584 J. PROTOZOOL., VOL. 3 1, NO. 4. NOVEMBER 1984

The most efficient method of isolating trypanosomes appears to be examination of residual blood fiom spleens (9) by single microhematocrit centrifbgation and subsequent centrifugation of the trypanosomes onto slides in a cytocentrifuge. In this way we demonstrated trypanosomes in each of the 15 spleens ex- amined.

LITERATURE CITED 1. Adams, A. R. D. & Lionnet, F. E. 1933. An outbreak of s u ~ a

among the wild d a r (Cervus unicolor var.) of Mauritius. J. Comp. Pathol. Ther., 46: 165-167.

2. Ayala, S. C., DAlessandro, A., Mackenzie, R. & Angel, D. 1973. Hemoparasite infections in 830 wild animals from the eastem llanos of Colombia. J. Parasitol.. 5 9 52-59.

3. Clark, G. G. 1972. Trypanosomes from mule dem in New Mex- ico and Colorado. J. Wildl. Dis., 8: 325-326.

4. DAlessandro, A. & Wells, E. A. 1971. Trypanosome infections in the family CeMdae. Tram. R. Sm. Trop. Med. Hyg.. 65: 845-846.

5. Deane, L. M. 1961. Tripanosomideos de mamiferos da re&o AmazBnica. I. A l g u n s flagelados encontrados no sangue de mamiferos silvestres do Estado do Para. Rev. Inst. Med. Trop. Srio Paulo. 3: 15- 28.

6. Herbert, J. V. 1961. Bovine trypanosomiasis due to Trypano- soma theileri Laveran, 1902 and its occurrence in Eire. Ir. Vet. J., 15: 230-236.

7. Hoare, C. A. 1972. The Trypanosomes ofMammals. Blackwell Scientific Publications, Oxford, UK.

8. Hoffmann, M. 1984. Trypanosomen bei Cerviden in Deutsch- land. Dr. Mad. Vet. Thesis, School of Veterinary Medicine, Hannover, west Germany.

9. Kingston, N. & Morton, J. K. 1975. Trypanosoma cewi s p a from elk (Cervus canadenrir) in Wyoming. J. Parasitol., 61: 17-23.

10. Kingston, N., Morton, J. K. & Dieterich, R. 1982. Trypano- soma cewi from Alaskan reindeer, Rangifer tarandus. J. Protozool., 29: 588-591.

1 I . Kingston, N., Thorne, E. T., Thomas, G. M., McHolland, L. & Trueblood, M. S. 1981. Further studies on trypanosomes in game animals in Wyoming 11. J. Wildl. Db., 17 539-546.

12. Kistner, T. P. & Hanson, W. L. 1969. Trypanosomiasis in white-tailed d a r . Bdl. Wildl. Dis. Assoc., 5: 398-399.

13. Kraneveld, F. C. & Mansjoer, M. 1952. Ondenoekingen over de gevoehgheid voor suna. 11. Het verloop der ziekte bij enkele in het wild levende dieren in Indonesie. Hemera Zoa, 59: 117-146.

14. Matthew, D. M., Kingston, N., Maki, L. & Nelms, G. 1979. Trypanosoma theileri Laveran, 1902, in Wyoming cattle. Am. J. Vet. Res.. 40: 623-629.

IS. Maza, S., Romaiia., C. & Fiora, A. 1932. Algunos hemopadt- sitos de mamiferos del No&. VII. R a n . Soc. Argent. Patol. Reg. Norie (B. Aires). Part 2: 990.

16. Petrich, J. 1976. Kultureller Nachweis von Trypanosoma thei- leri bei Rindern in Norddeutschland. Dr. Med. Vet. Thesis, School of Veterinary Mediche, Hannover, West Germany.

17. Saisawa, K, Take, K & Kaneko, K. 1933. Study on Trypano- soma theileri. Jpn. J. Exp. Med, 11: 101-105.

18. Siegel, S. 1956. Nonparametric Statistics for the Behavioral Sci- ences. McGraw-Hill Kogakusha Ltd., Tokyo.

19. Wells, E. A. 1969. Bovine trypanosomiasis with special refer- ence to Trypanosoma theileri. Ph.D. Thesis, University of Edinburgh.

