JOURNAL OF CLINICAL MICROBIOLOGY, Sept. 1995, p. 2501–2504 0095-1137/95/$04.0010 Copyright q 1995, American Society for Microbiology
Vol. 33, No. 9
Detection of Cowdria ruminantium in Blood and Bone Marrow Samples from Clinically Normal, Free-Ranging Zimbabwean Wild Ungulates NANCY D. KOCK,1,2 ARNOUD H. M.
VAN
VLIET,3 KRISTEN CHARLTON,1
AND
FRANS JONGEJAN3*
Department of Paraclinical Veterinary Studies, Faculty of Veterinary Science, University of Zimbabwe, Mount Pleasant, Harare, Zimbabwe1; International Wildlife Veterinary Services, Salinas, California2; and Department of Bacteriology, Institute of Infectious Diseases and Immunology, Faculty of Veterinary Medicine, University of Utrecht, 3508 TD Utrecht, The Netherlands3 Received 29 March 1995/Accepted 19 June 1995
Cowdria ruminantium causes severe, often fatal disease in domestic ruminants, whereas wildlife species usually are not affected. Blood and bone marrow samples from healthy, free-ranging Zimbabwean ungulates were taken during translocation from areas harboring Amblyomma ticks and tested for the presence of C. ruminantium, using a PCR assay based on the C. ruminantium map1 gene. Positive reactions were obtained in tsessebe (Damaliscus lunatus), waterbuck (Kobus ellipsiprymnus), and impala (Aepyceros melampus). Wildlife species may therefore be a reservoir for C. ruminantium thus contributing to the spread of cowdriosis. ATACAGGAAGAG-39) and 32R1 (59-CTATTCTTGGTCC ATTC-39) were based on the nucleotide sequence of the map1 gene of C. ruminantium (23). These primers amplified a fragment of 297 bp. PCR was carried out by using 35 cycles of 1 min of denaturation at 948C, 1 min of annealing at 558C, and 1 min of extension at 728C, using a thermal cycler (PerkinElmer Cetus, Norwalk, Conn.) and Taq polymerase (Promega, Madison, Wis.). Because antibodies to several Ehrlichia spp. recognize epitopes on the C. ruminantium MAP1 protein (8, 13, 24) and the MAP1 protein is homologous to the Anaplasma marginale MSP4 protein (23), primers were first tested for specificity. C. ruminantium (Senegal isolate [9]), Ehrlichia chaffeensis (HE17 isolate [7]), and Ehrlichia canis were cultivated in BUE9, Vero, and DH82 cells, respectively. Anaplasma marginale and Theileria annulata were purified from infected bovine erythrocytes. Genomic DNA from infected and uninfected cells was purified as described previously (22) and used as template in PCR. To show the presence of rickettsial DNA in the test samples, a fragment of 431 bp of the 16S rRNA gene was amplified by using primers FEhrl [59-TAAA(A/G)T(A/G) GGGAAGATAATG-39] and REhrl (59-GACCGTAGTCCC CAGGCGG-39). PCR products were separated by electrophoresis in an 1.5% agarose gel, denatured, and blotted onto Hybond-N membranes (Amersham, Buckinghamshire, England) by standard methods (18). The 1.2-kb HindIII insert of plasmid pCRS18 (23), containing two-thirds of the C. ruminantium map1 gene, was used as a probe after labelling with digoxigenin as described by the manufacturer (Boehringer, Mannheim, Germany). Hybridization, washing, and autoradiography were carried out under conditions described previously (23). A map1 amplification product was found only by using C. ruminantium DNA (Fig. 1), whereas rickettsial 16S rDNA was amplified from C. ruminantium, A. marginale, and the ehrlichial DNA samples (Fig. 1). Thus the 32F1 and 32R1 primer set is unique for C. ruminantium. PCR detection of C. ruminantium in blood from ruminants has so far only been successful on plasma samples taken from infected animals during the febrile period of the disease (14) but not on whole blood samples. We developed a PCR method for the detection of C. ruminantium in whole blood and bone marrow samples taken from experimentally infected sheep. A.
