Hostname: page-component-586b7cd67f-dlnhk Total loading time: 0 Render date: 2024-11-29T18:22:41.945Z Has data issue: false hasContentIssue false
  • English
  • Français

Leishmania parasites and their kinetoplast DNA (kDNA)

Published online by Cambridge University Press:  19 September 2011

N. N. Massamba ,
M. J. Mutinga  and
B. N. Odero
N. N. Massamba
Affiliation:
The International Centre of Insect Physiology and Ecology (ICIPE), P. O. Box 30772, Nairobi, Kenya
M. J. Mutinga
Affiliation:
The International Centre of Insect Physiology and Ecology (ICIPE), P. O. Box 30772, Nairobi, Kenya
B. N. Odero
Affiliation:
The International Centre of Insect Physiology and Ecology (ICIPE), P. O. Box 30772, Nairobi, Kenya
Get access

Abstract

Restriction endonuclease digestion of genomic DNAs from WHO Leishmania reference strains generates prominent kinetoplast DNA (kDNA) fragments ranging from 0.5–2.0 kb. Following fractionation by agarose gel electrophoresis, these DNA fragments were visualized as distinct bands in gels. Using as parameters the presence or absence of the prominent bands and their size and number, the Leishmania reference strains fell into three distinct genomic groups. These genomic groupings were applied to new Leishmania isolates. DNA analysis, involving restriction endonuclease digestion and Southern blot hybridization with probes generated from the prominent DNA bands of reference strains L. major IC-236 and L. major IC-235, cloned in plasmids, showed significant genetic variation within Leishmania and allowed some distinct isolates to be identified.

Résumé

La digestion aux enzymes de restriction des ADN génomiques obtenus à partir des souches de référence de Leishmania génère des fragments caractéristiques des ADN (ADN kinétoplastique) dont la taille est comprise entre 0.5–2.0 kb. Le fractionnement par électrophorèse sur gel d'agarose de ces fragments d'ADN révèle des bandes proéminentes distinctes. En se servant comme paramètres, la présence ou l'absence de ces bandes, leur taille et nombre, il a été possible de répartir les souches de référence de Leishmania en trois différents groupes génomiques. L'application de ce mode de partage aux nouveaux isolats est rapportée dans cette étude. L'analyse de l'ADN, faisant intervenir la digestion aux enzymes de restriction, l'hybridation à la Southern avec des sondes préparées à partir des fragments caractéristiques de l'ADN des souches de référence L. major IC-236 et L. major IC-235, clonés dans les plasmides a montré une variation génétique significative au sein des Leishmania et quel ques isolats ont été ainsi identifiés.

Keywords

WHO Leishmania reference strainskinetoplast DNA (kDNA)kilobase (kb)DNA probessouthern blot hybridizationSouches de référence de LeishmaniaADN kinétoplastiquekilobase (kb)sondes d'ADNhybridation selon southern
Type
Research Articles
Information
International Journal of Tropical Insect Science , Volume 13 , Issue 3 , June 1992 , pp. 293 - 306
Copyright
Copyright © ICIPE 1992

Access options

Get access to the full version of this content by using one of the access options below. (Log in options will check for institutional or personal access. Content may require purchase if you do not have access.)