Received 31 I 84; accepted 9 V 84

J Prorozwl.. 31(4), 1984. p p . 584-581 0 1984 by the Soany of RotozodogisU

Besnoitia darlingi (Apicomplexa, Sarcocystidae, Toxoplasmatinae): Transmission between Opossums and Cats'

D. D. SMITH d J. K. FRENKEL Department of Pathology and Oncology, University of Kama Medical Center, Kansas City, Kansas 66103

ABSTRACT. Opossums (Didelphrs marsupidis). act as intermediate hosts for Besnoitia darringi and could be infected orally with sporozoites (oocysts) and bradyzoites (tissue cysts), or intrapentoneally (i.p.) with tachyzoites. Infections could presumably be transmitted through cannibalism. Cats (Felis cutus), the definitive host, could be infected only with bradyzoites but not sporozoites. Oacysts shed by cats measure about 12 x 12 pm, resemble similarly sized oocysts of Toxoplarma gondii and Hammondia hammondi, and must be differentiated by the appearance of tissue cysts after experimental infection of intcrmcdiate hosts. Cats did not form tissue cysts of B. darlingi. Tachyzoites from the related B. jellisoni could be used in the Sabin-Feldman dye test to determine the development of antibody to B. darlingi in opossums after infection.

ESNOITIA darlingi (Brumpt, 1913) (4) Schneider, 1967 B (21-23) was described ftom the large, distinctive cyst con- taining lOOOs of zoites (Fig. I), recognized grossly in the con- nective tissue of Panamanian opossums (7, 16). In the U.S. it has also been found in opossums with nodules in the skin, tongue and adrenal glands (6,9). Recently, we described the cyclic trans- mission of cyst-bradyzoites to cats (Feh catus) and of oocyst- sporozoites to opossums (27), leading to the classification with the heteroxenous coccidia of the Toxoplasmatinae (1 0). To fur- ther define the cycle we now analyze the transmission of tachy- zoites, bradyzoites, and sporozoites to cats and opossums.

Supported by research grant AI-07489 from the National Institute of Allcrgy and Infedous Diseases. We thank Lynn Brann for rearing the opossums, Lilo Johnson for performing the serological tests, and the Missouri Department of Conservation and the Kansas Fish and Game Commission for permits to acquire and maintain opossums.

MATERIALS AND METHODS Seven opossums were obtained as nurslings from a road-killed

female from Jackson County, Missouri. They were hand-reared for use in these experiments. In addition, serum samples were obtained fiom 28 wild caught opossums obtained from the Kan- sas City area. Presence of infection was determined at necropsy by gross examination of ears, lips, tongue, skin, and the surface of viscera. In addition, tissues from most organs were fixed in formalin and examined in sections stained with hematoxylin and eosin. Laboratory mice that were chronically infected with B. darlingi and used as sources of infection for some of the experiments were CFl females obtained from Central Animal Research Facility, Inc., Shawnee, Kansas 66203.

Fecal collections were made from cats and some opossums at least two to three times weekly and floated in a sucrose so- lution (sp. gr. l. 15). Oocysts were stored in 2% (v/v) H,SO, or 2.5% (w/v) K,Cr,O,. Prior to inoculation, oocyst suspensions

SMITH & FRENKEL-BESNOITIA DARLINGI IN OPOSSUMS AND CATS 585

Fig. 1. Cyst of Besnoitia darlingi in intestinal smooth muscle of experimentally infected mouse. Cyst wall (CW) surrounds the host cell, showing hypertrophy and hyperplasia of nuclei 0. Within the host cytoplasm (Ha, a cytoplasmic vacuole (V) contains a large number of bradyzoites (B). H.&E. ~400. The bar equals 10 pm.

in H,SO, were neutralized with 3.3% (w/v) NaOH, using phenol red as an indicator. Dichromate suspensions were washed twice with normal saline prior to inoculation. Oocysts were admin- istered by means of a feeding tube.

For serological testing, opossums were bled from the heart while under ether anesthesia. Tachyzoites from the related B. jellisoni were used as antigen in the dye test because tachyzoites of B. darlingi were not available. These tachyzoites were ob- tained from a strain that was passed twice weekly in mice, as described by Frenkel et al. (1 1).