Cowdriosis, or heartwater, is a disease of ruminants, caused by the rickettsia Cowdria ruminantium and transmitted by ticks of the genus Amblyomma. The disease is present in most African countries south of the Sahara and on islands off the African mainland and in the Caribbean Sea (20). In addition, it threatens other parts of the world, because of its vector transmission and wide host range (3, 20). Fatal disease often occurs in cattle, sheep, and goats, particularly those of European origin, making C. ruminantium a major impediment against the introduction of exotic breeds into Africa for genetic improvement (20). Cattle, sheep, and African buffalo (Syncerus caffer) in Zimbabwe (1); goats in Guadeloupe (2); and some strains of laboratory mice (25) remain carriers of C. ruminantium for long periods and therefore serve as reservoirs of the infection. C. ruminantium would be expected to be found throughout the body during acute infection, given its tropism for endothelial cells, but it is conspicuously present in brain capillaries, kidney, lung, and heart only by light microscopy (19). The organism has also been found in neutrophils of acutely infected goats (12) and also in the spleen of mice (10). Because neutrophils circulate for only 6 to 8 h before entering various tissues (21), it seemed reasonable to assume that precursors of neutrophils and perhaps other leukocyte precursors become infected. Therefore we tested bone marrow samples as well as blood samples from the same animals. Wildlife species are suspected to be important reservoirs for C. ruminantium (1, 16), by being alternative hosts for Amblyomma ticks in areas where tick control methods on domestic ruminants are exercised (15). In Zimbabwe, extensive translocation of wildlife from areas in which heartwater is endemic to heartwater-free areas may facilitate the spread of heartwater (15). A PCR assay was developed to test for the presence of C. ruminantium in whole blood and bone marrow samples from carrier animals. Oligonucleotide primers 32F1 (59-GATGTA * Corresponding author. Mailing address: Department of Bacteriology, Institute of Infectious Diseases and Immunology, P.O. Box 80.165, 3508 TD Utrecht, The Netherlands. Phone: 31-30-532568. Fax: 31-30540784. Electronic mail (Internet):
[email protected]. 2501
2502
NOTES
J. CLIN. MICROBIOL. TABLE 1. Results from PCR detection of C. ruminantium in blood and bone marrow of acutely infected and immune sheep, healthy tsessebe (Damaliscus lunatus), waterbuck (Kobus ellipsiprymnus), impala (Aepyceros melampus), and reedbuck (Redunca arindinum) No. positive/no. tested Species
Tsessebe Waterbuck Impala Reedbuck Sheep Infected Noninfected a
FIG. 1. Demonstration of specificity of the map1-based PCR assay. (A) Ethidium bromide staining of DNA amplified with map1 primers (left panel) and rickettsial 16S rDNA primers (right panel). (B) Hybridization of map1 PCR products with a map1 probe (pCRS18). Lanes: 1, negative control (H2O); 2, C. ruminantium; 3, BUE9 cells; 4, E. chaffeensis; 5, Vero cells; 6, E. canis; 7, DH82 cells; 8, A. marginale; 9, T. annulata; M, marker (lambda DNA digested with PstI).
variegatum nymphal ticks infected with either the Senegal or the Lutale isolate of C. ruminantium (9) were fed on five adult female Tesselaar sheep. Four sheep, two infected with the Lutale isolate and two out of three infected with the Senegal isolate of C. ruminantium were treated with tetracycline (5 mg/kg once a day for 2 days) when the body temperature remained above 418C for 3 days. The fifth sheep was not treated and died of heartwater 21 days after infestation with ticks. Death due to heartwater was confirmed by detection of C. ruminantium in brain smears. A sixth sheep which had never been exposed to C. ruminantium served as negative control. Bone marrow and blood were collected under light anesthesia from the sheep infected with the Senegal isolate (2 months after infection), from the sheep infected with the Lutale isolate (4 months after infection), and from the negative control sheep. Aspirates of bone marrow (2 to 4 ml) were collected by a trephine punch into the right tuber coxae and placed directly into Vacutainer tubes containing sodium citrate. Blood was collected in sodium citrate by venipuncture. Blood was also collected during febrile disease from the sheep that died, and bone marrow was taken at necropsy. DNA was prepared from 1-ml samples after thawing from 2208C. Hemoglobin was removed by repeated washing with an equal volume of lysis buffer (0.22% NaCl, 0.015% saponin, 1 mM EDTA) and centrifuging for 5 min at 14,000 3 g. Cell pellets were then washed twice with distilled water, incubated with proteinase K and sodium dodecyl sulfate at final concentrations of 400 mg/ml and 0.5%, respectively, for 2 h at 558C. DNA was precipitated with ethanol and resuspended in 50 ml of H2O, and 10 ml was used as template in the PCR reactions. A map1 product was amplified from blood of the sheep which died from heartwater but not from the negative control sheep (Table 1). A map1 product was also amplified from bone marrow samples of all four carrier sheep and from blood of three out of four carrier sheep (Table 1). A representative sample of these PCRs is shown in Fig. 2. The second band visible upon hybridization with pCRS18 (Fig. 2b and 3b) was not visible on ethidium bromide staining of agarose gel-separated PCR prod-
Blood
Bone marrow
11/24 3/3 NAa 0/3
15/28 0/3 1/3 NA
4/5 0/1
5/5 0/1
NA, samples not available.