References

REFERENCES

Arnot, D. E. and Barker, D. C. (1981) Biochemical identification of cutaneous leishmaniasis by analysis of kinetoplast DNA. II. Sequence homologies in Leishmania kDNA. Mol. Biochem. Parasitol. 3, 47–56.Google Scholar
Barker, D. C. (1980) The ultrastructure of kinetoplast DNA with particular reference to the interpretation of darkfield electron microscopy images of isolated purified networks. Micron 11, 21–62.Google Scholar
Barker, D. C. (1987) DNA diagnosis of human leishmaniasis. Parasitol. Today 3, 177–184.Google Scholar
Barker, D. C. (1989) Molecular approaches to DNA diagnosis. Parasitology 99, S125–S146.CrossRefGoogle ScholarPubMed
Barker, D. C. and Arnot, D. E. (1981) Biochemical identification of cutaneous leishmaniasis by analysis of kinetoplast DNA. I. Ultrastructure and buoyant density analysis. Mol. Biochem. Parasitol. 3, 33–46.Google Scholar
Barker, D. C., Arnot, D. E. and Butcher, J. (1982) DNA characterization as a taxonomic tool for identification of kinetoplastic flagellate protozoans. In Proceedings of the Workshop of the Pan American Health Organization. Biochemical Characterization of Leishmania. Washington, DC (1982) (Edited by Chance, M. L. and Walton, B. C.), pp. 139–180. UNDP/ World Bank/WHO, Geneva.Google Scholar
Barker, D. C. and Butcher, J. (1983) The use of DNA probes in the identification of Leishmania: discrimination between isolates of the Leishmania mexicana and L. braziliensis complexes. Trans. R. Soc. Trop. Med. Hyg. 77, 285–297.Google Scholar
Barker, D. C., Gibson, L. J., Kennedy, W. P. K., Nasser, A. and Williams, R. H. (1986) The potential of using recombinant DNA species-specific probes for the identification of tropical leishmaniases. Parasitology 91, S139–S174.CrossRefGoogle Scholar
Bray, R. S., Ashford, R. W. and Bray, M. A. (1973) The parasite causing cutaneous leishmaniasis in Ethiopia. Trans. R. Soc. Trop. Med. Hyg. 67, 345–348.CrossRefGoogle ScholarPubMed
Beach, R., Kiilu, G., Hendricks, L. D., Oster, C. N. and Leeuwenburg, J. (1984) Cutaneous leishmaniasis in Kenya: transmission of Leishmania major to man by the bite of naturally infected Phlebotomus duboscqi. Trans. R. Soc. Trop. Med. Hyg. 78, 747–751.Google Scholar
Borst, P. and Hoeijmakers, J. H. J. (1979) Kinetoplast DNA (Review). Plasmid 2, 20–40.CrossRefGoogle Scholar
Borst, P., Fase-Fowler, F., Weijers, P. J., Barry, D. J., Tetley, L. and Vickerman, K. (1985) Kinetoplast DNA from Trypanosoma vivax and T. congolense. Mol. Biochem. Parasitol. 15, 129–142.CrossRefGoogle ScholarPubMed
Chance, M. L., Schnur, L. F., Thomas, S. C. and Peters, W. (1978) The biochemical and serological taxonomy of Leishmania from Aethiopian zoogeographical region. Ann. Trop. Med. Parasitol. 72, 533–542.CrossRefGoogle ScholarPubMed
Denhardt, D. T. (1966) A membrane-filter technique for the detection of complementary DNA. Biochem. Biophys. Res. Comm. 23, 641–646.Google Scholar
Fendall, N. R. E. (1952) Kala-azar in East Africa with particular reference to Kenya and Kamba country, I and II. J. Trop. Med. Hyg. 55, 193–204.Google ScholarPubMed
Gardener, P. J. and Howells, R. E. (1972) Isoenzyme variation in leishmanial parasites. J. Protozool. 19, (supplement), 47.Google Scholar
