Twenty blood samples of zebras ( Equus zebra zebra) from the Karoo National Park and the Bontebok National Park in South Africa, all seropositive for Theileria equi, were subjected to in vitro culture to identify carrier animals and to isolate the parasites. Sixteen animals had a detectable parasitaemia in Giemsa-stained blood smears examined before culture initiation, the remaining four animals were identified as T. equi carriers by in vitro culture. Cultures were initiated either in an oxygen-reduced gas mixture or in a 5% CO2-in-air atmosphere. Out of the 20 blood samples, 12 cultures of T. equi and two cultures of T. equi mixed with Babesia caballi were established. None of the four animals seropositive for B. caballi could be identified as carrier animals, whereas two seronegative samples became culture-positive for B. caballiThe articles have been scanned in colour with a HP Scanjet 5590; 600dpi. 
Adobe Acrobat v.9 was used to OCR the text and also for the merging and conversion to the final presentation PDF-format.mn201

Lopez-Rebollar, Laura M.

Meyer, P.

Zweygarth, Erich

Onderstepoort Journal of Veterinary Research

The articles have been scanned in colour with a HP Scanjet 5590; 600dpi. 
Adobe Acrobat v.9 was used to OCR the text and also for the merging and conversion to the final presentation PDF-format.Twenty blood samples of zebras ( Equus zebra zebra) from the Karoo National Park and the Bontebok National Park in South Africa, all seropositive for Theileria equi, were subjected to in vitro culture to identify carrier animals and to isolate the parasites. Sixteen animals had a detectable parasitaemia in Giemsa-stained blood smears examined before culture initiation, the remaining four animals were identified as T. equi carriers by in vitro culture. Cultures were initiated either in an oxygen-reduced gas mixture or in a 5% CO2-in-air atmosphere. Out of the 20 blood samples, 12 cultures of T. equi and two cultures of T. equi mixed with Babesia caballi were established. None of the four animals seropositive for B. caballi could be identified as carrier animals, whereas two seronegative samples became culture-positive for B. caballimn201