ucts (Fig. 2a and 3a). This phenomenon could not be explained. Blood and bone marrow were similarly taken from healthy, anesthetised tsessebe (Damaliscus lunatus), waterbuck (Kobus ellipsiprymnus), reedbuck (Redunca arindinum), and impala (Aepyceros melampus) prior to translocation within Zimbabwe from low-altitude regions in which A. hebraeum ticks occur alone or simultaneously with A. variegatum (6). Samples were prepared as described and used as template in PCR. map1 sequences were amplified from the blood samples of 11 out of 24 tsessebe and three out of three waterbuck (Table 1). Bone marrow samples were positive for map1 sequences with 15 out of 28 tsessebe and one out of three impala (Table 1). Bone marrow from the three waterbuck were negative, and the three blood samples from reedbuck were also negative (Table 1). Of the 17 tsessebe for which blood and bone marrow were both available, 6 were positive in both blood and bone marrow samples, 3 were positive in blood samples only, 5 were positive in bone marrow samples only, and 3 were negative in both blood and bone marrow samples. Figure 3 contains representative samples of these PCRs. Positivity for map1 sequences was confirmed by Southern hybridization with pCRS18 (Fig. 3b).
FIG. 2. Agarose gel electrophoresis and subsequent Southern hybridization of amplified map1 sequences from blood and bone marrow samples of sheep experimentally infected with C. ruminantium (Lutale or Senegal isolate). (A) Ethidium bromide staining. (B) Hybridization with map1 gene. Lanes: 1, positive control (C. ruminantium genomic DNA); 2, negative control (H2O); 3 and 5, blood from sheep infected with C. ruminantium (Lutale isolate) 4 months prior; 4 and 6, bone marrow from sheep infected with C. ruminantium (Lutale isolate) 4 months prior; 7 and 9, blood from sheep infected with C. ruminantium (Senegal isolate) 2 months prior; 8 and 10, bone marrow from sheep infected with C. ruminantium (Senegal isolate) 2 months prior; 11 and 12, blood and bone marrow from sheep with acute infection with C. ruminantium (Senegal isolate); 13 and 14, blood and bone marrow from uninfected sheep.
VOL. 33, 1995
FIG. 3. Agarose gel electrophoresis and subsequent Southern hybridization of amplified map1 sequences from blood and bone marrow samples of healthy tsessebe (Damaliscus lunatus), waterbuck (Kobus ellipsiprymnus), impala (Aepyceros melampus), and reedbuck (Redunca arindinum). (A) Ethidium bromide staining. (B) Hybridization with map1 gene. Lanes: 1, positive control (C. ruminantium genomic DNA); 2, negative control (H2O), 3 and 4, blood and bone marrow from waterbuck; 5, 7, 9, and 11, blood from tsessebe; 6, 8, 10, and 12, bone marrow from tsessebe; 13, blood from reedbuck; 14, bone marrow from impala; 15 and 16, blood and bone marrow from sheep with acute infection with C. ruminantium (Senegal isolate); 17 and 18, blood and bone marrow from uninfected sheep, M, molecular size marker.
The development of PCR for the detection of C. ruminantium in whole blood and bone marrow has practical applications for the diagnosis of heartwater during natural infections as well as under experimental conditions. At present, brain smears are often relied upon for definitive postmortem diagnosis of acute heartwater in domestic animals (20). Serodiagnosis is hampered by unavailability of wildlife species-specific conjugates and by relatively short duration of high antibody levels (50 to 200 days in experimental animals [24]) making it less suitable for the detection of carriers. Furthermore, wildlife species can be infected with Ehrlichia spp., and these are a cause of cross-reactive antibodies. For instance, the whitetailed deer (Oidocoileus virginianus) is susceptible to experimental infection with C. ruminantium (4) and is a possible reservoir host for E. chaffeensis (5), an organism eliciting crossreactive antibodies with the C. ruminantium MAP1 protein (24). PCR on ticks fed on infected animals or experimental transmission of C. ruminantium by ticks are both used experimentally to identify carriers, but both methods are more laborious than direct detection of the organism in blood or bone marrow samples. It is noteworthy that two isolates of C. ruminantium from different parts of Africa were detected by PCR in sheep. The Senegal isolate originates from Senegal and the Lutale isolate originates from Zambia, and neither of them may be the same as those detected here in the wild ungulates in Zimbabwe. As in the other diagnostic tests for heartwater, the question of cross-reactivity with Ehrlichia spp. arises (8, 13, 24). Whether infection with Ehrlichia spp. occurs in the wildlife studied here is unknown. DNA from other Ehrlichia spp. remains to be tested with the primers described herein. Isolation and subsequent characterization of the organism(s) causing the positive reactions with the PCR and also those causing false-positive serological reactions in domestic ruminants in heartwater-free areas of Zimbabwe (13) are needed before definitive conclusions can be drawn. As carriers for C. ruminantium have been demonstrated experimentally in mice (25), African buffalo (1), and domestic ruminants (1, 2), reservoir sites within the carrier host must exist. The successful amplification of C. ruminantium from bone marrow but not from blood in some instances in tsessebe and carrier sheep warrants further investigation of this site as a possible reservoir for the organism.
