a special threat in raising Bali mynahs in captivity. Efforts to raise Bali mynah chicks in captivity for later release in Bali Barat National Park have been seriously hampered by the widespread occurrence of Atoxoplasma spp in breeding programs and by high chick mortality associated with the disease. The free-ranging Bali Mynah is critically endangered in its native environment in Bali, due in part to poaching activity. Furthermore, it is unknown whether Atoxoplasma spp infects free-ranging mynahs[14]. Thus, it is critical to the survival of the Bali mynah that this parasite not be introduced into a naïve free-ranging population, where it could kill wild-bred chicks.

Since persistently infected birds can be completely asymptomatic, research into noninvasive, sensitive, and specific diagnostic tests for atoxoplasmosis has taken on a renewed urgency. The PCR test developed at The University of Georgia College of Veterinary Medicine may provide a reliable method to diagnose atoxoplasmosis in these infected birds. If so, this new diagnostic test will improve disease diagnosis, control, and prevention in captive breeding programs as well as prevent the accidental release of Atoxoplasma-infected birds into the wild.

References

  1. Box ED (1981) Isospora as an extraintestinal parasite of passerine birds. J Protozool 28:244-246
  2. Box ED (1970) Atoxoplasma associated with an isosporan oocyst in canaries. J Protozool 17:391-396
  3. Levine H, et al (1982) The genus Atoxoplasma (Protozoa, Apicomplexa). J Parasitol 68:719-723
  4. Box ED (1975) Exogenous stages of Isospora serini (Aragao) and Isospora canaria sp. in the canary (Serinus canarius linnaeus). J Protozool 22:165-169
  5. McNamee P, et al (1995) Clinical and pathological changes associated with Atoxoplasma in a captive bullfinch (Pyrrhula pyrrhula). Vet Rec 136:221-222
  6. Ball SJ, Brown MA, Daszak P, Pittilo RM (1998) Atoxoplasma (Apicomplexa: Eimeriorina: Atoxoplasmatidae) in the greenfinch (Carduelis chloris). J Parasitol 84:813-817
  7. Greiner EC, Ritchie BW (1994) Parasites. In Ritchie BW, Harrison GJ, Harrison LR (eds): Avian Medicine: Principles and Application. Lake Worth, FL: Wingers; pp:1007-1029
  8. Swayne DE, Getzy D, Slemons RD, Bocetti C, Kramer L (1991) Coccidiosis as a cause of transmural lymphocytic enteritis and mortality in captive Nashville warblers (Vermivora ruficapilla). J Wildl Dis 27:615-620
  9. Partington CJ (1989) Atoxoplasmosis in Bali mynahs. J Zoo Wildl Med 20:328-335
  10. MacWhorter, P.(1994) Passeriformes. In Ritchie BW, Harrison GJ, Harrison LR (eds): Avian Medicine: Principles and Application. Lake Worth, FL: Wingers; pp:1172-1199
  11. Little SE, Kelley LS, Norton TM, Terrell SP (2001) Developing diagnostic tools to further our understanding of Atoxoplasma species. Proc Assoc Avian Vet pp:157-159
  12. www.worldzoo.org/vetforum/baliatox.htm or www.riverbanks.org/aig/new.htm
  13. Quiroga MI, Aleman N, Vazquez S, Nieto JM (2000) Diagnosis of atoxoplasmosis in a canary (Serinus canaries) by histopathologic and ultrastructural examination. Avian Dis 44:465-469
  14. www.riverbanks.org/aig/new.htm