RESULTS Infections of Opossum (Table I)

Bradyzoites (tissue cysts). Three opossums were exposed to tissue cysts (containing bradyzoites) of B. darlingi. Two were fed mice infected with oocysts 48 to 238 days previously, and the third was injected intrapentoneally (i.p.) with triturated lung, liver, and spleen tissue from the 48-day infected mouse. These opossums were killed after 83 days and cysts were observed externally on ears, lips, tongue, and in the mouth, and internally on mesenteries, GI tract, kidneys, adrenals, and heart. Fecal specimens of these three opossums and of one infected with oocysts were examined for oocysts for 22 to 40 days after initial exposure to the inocula. None were ever observed to shed isos- poroid oocysts.

Sporozoites (oocysts). Oocysts were fed to two opossums. One opossum died after 14 days from bacterial endocarditis and pyelonephritis, similar to spontaneous infections that have been described (25). The other opossum was sacrificed at 124 days and numerous cysts were found in ears, lips, skin, nasal passages, and lungs, with fewer in heart, GI tract, liver, kidneys, adrenals, and mesenteries.

Tachyzoites. Peritoneal exudate was aspirated seven days postinfection from an opossum inoculated i.p. with infected

triturated tissues. This fluid was then injected i.p. into another opossum. When sacrifid at 92 days, numerous cysts and gran- ulomas were observed on ears, lips, tongue, mouth, liver, mes- enteries, and spleen of t h i s animal. Mice were similarly suscep tible although serial passage of tachyzoites was not achieved in mice. Two control opossums that remained uninoculated were examined after 200+ days; no tissue cysts were found.

Serologic response. Sera of all opossums were tested for an- tibody in the dye test using B. jellisoni antigen. Opossums fed bradyzoites attained a titer of 1:32; the one given tachyzoites i.p. developed a titer of 1:64, and the other three fed oocysts acquired a titer ofgreater than 1: 128. The highest titer observed

TABLE I. Besnoitia darlingi infictiom of opossums in relation to stage administered.

Stage administered Tachy- zoite

Sporo- (prolif- Bradyzoite zoite erative None (tissue cyst) (oocyst) stage) (controls)

~~

Number of opossums 3 2’ 1 2 Route PO/PO/IP Po IP -

1/1 1/1 0/2 Tissue cysts formed 313 oocyst shedding 013 0/2 NIY ND Antibody titers:

Pre-infection 4/<2/<2 4/4 <2 <2/<2 Post-infection 128/32/128 >128 64 8/8 * One opossum died at 14 days with bacterial endocarditis and pyelo-

nephritis. ND = not done.

586 J. PROTOZOOL., VOL. 31, NO. 4, NOVEMBER 1984

TABLE 11. Besnoitia darlingi infmtions of cats administered bradyzoites and sporozoites orally.

Stage administered Bradyzoite Sporozoite (tissuecyst) (oocyst)

Number of cats 14 2 Oocyst shedding 14/14 012 Oocyst shedding after challenge 3 4 2/2 Antibody titers:

Re-infection Post-infection

5 1:8 < 1:2 5l:8 (9) < 1:2 1:16 (3) 1:128 (2)

prior to infection or in controls was 1%. Thirteen of twenty- eight wild caught opossums from within a 50-mile radius of the Kansas City metropolitan area exhibited titers greater than 1:8 with the highest being 1 : 128.

Infections of Cats (Table II) Bradyzoites (tissue cysts). After the ingestion of mouse or

opossum tissues containing cysts, all of 14 cats shed oocysts of B. darlingi. Oocysts were identical to those described previously (27). Oocysts appeared after 9-14 days (2 = 1 1.5 days) and per- sisted from 3-1 3 days (2 = 8 days). Four cats that had previously shed oocysts were challenged with bradyzoites. Two failed to shed oocysts when challenged 66 and 69 days after initial in- fection. The other two cats challenged 66 and 76 days after initial infection did re-shed oocysts starting at 9 and 13 days and continued shedding for 7 and 14 days, respectively.