UPSpace at the University of Pretoria

Onderstepoort Journal of Veterinary Research, 69:197-200 (2002) In vitro isolation of equine piroplasms derived from Cape Mountain zebra (Equus zebra zebra) in South Africa E. ZWEYGARTH*1, LAURA M. LOPEZ-REBOLLAR1 and P. MEYER2 ABSTRACT ZWEYGARTH, E. , LOPEZ-REBOLLAR, L.M. & MEYER, P. 2002. In vitro isolation of equine piro-plasms derived from Cape Mountain zebra (Equus zebra zebra) in South Africa. Onderstepoort Journal of Veterinary Research, 69:197-200 Twenty blood samples of zebras (Equus zebra zebra) from the Karoo National Park and the Bonte-bok National Park in South Africa, all seropositive for Theileria equi, were subjected to in vitro cul-ture to identify carrier animals and to isolate the parasites. Sixteen animals had a detectable para-sitaemia in Giemsa-stained blood smears examined before culture initiation, the remaining four ani-mals were identified as T. equi carriers by in vitro culture. Cultures were initiated either in an oxygen-reduced gas mixture or in a 5% CO2-in-air atmosphere. Out of the 20 blood samples, 12 cultures of T. equi and two cultures of T. equi mixed with Babesia cabal/i were established. None of the four ani-mals seropositive for B. cabal/i could be identified as carrier animals, whereas two seronegative sam-ples became cUlture-positive for B. cabal/i. Keywords: Babesia caballi, Babesia equi, Cape Mountain zebra, equine piroplasmosis, Equus zebra zebra, in vitro cultivation , Theileria equi INTRODUCTION Equine piroplasmosis is caused by two intraery-throcytic protozoa parasites, Babesia cabal/i and Theileria equi. The latter, previously known as Bab-esia equi, was recently reclassified (Mehlhorn & Schein 1998) due to the fact that it multiplies in lym-phoid cells before invading erythrocytes (Schein, Rehbein, Voigt & Zweygarth 1981). Both parasites are tick-transmitted and infect horses, mules, don-keys as well as zebras. * Author to whom correspondence is to be directed 1 Parasitology Division , Onderstepoort Veterinary Institute, Private Bag X05, Onderstepoort, 0110 South Africa 2 South African National Parks, PO. Box 110040, Hadison-park, 8306 South Africa Accepted for publication 13 May 2002-Editor Soon after T. equi was described in horses (Laver-an 1901) it was also described in an east African zebra (species unnamed)(Kudicke cited by Koch 1905). Inoculation of zebra blood into a susceptible horse caused high fever and the appearance of "Piroplasma equl', a synonym of T. equi, in the recipient horse. These findings led to the conclusion that zebras are carriers of T. equi, as are horses, donkeys and their crosses (Government Veterinary Bacteriologist 1905-1906). Subsequently, small piroplasms, presumably T. equi, were found in the blood of zebras (Equus quagga wahlbergi Pocock) (Neitz 1931). Young, Zumpt, Boomker, Penzhorn & Erasmus (1973) described T. equi in another zebra subspecies, the Cape Mountain zebra (Equus zebra zebra). In contrast to T. equi, B. caballi was only described in zebras more than 50 years later and citations in the literature are very rare. To our knowl-edge, Van Niekerk (1962) may have been the first to be mentioned as having found B. cabafli in 197 In vitro isolation of equine piroplasms from Cape Mountain zebra (Equus zebra zebra) in South Africa zebras (cited by Neitz 1965). On the basis of mor-phological characteristics Salabarria, Godshaev, Ferrer, Jimenez, Villalba & Jorge (1981) reported having found B. caballi in zebras exported from South Africa into Cuba. The aim of this study was to demonstrate and to isolate piroplasms from zebras using in vitro culture techniques. MATERIALS AND METHODS Blood samples Blood and serum samples from 20 Cape Mountain zebras (Equus zebra zebra) were collected by veni-puncture into sterile vacuum tubes (Vacutainer®, Becton Dickinson, Meylan, France) in the Karoo National Park (six samples in January 2000) and the Bontebok National Park (14 samples in March 2000). They were brought to the Onderstepoort Vet-erinary Institute (OVI) on ice in a polystyrene con-tainer. The blood samples from the Karoo National Park were severely haemolysed when processed. Upon arrival, all samples were processed as out-lined below. Thin blood smears were prepared from all of them, and were stained with Giemsa stain and examined microscopically. Serum was prepared and examined by the indirect fluorescent antibody (IFA) test for anti- T. equi and anti-B. caballi antibod-ies. This test was performed as described for B. caballi by Madden & Holbrook (1968). A titre of 1 :80 was considered to be positive. Culture medium A modified HL-1 medium (BioWhittaker, Walkers-ville, MD, USA) as described recently for the culture initiation of T. equi (Babesia eqw) (Zweygarth, Just, De Waal & Lopez-Rebollar 1999), was used. It was supplemented with 20% horse serum, 2 mM L-glut-amine, 0.2 mM hypoxanthine, 1 mM L-cysteine hydrochloride, 0.02 mM 2,9-dimethyl-4,7-diphenyl-1,10-phenanthrolinedisulphonic acid disodium salt (bathocuproine sulphonate, BCS; Serva