Eradication of Diseases
Human Genetics: Concepts And Application
Because of natural selection, different alleles are more likely to confer a survival advantage in different environments. Cycles of infectious disease prevalence and virulence often reflect natural selection.
Balanced PolymorphismIf natural selection eliminates individuals with detrimental phenotypes from a population, then why do harmful mutant alleles persist in a gene pool? A disease can remain prevalent when heterozygotes have some other advantage over individuals who have two copies of the wild type allele. When carriers have advantages that allow a detrimental allele to persist in a population, balanced polymorphism is at work. This form of polymorphism often entails heterozygosity for an inherited illness that protects against an infectious illness. Examples are fascinating.
Sickle Cell DiseaseSickle Cell disease is an autosomal recessive disorder that causes anemia, joint pain, a swollen spleen, and frequent, severe infections. It illustrates balanced polymorphism because carriers are resistant to malaria, an infection by the parasite Plasmodium falciparum that causes cycles of chills and fever. The parasite spends the first stage of its life cycle in the salivary glands of the mosquito Anopheles gambiae. When an infected mosquito bites a human, the malaria parasite enters the red blood cells, which transport it to the liver. The red blood cells burst, releasing the parasite throughout the body.
In 1949, British geneticist Anthony Allison found that the frequency of sickle cell carriers in tropical Africa was higher in regions where malaria raged all year long. Blood tests from children hospitalized with malaria found that nearly all were homozygous for the wild type of sickle cell allele. The few sickle cell carriers among them had the mildest cases of malaria. Was the presence of malaria somehow selecting for the sickle cell allele by felling people who did not inherit it? The fact that sickle cell disease is far less common in the United States, where malaria is rare, supports the idea that sickle cell heterozygosity provides a protective effect.
Further evidence of a sickle cell carrier's advantage in a malaria-ridden environment is the fact that the rise of sickle cell disease parallels the cultivation of crops that provide breeding grounds for Anopheles mosquitoes. About 1,000 B.C., Malayo-Polynesian sailors from southeast Asia traveled in canoes to East Africa, bringing new crops of bananas, yams, taros, and coconuts. When the jungle was cleared to grow these crops, the open space provided breeding ground for mosquitoes. The insects, in turn, offered a habitat for part of the life cycle of the malaria parasite.
The sickle cell gene may have been brought to Africa by people migrating from Southern Arabia and India, or it may have arisen by mutation directly in East Africa. However it happened, people who inherited one copy of the sickle cell allele had red blood cell membranes that did not admit the parasite. Carriers had more children and passed the protective allele to approximately half of them. Gradually, the frequency of the sickle cell allele in East Africa rose from 0.1 percent to a spectacular 45 percent in thirty-five generations. Carriers paid the price for this genetic protection, whenever two produced a child with sickle cell disease.
A cycle set in. Settlements with large numbers of sickle cell carriers escaped debilitating malaria. They were therefore strong enough to clear even more land to grow food- and support the disease-bearing mosquitoes. Even today, sickle cell disease is more prevalent in agricultural societies than among people who hunt and gather their food.
Glucose-6-Phosphate Dehydrogenase DeficiencyG6PD deficiency is a sex-linked enzyme deficiency that affects 400 million people worldwide. It causes life-threatening hemolytic anemia, in which red blood cells burst. However, it develops only under specific conditions- eating fava beans, inhaling certain types of pollen, taking certain drugs, or contracting certain infections. Studies on African children with severe malaria show that heterozygous females and hemizygous males for G6PD deficiency are underrepresented. This suggests that inheriting the enzyme deficiency gene somehow protects against malaria.
The fact that G6PD deficiency is sex-linked introduces a possibility we do not see with sickle cell disease, which is autosomal recessive. Because both heterozygotes and hemizygotes are selected for, the mutant allele should eventually predominate in a malaria-exposed population. However, this doesn't happen- there are still males hemizygous and females homozygous for the wild type allele. The reason again relates to natural selection. People with the enzyme deficiency- hemizygous males and homozygous females- are selected out of the population by the anemia. Therefore, natural selection acts in two directions on the hemizygous males- selecting for the mutant allele because it protects against malarial infection, yet selecting against it because an enzyme deficiency. This is the essence of balanced polymorphism.