Development of tissue cysts in cats. Two cats that had previ- ously shed oocysts were sacrificed and portions of lung, heart, liver, spleen, pancreas, kidney, skeletal muscle, GI tract, and mesenteries were fed respectively to two cats. During 31 days of observation, neither of these cats discharged oocysts of B. darlingi. One of them was challenged with bradyzoites and later shed oocysts from day 13 through day 20.

Infection with sporozoites (oocysts). Two cats were fed spor- dated oocysts but neither discharged o a q s t s during 39 days of observation. An opossum fed inoculum oocysts from the same suspension seroconverted and developed cysts. Both cats were challenged with bradyzoites and both shed oocysts.

Serologic response. Of the 14 cats exposed to bradyzoites only two seroconverted strongly with titers at 1: 128. Seven other cats showed slight increases in titer, from one to three dilutions over their pre-exposure sample, to 1 : 16. Five cats showed no increase in titer. The two cats fed tissues from previously infected cats displayed postexposure titers of 1:8 and 1:16. Neither of the two cats exposed to oocysts of B. darlingi seroconverted in the dye test.

DISCUSSION From the experiments described, the life cycle of B. darlingi

may be characterized as heteroxenous, with oocysts developing only in cats and tissue cysts only in opossums and other inter- mediate hosts. Oocysts are infectious to the intermediate host, but not to the 6nal host. On the other hand, bradyzoites from tissue cysts are highly infectious to both intermediate and 6nal hosts; hence, cannibalism may be a factor in the transmission of this parasite between intermediate hosts.

When comparing the known life cycles of the other three congeners, B. jellisoni, B. wallacei. and B. besnoiti. the life cycle of B. darlingi most closely resembles that of B. jellisoni of ro-

TABLE 111. Key to distinguishing charactersitics of Sarcocystidae and genera of heteroxenour tirsue cyst-forming coccidia with isosporoid

oixysts.

la. Tissuc cyst contains a single meront --CLsloiraspoa Frmkel, 1977 2

2a. Cycle facultatively heteroxenous with oocyst capable of infecting the hnal host - Toxoplasmp gondii Nicolle & M a n m u , 1909*

3

b. lntracystic meronts dimorphic: metrocytes and bradyzoites - 5 4a. Host cell nucleus external to cyst, which is thin walled (< 1 cm);

cysts found mainly in striated muscle Hammondia Frenkel& Dubey, 197Sb

b. Host cell nuclei hypcmophicd and hyperplastic and enclosed by “cyst wall,” which is 1-10 pm thick; cyst found in many tis- sues Besnoitia Henry, 19 13”

5a. Cyst found principally in b& and spinal cord --- Frenkelia Biocca, 1968

6 6a. Cystsunjointed Surcocystis Lankester, 1 882d*

a r t h r o c y s t i s Levine, Beamer & Simon, 1970 Synonym: Isospora gondii Tadros & Laarman, 1976 (28). Synonym: Toxoplasma hammondi Levine, 1977 (1 4); Isospora da-

Synonym: Isospora jellisoni Tadms & Laarman, 1976 (28). Synonym: Endorimospora Tadros & Laarman, 1976 (28).

b. Tissue cyst is polyzoic

b. Oocpt not infectious to 6nal host 3a. Intracystic memnts are monomorphic: bradyzoites 4

b. Cyst found principally in striated muscle ___

b. Cysts jointed like bamboo

tusi Overdulve, 1978 (19).

Known to infect opossums.

dents (lo), at least in respect to the intermediate host (the de- finitive host of B. jellisoni has not yet been found), since infec- tions are readily transmitted between intermediate hosts via ingestion of infected tissue. Besnoitia wallacei may exception- ally be transmitted from one to another intermediate host ( 10). The degree of tachyzoite proliferation of B. darlingi is probably intermediate to that of B. jellisoni and B. wallacek B. besnoiti of cattle and sheep is apparently similar to B. darlingi in this relation and may be passaged in the tachyzoite stage from one intermediate host to another (1, 18). Domestic cats have been reported to be de6nitive hosts of B. besnoiti, B. wallacei. and B. darlingi (20, 27, 29).

The Sabin-Feldman dye test using B. jellisoni tachyzoites as antigen is a good indicator of B. darlingi infection in mice. In opossums antibody response is poor, but it was deemed ade- quate for diagnosis; however, in cats, it was an uncertain in- dicator of previous enteric infection.

Considering the ease with which B. darlingi may be trans- mitted from one intermediate host to another, individual hous- ing is advisable. Breeding colonies derived from wild caught animals in gang cages are most commonly used (8,24) (but see 12, 13). Cannibalism could result in infection of the entire col- ony. Indeed, judging from the fi-equency of tom ears and dam- aged tails in group-caged opossums, fighting could result in some transmission of this parasite. There are reports of the persistence of the related B. jellisoni in two laboratory colonies of Pero- myscus although inapparent transmission by a cryptic definitive host has not been excluded.

Although oocysts of Cystoisospora felis and C. rivolta can be recogILized by their larger sue, the similarity in size and ap- pearance of the smaller oocysts of B. darlingi, H. hammondi, and Toxoplawna gondii. also of cats, requires examination of tissue cysts in intermediate hosts for definitive diagnosis. Lab- oratory mice are fed or injected with sporulated oocysts, and after a period of at least 30 days, tissue cysts can be searched for. The possibility of double and triple infections must be kept in mind. Prophylaxis with sulfadiazine &om days 3 and 14 may be necessary to sustain mice through the acute phase ofinfection.

587 SMITH & FRENKEL-BESNOITIA DARLINGI IN OPOSSUMS AND CATS

The genera of cyst-forming coccidia are defined by their tissue cyst: its morphology and location, its Occurrence only in the intermediate or also in the final host, the infectivity of brady- zoites and sporozoites, and the degree of heteroxeny, whether obligatory or facultative (1 0, 26) (Table 111). More recently, a slightly different classification of genera has been proposed by Levine (1 5), who regards the genus Hammondia as a synonym of Toxoplasma and adds 5 taxa from amphibia and reptiles, the cycles of which are unknown. The genera Toxoplasma, Bes- noitia, and Hammondia have been synonymized with Isospora by Tadros & Laarman (28), and to avoid secondary homonymy with Isospora hammondi of the "marsh rat" (Oryzomys palus- tris), Overdulve (1 9) proposed I. datusi. Because we consider the cycle and the manner of transmission to be important, we continue to recognize these attributes in separate generic con- cepts (Table 111).

Two other cyst-forming coccidia, Toxoplasma and Sarcocys- tis, have been found in opossums as the intermediate host (1 6, 17, 30). The cyst of Toxoplasma is thin-walled, measuring less than 1 pm, and does not enclose the host cell nucleus. The cyst of Sarcocystis is located in skeletal muscles, the nuclei of which are unaltered in size. We have also observed opossums to serve as intermediate hosts for Cystoiospora rivolta.

Opossums may also act as definitive hosts for heteroxenous coccidia, shedding oocysts in their feces. Sarcocystis debonei form tissue cysts in grackles and blackbirds in the United States (2) and experimentally in a wide variety of birds from four families (3); S. columbididelphis infects doves of the genus Co- lumbina in Colombia, S.A. (5).

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2. Box, E. D. & Duszynski, D. W. 1980. Sarcocystis of passerine birds: sexual stages in the opossum (Didelphis virginiana). J. Wildl. Dis.,

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5. Carvajal, H. 198 1. Ciclo de vida de SarcoLystis columbididt'lphis n. sp. (Protozoa: Apicomplexa: Sarcocystidae) (Linneo 1758) entre la chucha Didelphis marsupialis y la torcaza c o m b Columbina tdpacoti (Temminick 18 1 1). Diss., University of Valle, Colombia, 37 pp.

6. Conti-Diaz, I. A., Turner, C. & Tweeddale, D. T. 1970. Bes- noitiasis in the opossum (Didelphis marsupidis). J. Paraitol.. 56: 457- 460.

7. Darling, S. T. 1910. Sarcosporidiosis in the opossum and its experimental production in the guinea pig by the intra-muscular injec- tion of sporozoites. Bull. Soc. Pathol. Exot.. 3: 5 13-5 18.

8. Farris, E. J. 1950. The opossum, in Farris, E. J., ed., The Care and Breeding of Laboratory Animals. New York, John Wiley and Sons,

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10. Frenkel, J. K. 1977. Besnoitia wallaceiofcats and rodents: with a reclassification of other cyst-forming isosporoid coccidia. J. Parasitol., 63: 61 1428.