Feinbio-chemica, Heidelberg, Germany), 100 IU/m£ penicil-lin and 100 mg/m£ streptomycin. The medium was buffered with 15 mM HEPES and 2.2 g/i! NaHC03. In vitro culture To initiate cultures, the red blood cells (RBC) were washed four times by centrifugation (650 x 9 for 10 min at room temperature) and resuspension in a modified Vega y Martinez phosphate-buffered saline solution (Vega, Buening, Green & Carson 1985) omitting adenine, guanosine and the antibiotics (mVYM). After each wash, the white blood cell layer overlaying the RBC was removed. After the fourth and final wash, the RBC were resuspended in the culture medium at a concentration of 10 % (v/v). Aliquots of 1 m£ were distributed into wells of 24-well culture plates (Corning, Bibby Sterilin, Staffordshire, England). Two different culture conditions were then applied: the plates were incubated at 37°C, one in a humidified 5% CO2-in-air atmosphere (Zweygarth et al. 1999), the other in a humidified gas mixture of 5 % CO2, 2 % O2 and 93 % N2 (Zweygarth, Just & De Waal 1995). The culture medium was changed daily by replacing 700 1-1£ medium overlaying the erythrocytes in each well. Every 4 days, 25 1-1£ of uninfected horse RBC suspension was added to each well. For this purpose, blood from an uninfect-ed donor horse, kept under tick-free conditions at the OVI, was collected and processed as described above. After the fourth wash, the horse RBC were suspended in mVYM solution and stored at 4 °C until used. Cultures were discontinued after 15 days for those samples in which no parasites were found in a Giemsa-stained thin blood smear, i.e. when no parasite was detected in approximately 20 000 RBC. Cultures were first examined on day 3 after culture initiation, and then at least every other day. RESULTS Before culture initiation, T. equi parasites were demonstrated in Giemsa-stained thin blood smears in 16 out of the 20 zebras. Of the cultures initiated with blood from the four zebras in which no para-sites had been demonstrated before culture initia-tion, all became positive. In four zebras (4, 13, 18 and 19) parasites were not detected beyond the examination period of at least 15 days although parasites had been demonstrated before culture initiation. Two cultures became contaminated (2 and 5) and were terminated prematurely. All of the zebra serum samples were serologically positive for T. equi, whereas antibodies against B. caballi were demonstrated only in zebras 1, 9, 15 and 20. No B. caballi parasites, however, were demonstrated in any of the samples serologically positive for B. caballi. In contrast, the blood of zebra 3, serologically negative for B. caballi, proved to be positive for both piroplasma species in culture. The results using an oxygen-reduced atmosphere for culture initiation are summarised in Table 1. E. ZWEYGARTH et al. TABLE 1 Culture initiation of blood samples1 obtained from zebras (Equus zebra zebra) from the Karoo National Park and the Bontebok National Park IFN Smear before Animal no. Culture results culture initiation T. equi B. caballi 1 - 640 160 Te 2 Te 640 - C(Te) 3 - 640 - Te + Bc 4 Te 640 - TeL14 5 Te 640 - C(Te) 6 Te 320 - Te 7 - 640 - Te 8 Te 320 - Te 9 Te 640 80 Te 10 Te 320 - Te 11 - 640 - Te 12 Te 320 - Te 13 Te 320 - Te L3 14 Te 640 - Te 15 Te 320 160 Te 16 Te 320 - Te 17 Te 320 - Te 18 Te 320 - TeL3 19 Te 160 - -20 Te 320 160 Te Samples 1-6 were derived from the Karoo National Park; samples 7-20 from the Bontebok National Park 2 Reciprocal titres obtained by the indirect fluorescent antibody test Te T. equi paraSites were detected in Giemsa-stained smears prepared from culture materi-al Bc B. caballi parasites were detected in Giemsa-stained smears prepared from culture material C Culture terminated due to bacterial contamination Parasites were positively identified in smears prepared from culture material but were no longer detectable on the day indicated In cultures initiated using a 5 % CO2-in-air atmos-phere, T. equi grew beyond day 15 in only four samples (1, 6, 9 and 14). However, B. caballi was identified in sample 6, which was serologically negative. DISCUSSION Of the 20 blood samples drawn from Cape Moun-tain zebra from two different National Parks in South Africa, 16 tested positive for T. equi in Giemsa-stained blood smears. The remaining four samples were identified as positive by in vitro culture. In an investigation into diseases of zebras in East Africa (Equus grevyi and Equus burchell1), Dennig (1966) found that 108 of 121 blood smears were positive for T. equi. He assumed that 100 % of the zebra population may have been infected but not all could be identified due to the poor quality of some blood smears. De Waal (1995) demonstrated that 61 % of the blood smears examined from zebras were pos-itive for T. equi, whereas in blood smears prepared from horses, parasites could only be detected in a few (~ 2 %). Similar low detection rates were found in 26 blood samples from carrier horses, only four being positive in thin Giemsa-stained blood smears (Zweygarth et al. 