Gardener, P. J., Chance, M. L. and Peters, W. (1974) Biochemical taxonomy of Leishmania II. Electrophoretic variation of malate dehydrogenase. Ann. Trop. Parasitol. 68, 317–325.Google Scholar
Gibson, W., Borst, P. and Fase-Fowler, F. (1985) Further analysis of intraspecific variation in Trypanosoma brucei using restriction site polymorphisms in the maxicircle of kinetoplast DNA. Mol. Biochem. Parasitol. 15, 21–36.CrossRefGoogle ScholarPubMed
Gomez-Eichelman, C. M., Holz, G. Jr, Beach, D., Simpson, A. M. and Simpson, L. (1988) Comparison of several lizard Leishmania species and strains in terms of kinetoplast minicircle and maxicircle DNA sequences, nuclear chromosomes, and membrane lipids. Mol. Biochem. Parasitol. 27, 143–158.CrossRefGoogle Scholar
Grimaldi, G. Jr, Momen, H., Soares, M. J. and Moriearty, P. L. (1982) Enzyme variation and difference in infectivity within a single strain of Leishmania mexicana. Int. J. Parasitol. 12, 185–189.Google Scholar
Grunstein, M. and Hogness, D. (1975) Colony hybridization: A method for the isolation of cloned DNAs that contain a specific gene. Proc. Natl. Acad. Sci. (USA). 72, 3961–3965.CrossRefGoogle ScholarPubMed
Ho, M., Leeuwenburg, J., Mbugua, A., Wamachi, A. and Voller, A. (1983) An enzyme-linked immunosorbent assay (Elisa) for field diagnosis of visceral leishmaniasis. Am. J. Trop. Med. Hyg. 32, 943–946.Google Scholar
Hoeijmakers, J. H. J., Borst, P., Van de Burg, J., Weismann, C. and Cross, G. A. M. (1980) The isolation of plasmids containing DNA complementary to messenger RNA for variant surface glycoproteins of Trypanosome brucei. Gene 8, 391–417.CrossRefGoogle Scholar
Jackson, P. R., Wohlhieter, J. A., Jackson, J. E., Sayles, P., Diggs, C. L. and Hockmeyer, W. T. (1984) Restriction endonuclease analysis of Leishmania kinetoplast DNA characterizes parasites responsible for visceral and cutaneous disease. Am. J. Trop. Hyg. 33, 808–819.CrossRefGoogle ScholarPubMed
Jackson, P. R., Lawrie, J. M., Stiteler, J. M., Hawkins, D. N., Wohlhieter, J. A. and Rowton, E. D. (1986) Detection and characterization of Leishmania species and strains from mammals and vectors by hybridization and restriction endonuclease digestion of kinetoplast DNA. Vet. Parasitol. 20, 195–215.CrossRefGoogle ScholarPubMed
Jaffe, C. L. and Zallis, M. (1988) Use of purified parasite proteins from Leishmania donovani for rapid serodiagnosis of visceral leishmaniasis. J. Infect. Dis. 157, 1212–1220.Google Scholar
Kaddu, J. B. and Mutinga, M. J. (1984) Leishmania in Kenyan Phlebotomine sandflies — II. Natural infection in the malpighian tubules of Sergentomyia garnhami and Sergentomyia antennatus. Insect Sci. Applic. 5, 239–243.Google Scholar
Kaddu, J. B., Mutinga, M. J., Chimtawi, M., Okot-Kotber, B. M., Nyamori, M. P. and Musyoki, R. (1988) Leishmania in Kenyan Phlebotomine sandflies — V. Leishmania aethiopica in the oesophagus of Phlebotomus pedifer. Insect Sci. Applic. 9, 117–121.Google Scholar
Kungu, A., Mutinga, M. J. and Ngoka, J. M. (1972) Cutaneous leishmaniasis in Kenya. East Afr. Med. J. 49, 458–465.Google ScholarPubMed
Laurent, M., Van Assel, S. and Steinert, M. (1971) Kinetoplast DNA. A unique macromolecular structure of considerable size and mechanical resistance. Bioch. Biophys. Res. Com. 43, 278–284.Google Scholar