16: 209-215.

pp. 256-266.

11. FrenkeL J. K-, D U ~ Y , J. P. & Hoff, R. L. 1976. LOSS of stages after cCdnuoW Passage Of Toxo~lasma gondii and Besnoitia jellisoni, J. Protozool.. 23: 421-424.

12. Jurgelski, w. 1974. The O p o s S ~ m (Didelphis virginiana Kern) as a biomedical model. I. Research perspective, husbandry, and l a b ratory technics. Lab. Animal Sci.. 24: 376403.

13. Jurgelski, W., Forsytbe, W., & Dahl, D. 1974. The opossum (Didelphis virginiana Kerr) as a biomedical model. 11. Breeding the opossum in captivity: facility design. Lab. Animal Sci., 24: 404-41 1.

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15. - 1982. Taxonomy and life cycles of coccidia, in Long, P. L., ed., The Biologyojthe Coccidia, University Park Press, Baltimore, pp. 1-33.

16. Mandour, A. M. 1965. Sarcocystis garnhami n. sp. in the skel- etal muscle of an opossum, Didelphis marsupialis. J. Protozool.. 12: 606-609.

17. McCulloch, W. F., Foster, B. G. & Diesch, S. L. 1967. Prev- alence of Toxoplama antibodies in wild mammals collected on three Iowa farms. Bull. Wildl, Dis. Assoc., 3: 30-31.

18. Neuman, M. 1965. Infection of laboratory white mice with Besnoitia besnoiti. Refu. Vet., 22: 201-205.

19. Overdulve, J. P. 1978. Prudish Parasites. The Discovery ofsex- uality and Studies on the Life Cycle, PURiCUhdy the Sexual Stages in Cats, of the Sporozoan Parasite Isospora (Toxoplasma) gondii. Elink- wijk, Utrecht, 83 pp.

20. Peteshev, V. M., Galuzo, I. G. & Polomoshnov, A. P. 1974. Cats-definitive hosts of Besnoitia (Besnoitia besnoiti]. IN. Akad. Nauk Az. SSR Ser. Biol., 1974: 33-38. (in Russian)

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22. - 1967a. Besnoitia darlingi (Brumpt, 19 13) in Panama. J. Protozool., 14 78-82.

23. - 1967b. The distribution of lizard besnoitiosis in Pan- ama, and its transfer to mice. J. Protozool., 14: 674-678.

24. Sherwood, B. F., Rowlands, D. T. & Hackel, D. B. 1969. The opossum (Didelphis virginiana) as a laboratory animal. Lab. Animal Care. 19: 494-499.

25. Sherwood, B. F., Rowlands, D. T. & Vakilzadeh, J. 1971. Ex- perimental bacterial endocarditis in the opossum (Didelphis virginiana). 111. Comparison of spontaneously occurring endacarditis with that in- duced experimentally by pyogenic bacteria and fungi. Am. J . Pathol.,

26. Smith, D. D. 1981. The Sarcocystidae: Sarcocystic, Frenkelia, Toxoplasma, Besnoitia, Hammondia and Cystoisospora. J. Protozool.,

27. Smith, D. D. & Frenkel, J. K. 1977. Besnoitia darlingi (Pro- tozoa: Toxoplasmatinae): cyclic transmission by cats. J. Parasitol.. 63: 1066-1071.

1976. Sarcocystis and related coccidian parasites: a brief general review, together with a discussion on some biological aspects of their life cycles and a new proposal for their classification. Acta L.ei&n., 44: 1-1 37.

29. Wallace, G. D. & Frenkel, J. K. 1975. Besnoitia sp. (Protozoa, Sporozoa, Toxoplasmatidae): recognition of cyclic transmission by cats. Science, 188: 369-37 1.

30. Walton, B. C. & Walls, K. W. 1964. Prevalence of toxoplas- mosis in wild animals from Fort Stewart, Georgia as indicated by se- rological tests and mouse inoculation. Am. J. Trop. Med. Hyg. .. 13: 530- 533.

64: 5 13-520.

28: 262-266.

28. Tadros, W. & Laarman, J. J.

Received 23 I 84; accepted 21 V 84