PKUPhenylketnonuria is an inborn error of metabolism in which a missing enzyme causes the amino acid phenylalanine to build up, with devastating effects on the nervous system unless the individual follows a restrictive diet. Carriers of this autosomal recessive condition have elevated phenylalanine levels- levels that are not sufficiently high to cause symptoms, but that are high enough that they may have a protective effect during pregnancy. Physicians have observed that women who are PKU carriers have a much lower-than�average incidence of miscarriage. One theory is that excess phenylalanine somehow inactivates a poison, called ochratoxin A, that certain fungi produce and that is known to cause spontaneous abortion.
History provides the evidence that links PKU heterozygosity to protection against a fungal toxin. PKU is most common in Ireland and western Scotland, and many affected families living elsewhere trace their roots to this part of the world. If PKU carriers were most likely to have children than non-carriers because of the protective effects of the PKU gene, over time, the disease-causing allele would increase the population.
Tay-Sachs DiseaseCarrying Tay-Sachs disease may protect against tuberculosis (TB). In Ashkenazim populations, up to 11 percent of the people are Tay-Sachs carriers. During World War II, TB ran rampant in Eastern European Jewish settlements. Often, healthy relatives of children with Tay-Sachs disease did not contact TB, even when repeatedly exposed. The protection against TB that Tay-Sachs disease heterozygosity apparently offered remained among the Jewish people because they were prevented from leaving the ghettos. The mutant allele increased in frequency as TB selectively felled those who did not carry it and the carriers had children with each other. Genetic drift may also have helped isolate the Tay-Sachs allele, by chance, in groups of holocaust survivors. Precisely how lowered levels of the gene product, an enzyme called hexoseaminidase A, protect against TB isn't known.
Cystic FibrosisBalanced polymorphism may explain why cystic fibrosis is so common- the anatomical defect that underlies CF protects against diarrheal illnesses, such as cholera.
Cholera epidemics have left their mark on human populations, causing widespread death in just days. In the summer of 1831, an epidemic killed 10 percent of the population of St. Louis, and in 1991, an epidemic swept Peru. Cholera bacteria causes diarrhea, which rapidly dehydrates the body and can lead to shock and kidney and heart failure. The bacterium produces a toxin that opens chloride channels in the small intestine. As salt (NaCl) leaves the cells, water follows, in a natural chemical tendency to dilute the salt. Water rushing out of intestinal cells leaves the body as diarrhea.
In 1989, when geneticists identified the CF gene and described its protein product as a regulator of a chloride channel in certain secretory cells, a possible explanation for the prevalence of the inherited disorder emerged. Cholera opens chloride channels, letting chloride and water leave cells. The CFTR protein does just the opposite, closing chloride channels and trapping salt and water in cells, which dries out mucus and other secretions. A person with CF cannot contract cholera, because the toxin cannot open the chloride channels in the small intestine.
Carriers of CF enjoy the mixed blessing of a balanced polymorphism. They do not have enough abnormal chloride channels to cause the labored breathing and clogged pancreas of cystic fibrosis, but they do have enough of a defect to prevent the cholera from taking hold. During the devastating cholera epidemics that have peppered history, individuals carrying mutant CF alleles had a selective advantage, and they disproportionately transmitted those alleles to future generations. However, because CF arose in Western Europe and cholera in Africa, perhaps an initial increase in CF herterozygosity was a response to a different diarrheal infection.
Malaria And The Liver: Exploring The Silent Pathway
Cite this articleO'Connell, D. Malaria and the liver: exploring the silent pathway. Nat Rev Microbiol 5, 909 (2007). Https://doi.Org/10.1038/nrmicro1809
Share this article Get shareable linkGenetic Linkage And Association Analyses For Trait Mapping In Plasmodium Falciparum
Levine, N. D. Progress in taxonomy of the Apicomplexan protozoa. J. Protozool. 35, 518–520 (1988).
Bruce-Chwatt, L. J. Essential Malariology (Oxford Univ. Press, New York, 1993).