1999). On the other hand, Hol-man, Becu, Bakos, Polledo, Cruz & Wagner (1998) could not detect parasites in any of 23 horse blood samples, but T. equi was subsequently observed after culture in all of them. In the case of the zebras serologically positive for both B. caballi and T. equi, all four samples gave positive culture results for T. equi only. Babesia 199 In vitro isolation of equine piroplasms from Cape Mountain zebra (Equus zebra zebra) in South Africa caballi was not detected in any of these samples. However, two B. cabal/i-seronegative zebras were identified by culture as carriers for this parasite. The reason for the low in vitro recovery rate of B. cabal-Ii remains speculative: false positive serological reactions, clearance of B. cabal/i parasites from the animals' circulation or other unknown factors relat-ed to the sampling, the transportation, the storage before cultivation or the culture system itself. The comparison of the effects of an oxygen-reduced atmosphere and a 5 % CO2-in-air atmosphere on culture initiation of T equi from zebra blood showed that only the reduced oxygen system gave satisfac-tory results. This is in contrast with results of a recent survey of 43 horses of which 41 horses were identified as T equi carriers using the technique applying a normal oxygen tension (Zweygarth, un-published results 2001). Why these results could not be repeated with piroplasms derived from zebra blood is not clear. Possible reasons may be related to the host celis, the parasites or both. Babesia caballi was found in one culture sample each initi-ated in an oxygen-reduced atmosphere and in a 5 % CO2-in-air atmosphere. ACKNOWLEDGEMENTS We thank Drs D.T. de Waal and B.A. Allsopp for helpful comments on the manuscript. REFERENCES DENNIG, H.K. 1966. The isolation of Babesia species from wild animals. Proceedings of the First International Congress of Parasitology, Rome 1964, edited by A. Corradetti. Oxford: Pergamon Press: 262-263. DE WAAL, D.T. 1995. Distribution, transmission and serodiagno-sis of Babesia equi and Babesia cabal/i in South Africa. Ph.D. thesis, University of Pretoria. GOVERNMENT VETERINARY BACTERIOLOGIST 1905-1906. Inoculation of zebra blood into a susceptible horse. Report of 200 the Government Veterinary Bacteriologist. Pretoria: Government Printer. HOLMAN P.J. , BECU, 1., BAKOS, E., POLLEDO, G., CRUZ, D. & WAGNER, G.G. 1998. Babesia equifield isolates cultured from horse blood using a microcentrifuge method. Journal of Parasitology, 84:696-699. KOCH, R. 1905. Vorlaufige Mitteilungen Ober die Ergebnisse einer Forschungsreise nach Ostafrika. Deutsche Medizin-ische Wochenschrift, 31 :1865-1869. LAVERAN, A. 1901. Contribution a I' etude du Piroplasma equi. Comptes rendus des seances. Societe de Biologie et de ses Filiales, 53:385-388. MADDEN, PA. & HOLBROOK, A.A. 1968. Equine piroplasmo-sis: indirect fluorescent antibody test for Babesia cabal/i. American Journal of Veterinary Research, 29:117-123. MEHLHORN, H. & SCHEIN, E. 1998. Redescription of Babesia equi Laveran, 1901 as Theileria equi Mehlhorn, Schein 1998. Parasitology Research, 84:467-475. NEITZ, WO. 1931. Blood parasites of game in Zululand. 17th Report of the Director of Veterinary Services and Animal Industry, Union of South Africa. Pretoria: Government Printer. NEITZ, WO. 1965. Classification, transmission, and biology of piroplasms of domestic animals. Annals of the New York Academy of Science, 64:56-111 . SALABARRIA, F.F, GODSHAEV, A. , FERRER, J., JIMENEZ, 1., VILLALBA, G. & JORGE, J. 1981. Babesia (Nuttalia) equiy Babesia caballi en cebras (Equus zebra) . Diagnostico mor-fologico y serologico. Revista Cubana de Ciencias Vet-erinarias, 12: 171-176. SCHEIN, E., REHBEIN, G., VOIGT, WP & ZWEYGARTH, E. 1981. Babesia equi (Laveran 1901) 1. Development in hors-es and in lymphocyte culture. Tropenmedizin und Parasito-logie, 32:223-227. YOUNG, E. , ZUMPT, F. , BOOMKER, J., PENZHORN, B.L. & ERASMUS, B. 1973. Parasites and diseases of Cape Moun-tain zebra, black wildebeest, mountain reedbuck and bles-bok in the Mountain Zebra National Park. Koedoe, 16:77-81. VEGA, C.A., BUENING, G.M., GREEN, TJ. & CARSON, CA 1985. In vitro cultivation of Babesia bigemina. American Journal of Veterinary Research, 46:416-420. ZWEYGARTH, E., JUST, M.C. & DE WAAL, D.T 1995. Continu-ous in vitro cultivation of erythrocytic stages of Babesia equi. Parasitology Research, 81 :355-358. ZWEYGARTH, E. , JUST, M.C., DE WAAL, D.T. & LOPEZ-REBOLLAR, L.M. 1999. In vitro diagnosis of Babesia equi, in Equine Infectious Diseases VIII, edited by U. Wernery, J.F. Wade, JA Mumford & O.-R. Kaaden. Newmarket: R & W Publications (Newmarket) Ltd. 

In vitro isolation of equine piroplasms derived from Cape Mountain zebra (Equus zebra zebra) in South Africa

https://repository.up.ac.za/bitstream/2263/18339/1/21zweygarth2002.pdf

In vitro isolation of equine piroplasms derived from Cape Mountain zebra (Equus zebra zebra) in South Africa

Abstract

Similar works

Full text

Available Versions

UPSpace at the University of Pretoria