Lawrie, J. M., Jackson, P. R., Stiteler, J. M. and Hockmeyer, W. T. (1985) Identification of pathogenic Leishmania promastigotes by DNA:DNA hybridization with kinetoplast DNA cloned in E. coli plasmids. Am. J. Trop. Med. Hyg. 34, 257–265.CrossRefGoogle ScholarPubMed
Le Blancq, S. M., Schnur, L. F. and Peters, W. (1986) Leishmania in the Old World: 1. The geographical and hostal distribution of L. major zymodemes. Trans. R. Soc. Trop. Med. Hyg. 80, 99–112.Google Scholar
Lopez, U. G. and Wirth, D. F. (1986) Identification of visceral Leishmania species with cloned sequences of kinetoplast DNA. Mol. Biochem. Parasitol. 20, 77–84.CrossRefGoogle Scholar
Mackinnon, J. A. and Fendal, N. R. E. (1955) Kala-azar in Baringo District of Kenya. J. Trop. Med. Hyg. 58, 205–209.Google Scholar
Maniatis, T., Fritsch, E. F. and Sambrook, J. (1982) Molecular Cloning: a Laboratory Manual. Cold Spring Harbor Laboratory, New York.Google Scholar
McMahon-Pratt, D., Bennett, E. and David, J. R. (1982) Monoclonal antibodies that distinguish subspecies of Leishmania braziliensis. J. Immunol. 129, 926–927.CrossRefGoogle ScholarPubMed
Mebrahtu, Y., Oster, C.N., Sharty, A.M., Hendricks, L. D., Githure, J. I., Rees, P. H., Perkins, P.V., Leeuwenburg, J. (1987) Cutaneous leishmaniasis caused by Leishmania tropica in Kenya. Trans. R. Soc. Trop. Med. Hyg. 81, 923–924.CrossRefGoogle ScholarPubMed
Mebrahtu, Y., Lawyer, P., Githure, J., Kager, P., Leeuwenburg, J., Perkins, P., Oster, C. and Hendricks, L. (1988) Indigenous human cutaneous leishmaniasis caused by Leishmania tropica in Kenya. Am. J. Trop. Med. Hyg. 39, 267–273.CrossRefGoogle ScholarPubMed
Mebrahtu, Y., Lawyer, P., Githure, J., Were, J. B. O., Muigai, R., Hendricks, L., Leeuwenburg, J., Koech, D. and Roberts, C. (1989) Visceral leishmaniasis unresponsive to pentostam caused by Leishmania tropica in Kenya. Am. J. Trop. Med. Hyg. 41, 289–294.Google Scholar
Mebrahtu, Y., Lawyer, P., Hendricks, L., Oster, C., Perkins, P., Koech, D., Pamba, H. and Roberts, C. (1990) Concurrent infection with Leishmania donovani and Leishmania major in a Kenyan patient. Am. J. Trop. Med. Hyg. In press.Google Scholar
Miles, M. A., Povoa, M. M., De Souza, A. A., Lainson, R. and Shaw, J. J. (1980) Some methods for enzymic characterization of Latin-American Leishmania with particular reference to Leishmania mexicana amazonensis and subspecies of Leishmania hertigi. Trans. R. Soc. Trop. Med. Hyg. 74, 243–252.Google Scholar
Minter, D. M. and Wijers, J. B., Heisch, R. B. and Manson-Bahr, P. E. C. (1962) Phlebotomus martini — a probable vector of kala-azar in Kenya. Brit. Med. J. 2, 285.CrossRefGoogle Scholar
Muigai, R., Githure, J. I., Gachihi, G. S., Were, J. B. O., Leeuwenburg, J. and Perkins, P. V. (1987) Cutaneous leishmaniasis caused by Leishmania major in Baringo District, Kenya. Trans. R. Soc. Trop. Med. Hyg. 81, 600–602.Google Scholar
Mutinga, M.J. and Ngoka, J.M. (1970) Culture isolation and description of cutaneous leishmaniasis in Kenya. Proc. E. Afr. Med. Res. Council 4, 72–74.Google Scholar
Mutinga, M. J. and Ngoka, J. M. (1978) Incrimination of the vector of visceral leishmaniasis in Kenya. East Afr. Med. J. 55, 337–340.Google ScholarPubMed