Singh, B. Et al. A large focus of naturally acquired Plasmodium knowlesi infections in human beings. Lancet 363, 1017–1024 (2004).
World Health Organization. The World Health Report 2005: Make Every Mother and Child Count. World Health Organization, Geneva. [online], (2005).
Trager, W. & Jensen, J. B. Human malaria parasites in continuous culture. Science 193, 673–675 (1976). A landmark paper describing the methods for in vitro culture of P. Falciparum.
Vanderberg, J. P. & Gwadz, R. W. In Malaria, Volume 2: Pathology, Vector Studies and Culture. (ed. Kreier, J. P.) 153–234 (Academic, New York, 1980).
Walliker, D. Et al. Genetic analysis of the human malaria parasite Plasmodium falciparum. Science 236, 1661–1666 (1987). Description of the first genetic cross in P. Falciparum.
Peterson, D. S., Walliker, D. & Wellems, T. E. Evidence that a point mutation in dihydrofolate reductase-thymidylate synthase confers resistance to pyrimethamine in falciparum malaria. Proc. Natl Acad. Sci. USA 85, 9114–9118 (1988).
Wellems, T. E. Et al. Chloroquine resistance not linked to mdr-like genes in a Plasmodium falciparum cross. Nature 345, 253–255 (1990).
Wellems, T. E., Walker-Jonah, A. & Panton, L. J. Genetic mapping of the chloroquine-resistance locus on Plasmodium falciparum chromosome 7. Proc. Natl Acad. Sci. USA 88, 3382–3386 (1991).
Vaidya, A. B. Et al. A genetic locus on Plasmodium falciparum chromosome 12 linked to a defect in mosquito infectivity and male gametogenesis. Mol. Biochem. Parasitol. 69, 65–71 (1995).
Su, X.-Z., Kirkman, L. A., Fujioka, H. & Wellems, T. E. Complex polymorphisms in an ∼330 kDa protein are linked to chloroquine-resistant P. Falciparum in Southeast Asia and Africa. Cell 91, 593–603 (1997).
Wang, P., Read, M., Sims, P. F. G. & Hyde, J. E. Sulfadoxine resistance in the human malaria parasite Plasmodium falciparum is determined by mutations in the dihydropteroate synthase and an additional factor associated with folate utilization. Mol. Microbiol. 23, 979–986 (1997).
Su, X.-Z. Et al. A genetic map and recombination parameters of the human malaria parasite Plasmodium falciparum. Science 286, 1351–1353 (1999). Description of the first high-resolution genetic map for P. Falciparum.
Fidock, D. A. Et al. Mutations in the P. Falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Mol. Cell 6, 861–871 (2000). The original description of the determinant of CQ resistance in P. Falciparum.
Ferdig, M. T. Et al. Dissecting the loci of low-level quinine resistance in malaria parasites. Mol. Microbiol. 52, 985–997 (2004).
Wang, P., Nirmalan, N., Wang, Q., Sims, P. F. & Hyde, J. E. Genetic and metabolic analysis of folate salvage in the human malaria parasite Plasmodium falciparum. Mol. Biochem. Parasitol. 135, 77–87 (2004).
Furuya, T. Et al. Disruption of a Plasmodium falciparum gene linked to male sexual development causes early arrest in gametocytogenesis. Proc. Natl Acad. Sci. USA 102, 16813–16818 (2005).
Hoffman, S. L., Subramanian, G. M., Collins, F. H. & Venter, J. C. Plasmodium, human and Anopheles genomics and malaria. Nature 415, 702–709 (2002).
Crabb, B. S. Transfection technology and the study of drug resistance in the malaria parasite Plasmodium falciparum. Drug Resist. Updat. 5, 126–130 (2002).
Bozdech, Z. Et al. The transcriptome of the intraerythrocytic developmental cycle of Plasmodium falciparum. PLoS Biol. 1, e5 (2003).
Le Roch, K. G. Et al. Discovery of gene function by expression profiling of the malaria parasite life cycle. Science 301, 1503–1508 (2003).
Yeh, I., