Before oocysts can be studied critically, they must be properly maintained to keep them viable so that their structural integrity remains intact. in our experience, oocysts from different vertebrate host species fall into two groups, which, of necessity, need to be handled differently when collected under field conditions.

These oocysts keep best when fresh feces are placed directly in 2-2.5% aqueous (w/v) potassium dichromate (K2Cr2O7) in a ratio of 1 volume of feces : greater than or equal to 5 volumes postassium dichromate. In field collections, either snap-cap or screw-cap 16-25 ml vials work well, but one should not fill the vial all the way to the top; leave a layer of air between the top of the feces-dichromate mixture and the cap to allow the oocysts some atmospheric oxygen. Unfortunately, other solutions for feces, for example, 2% aqueous sulfuric acid (see Wash et al., 1985) or common laboratory fixatives for oocysts (see Duszynski and Gardner, 1991) have proven unsatisfactory either for keeping oocysts viable or for preserving them as types.

These oocysts often are very thin-walled and fragile and sometimes prove difficult to sporulate. When examining hosts from freshwater environments, fresh mucus and feces from the intestinal tract should be placed in vials with tap water or with filtered river water at room temperature. Likewise, mucus and gut contents of marine animals should be placed in containers with filtered seawater. These fecal-water solutions must be supplemented with 200 IU penicillin g/ml, 200 ug streptomycin/ml, and 0.5 ug Fugizone/ml (see Upton et al., 1988; Molnár, 1996).

Upon return to the laboratory, the fecal-dichromate or fecal-water-antibiotic mixtures should be placed into a petri dish, any fecal pellets should be broken up, and the fecal material spread out in the dish and covered (Duszynski and Conder, 1977). The petri dishes generally should be maintained at room temperature (20-23 º C) for 7-10 days, which will allow any oocysts present to sporulate. Fecal-dichromate mixtures (terrestrial hosts) should not be refrigerated prior to the sporulation process as, in our experience, this will interfere with sporulation success. However, oocysts of some marine fishes were found to sporulate adequately only when the fecal-supplemented seawater mixture was placed on ice for 7-8 days (Upton et al., 1988); in this instance, the oocyst wall ruptured shortly after sporulation, releasing free sporocysts. In most species, however, after about 7-10 days, the mixture can be washed from the petri dish with clean potassium dichromate into a screw-cap jar (disposable baby bottle jars work well) filled only about half way and then put into a standard refrigerator (4-7 º C) until the material can be examined (sugar flotation) for the presence of oocysts. In our experience, oocysts of terrestrial vertebrates can remain viable, or at least stucturally intact, in the refrigerator for 3-4 years, whereas oocysts of certain fish coccidia (Upton et al., 1988, Molnár, 1996) may deteriorate soon after sporulation and die within a few days or weeks. Thus, it is probably best to study and document the structure of sporulated oocysts as soon as possible after they are sporulated.

Sporulated oocysts are best separated from the dichromate-fecal mixture by suspending an aliquot (1-3 ml) from the sample in 14-12 ml of modified Sheather's (Sheather, 1923) sugar flotation solution (500 g sucrose, 350 ml tap water, 5 ml phenol) via centrifugation (5 min at 2,000 rpm). It is important to use only number 1, 18 mm2 coverslips on top of the 15 ml centrifuge tubes (those with a smooth, beaded edge work best) as this reduces the surface area that needs to be scanned for oocysts. After centrifugation, lift the coverslip carefully from the centrifuge tube, place onto a glass slide, and set aside for 5-10 minutes; this allows the sugar along the edges of the coverslip to harden and minimizes movement of the oocysts during observation, measurement, and photography. The coverslip should be scanned systematically (100-400x total magnification) until oocysts are located. Measuring and detailing the structure of sporulated oocysts should always be done only under an oil immersion objective (Neofluor and Nomarski optics are both useful). Apochromatic lenses are superior to achromats and the higher the numerical aperture on the objective lens, the more accurate will be the measurements.


We strongly suggest that the following criteria be presented to allow accurate evaluation of a proposed new species description for coccidia (family Eimeriidae). In the list of features below, we have followed the example of Lom and Arthur (1989) by marking those features that are indispensable with a solid circle, while those recommended for inclusion are marked with an open circle.