NOTES
2503
Naturally infected wildlife carriers of C. ruminantium have been suspected in Africa for some time (11, 16). Natural or experimental infection with heartwater has been reported in springbuck, eland, sitatunga, and lechwe, whereas subclinical infection has been demonstrated in African buffalo and giraffe (16, 20). One natural outbreak of disease has been described in lechwe (Kobus leche kafuensis) during translocation in Zambia (17). Given the positive results in this study, it is likely that tsessebe and waterbuck, and possibly impala, carry C. ruminantium asymptomatically. The presence of naturally infected, asymptomatic carriers may make translocation of these species into heartwater-free areas risky. This is of particular concern for Zimbabwe itself, which translocates wildlife within the country as well as abroad. The wild ungulates in this study originated from low-altitude regions that harbor A. hebraeum alone, or concurrently with A. variegatum (6, 15). Further use of the methods developed here will be applied to other wildlife species in Zimbabwe, attempting to further define the role of wildlife in the epidemiology of heartwater. This work was supported by the Commission of the European Union (EU), Directorate General XII, STD-3 program under contract no. TS3*-CT91-0007, the EU-funded University of Utrecht-University of Zimbabwe Link programme, the University of Zimbabwe Research Board, and the International Wildlife Veterinary Services. A. Rijkenhuizen, B. Ahmer, and J. B. de Kok are acknowledged for technical assistance, G. Uilenberg and B. A. M. van der Zeijst for critical reading of the manuscript, and P. Crocquet-Valdes and D. H. Walker (Galveston, Tex.) for providing Ehrlichia chaffeensis HE17 DNA. REFERENCES 1. Andrew, H. R., and R. A. I. Norval. 1989. The carrier status of sheep, cattle and African buffalo recovered from heartwater. Vet. Parasitol. 34:261–266. 2. Barre´, N., and E. Camus. 1987. The reservoir status of goats recovered from heartwater. Onderstepoort. J. Vet. Res. 54:435–437. 3. Barre´, N., G. Uilenberg, P. C. Morel, and E. Camus. 1987. Danger of introducing heartwater onto the American mainland: potential role of indigenous and exotic Amblyomma ticks. Onderstepoort. J. Vet. Res. 54:405– 417. 4. Dardiri, A. H., L. L. Logan, and C. A. Mebus. 1987. Susceptibility of whitetailed deer to experimental heartwater infections. J. Wildl. Dis. 23:215–219. 5. Dawson, J. E., D. E. Stallknecht, E. W. Howerth, C. Warner, K. Biggie, W. R. Davidson, J. M. Lockhart, V. F. Nettles, J. G. Olson, and J. E. Childs. 1994. Susceptibility of white-tailed deer (Odocoileus virginianus) to infection with Ehrlichia chaffeensis, the etiologic agent of human ehrlichiosis. J. Clin. Microbiol. 32:2725–2728. 6. de Vries, N., S. M. Mahan, U. Ushewokunze-Obatolu, R. A. I. Norval, and F. Jongejan. 1993. Correlation between antibodies to Cowdria ruminantium (Rickettsiales) in cattle and the distribution of Amblyomma vector ticks in Zimbabwe. Exp. Appl. Acarol. 17:799–810. 7. Dumler, J. S., and D. H. Walker. 1993. Isolation and partial characterization of an Ehrlichia isolate from a patient with near-fatal ehrlichiosis, abstr. D-89, p. 111. In Abstracts of the 93rd General Meeting of the American Society for Microbiology. American Society for Microbiology, Washington, D.C. 8. Jongejan, F., N. de Vries, J. Nieuwenhuijs, A. H. M. van Vliet, and L. A. Wassink. 1993. The immunodominant 32-kilodalton protein of Cowdria ruminantium is conserved within the genus Ehrlichia. Rev. Elev. Med. Vet. Pays Trop. 46:145–152. 