Mutinga, M. J. (1986) Epidemiology of leishmaniases in Kenya. Advances in research of vectors and animal reservoirs and possible control measures. Insect Sci. Applic. 7, 199–206.Google Scholar
Mutinga, M. J. and Odhiambo, T. R. (1986) Cutaneous leishmaniasis in Kenya — II. Studies on vector potential of Phlebotomies pedifer (Diptera:Phlebotominae) in Kenya. Insect Sci. Applic. 7, 171–174.Google Scholar
Mutinga, M. J., Mutero, C. M., Ngindu, A., Amimo, F. A. (1988) The isolation of leishmanial parasites from domestic goats and wild hosts and possible role of goats as reservoirs of leishmaniases. Insect Sci. Applic. 9, 339–344.Google Scholar
Mutinga, M. J., Kihara, S. M., Lohding, A., Mutero, C. M., Ngatia, T. A. and Karanu, F. (1989) Leishmaniasis in Kenya: description of leishmaniasis of a domestic goat from Transmara, Narok District, Kenya. Trop. Med. Parasitol. 40, 91–96.Google Scholar
Okot-Kotber, B. M. (1985) A rapid chromatographic method for elimination of fungal contamination in in vitro cultures of Leishmania spp. Parasitology 91, 1–7.CrossRefGoogle ScholarPubMed
Okot-Kotber, B. M., Mutinga, M. J. and Kaddu, J. B. (1989) Biochemical characterization of Leishmania spp. isolated from man and wild animals in Kenya. Int. J. Parasitol. 19, 657–663.CrossRefGoogle ScholarPubMed
Rigby, P. W. J., Dickmann, M., Rhodes, C. and Berg, P. (1977) Labelling deoxyribonucleic acid to high specific activity in vitro nick translation with DNA polymerase I. J. Mol. Biol. 113, 237–251.CrossRefGoogle ScholarPubMed
Riou, G. and Detain, E. (1969) Electron microscopy of the circular kinetoplast DNA from Trypanosoma cruzi: occurrence of catenated forms. Biochemistry 62, 210–217.Google Scholar
Rogers, W. O., Bumheim, P. F. and Wirth, D. F. (1988) Detection of Leishmania within sandflies by kinetoplast DNA hybridization. Am. J. Trop. Med. Hyg. 39, 434–139.Google Scholar
Rogers, W. O. and Wirth, D. F. (1988) Generation of sequence diversity in the kinetoplast DNA minicircles of Leishmania mexicana amazonensis. Mol. Biochem. Parasitol. 30, 1–8.Google Scholar
Schottelius, J. (1982) Lectin binding strain-specific carbohydrates on the cell surfaces of Leishmania strains from the Old World. Z. Parasitenk. 66, 237–247.CrossRefGoogle Scholar
Southern, E. M. (1975) Detection of specific sequences among DNA fragments separated by gel electrophoresis. J. Mol. Biol. 98, 501–517.CrossRefGoogle ScholarPubMed
Spithill, T. W. and Grumont, R. J. (1984) Identification of species, strains and clones of Leishmania by characterization of kinetoplast DNA minicircles. Mol. Biochem. Parasitol. 12, 217–236.Google Scholar
Van Eys, G. J. J. M., Schoone, G. J., Ligthart, G. S., Alvar, J., Evans, D. A. and Terpstra, W. J. (1989) Identification of “Old World” Leishmania by DNA recombinant probes. Mol. Biochem. Parasitol. 34, 53–62.Google Scholar
Wahl, G. M., Stern, M. and Stark, G. R. (1979) Efficient transfer of large DNA fragments from agarose gels to diazobenzyl oxymethyl-paper and rapid hybridization using dextran sulfate. Proc. Natl. Acad. Sci. (USA) 76, 3683–3687.Google Scholar
Wirth, D. F. and McMahon-Pratt, D. (1982) Rapid identification of Leishmania species by specific hybridization of kinetoplast DNA in cutaneous lesions. Proc. Natl. Acad. Sci. (USA) 79, 6999–7003.CrossRefGoogle ScholarPubMed