The Host

The Sporulated Oocyst

Also see Duszynski and Wilber, 1997.

Literature Cited

Bandoni, S.M. and D.W. Duszynski. 1988. A plea for improved presentation of type material for coccidia. Journal of Parasitology 74: 519-523.

Box, E.D., A.A. Marchiondo, D.W. Duszynski, and C.P. Davis. 1980. Ultrastructure of Sarcocystsis sporocysts from passerine birds and opossums: Comments on classification of the genus Isospora. Journal of Parasitology 66: 68-74.

Couch, L., D.W. Duszynski and E. Nevo. 1993. Coccidia (Apicomplexa), genetic diversity, and environmental unpredictability of four chromosmal species of the subterranean superspecies Spalax ehrenbergi (mole-rat) in Israel. Journal of Parasitology 79: 181-189.

Duszynski, D.W. 1986. Host specificity in the coccidia of small mammals: Fact or fiction? In Advances in protozoological research. M. Bereczky (ed.). Symposia Biologica Hungarica, Vol. 33. Akademiai Kiado, Budapest, Hungary, p. 325-337.

Duszynski, D.W. and G.A Conder. 1977. External factors and self-regulating mechanisms which may influence the sporulation of oocysts of the rat coccidium, Eimeria nieschulzi. International Journal of Parasitology 7: 83-88.

Duszynski, D.W. and S.L. Gardner. 1991. Fixing coccidian oocysts is not an adequate solution to the problem of preserving protozoan type material. Journal of Parasitology 77: 52-57.

Frey, J.K., T.L. Yates, D.W. Duszynski, W.L. Gannon and S.L. Gardner. 1992. Designation and curatorial management of type host specimens (symbiotypes) for new parasite species. Journal of Parasitology 78: 930-932.

Lom, J. and J.R. Arthur. 1989. A guideline for the preparation of species descriptions in Myxosporea. Journal of Fish Diseases 12: 151-156.

McAllister, C.T. and S.J. Upton. 1989. The coccidia (Apicomplexa: Eimeriidae) of testudines, with descriptions of three new species. Canadian Journal of Zoology 67: 2459-2467.

Molnár, K. 1996. Eimerian infection in the gut of the tube-nosed goby Proterorhinus marmoratus (Pallas) of the River Danube. Systematic Parasitology 34: 43-48.

Parker, B.B. and D.W. Duszynski. 1986. Coccidiosis of sandhill cranes (Grus canadensis) wintering in New Mexico. Journal of Wildlife Diseases 22: 25-35.

Relman, D.A., T.M. Schmidt, A. Gajadhar, M. Sogin, J. Cross, K. Yoder, O. Sethabutr and P. Echeverria. 1996. Molecular phylogenetic analysis of Cyclospora, the human intestinal pathogen, suggests that it is closely related to Eimeria spp. The Journal of Infectious Diseases 173: 440-445.

Sheather, A.L. 1923. The detection of intestinal protozoa and mange parasites by a flotation technique. Journal of Comparative Pathology 36: 266-275.

Upton, S.J., S.L. Gardner and D.W. Duszynski. 1988. The round stingray, Urolophus halleri (Rajiformes: Dasyatidae), as a host for Eimeria chollaensis sp. nov. (Apicomplexa: Eimeriidae). Canadian Journal of Zoology 66: 2049-2052.

Wash, C.D., D.W. Duszynski and T.L. Yates. 1985. Eimerians from diffrent karyotypes of the Japanese wood mouse (Apodemus spp.), with descriptions of two new species and a redescription of Eimeria montgomeryae Lewis and Ball, 1983. Journal of Parasitology 71: 808-814.

Wilber, P.G., B. Hanelt, B. Van Horne and D.W. Duszynski. 1994. Two new species and temporal changes in the prevalence of eimerians in a free-living population of Townsend's ground squirrels Spermophilus townsendii) in Idaho. Journal of Parasitology 80: 251-259.

Wilber, P.G., K. McBee, D.J. Hafner and D.W. Duszynski. 1994. A new coccidian (Apicomplexa: Eimeriidae) in the northern pocket gopher (Thomomys talpoides) and a comparison of oocyst survival in hosts from radon-rich and radon-poor soils. Journal of Wildlife Diseases 30: 359-364.