9. Jongejan, F., G. Uilenberg, F. F. J. Franssen, A. Gueye, and J. Nieuwenhuijs. 1988. Antigenic differences between stocks of Cowdria ruminantium. Res. Vet. Sci. 44:186–189. 10. Kock, N. D. Unpublished results. 11. Kock, N. D., F. Jongejan, M. D. Kock, R. A. Kock, and P. Morkel. 1993. Serological evidence for Cowdria ruminantium infection in free-ranging black (Diceros bicornis) and white (Ceratotherium simum) rhinoceroses in Zimbabwe. J. Zoo Wildl. Med. 23:409–413. 12. Logan, L. L., T. C. Whyard, J. C. Quintero, and C. A. Mebus. 1987. The development of Cowdria ruminantium in neutrophils. Onderstepoort. J. Vet. Res. 54:197–204. 13. Mahan, S. M., N. Tebele, D. Mukwedeya, S. Semu, C. B. Nyathi, L. A. Wassink, P. J. Kelly, T. Peter, and A. F. Barbet. 1993. An immunoblotting diagnostic assay for heartwater based on the immunodominant 32-kilodalton protein of Cowdria ruminantium detects false positives in field sera. J. Clin. Microbiol. 31:2729–2737. 14. Mahan, S. M., S. D. Waghela, T. C. McGuire, F. R. Rurangirwa, L. A.
2504
15.
16. 17. 18. 19. 20.
NOTES
Wassink, and A. F. Barbet. 1992. A cloned DNA probe for Cowdria ruminantium hybridizes with eight heartwater strains and detects infected sheep. J. Clin. Microbiol. 30:981–986. Norval, R. A. I., B. D. Perry, M. I. Meltzer, R. L. Kruska, and T. H. Booth. 1994. Factors affecting the distributions of the ticks Amblyomma hebraeum and A. variegatum in Zimbabwe: implications of reduced acaricide usage. Exp. Appl. Acarol. 18:383–407. Oberem, P. T., and J. D. Bezuidenhout. 1987. Heartwater in hosts other than domestic ruminants. Onderstepoort. J. Vet. Res. 54:271–275. Pandey, G. S., D. Minyoi, F. Hasebe, and E. T. Mwase. 1992. First report of heartwater (cowdriosis) in Kafue lechwe (Kobus leche kafuensis) in Zambia. Rev. Elev. Med. Vet. Pays Trop. 45:23–25. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning, a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Uilenberg, G. 1983. Heartwater (Cowdria ruminantium infection): current status. Adv. Vet. Sci. Comp. Med. 27:428–455. Uilenberg, G., and E. Camus. 1993. Heartwater (cowdriosis), p. 293–332. In Z. Woldehiwet and M. Ristic (ed.), Rickettsial and chlamydial diseases of domestic animals. Pergamon Press, Oxford.
J. CLIN. MICROBIOL. 21. Valli, V. E. O. 1985. The hematopoietic system, p. 88. In K. V. F. Jubb, P. C. Kennedy, and N. Palmer (ed.), Pathology of domestic animals. Academic Press, London. 22. van Vliet, A. H. M., F. Jongejan, and B. A. M. van der Zeijst. 1992. Phylogenetic position of Cowdria ruminantium (Rickettsiales) determined by analysis of amplified 16S ribosomal DNA sequences. Int. J. Syst. Bacteriol. 42:494–498. 23. van Vliet, A. H. M., F. Jongejan, M. van Kleef, and B. A. M. van der Zeijst. 1994. Molecular cloning, sequence analysis, and expression of the gene encoding the immunodominant 32-kilodalton protein of Cowdria ruminantium. Infect. Immun. 62:1451–1456. 24. van Vliet, A. H. M., B. A. M. van der Zeijst, E. Camus, S. M. Mahan, D. Martinez, and F. Jongejan. 1995. Use of a specific immunogenic region on the Cowdria ruminantium MAP1 protein in a serological assay. J. Clin. Microbiol. 33:2405–2410. 25. Wassink, L. A., F. Jongejan, E. Gruys, and G. Uilenberg. 1990. Observations on mouse-infective stocks of Cowdria ruminantium: microscopical demonstration of the Kwanyanga stock in mouse tissue and the carrier-status of the Senegal stock in mice. Res. Vet. Sci. 48:389–390.
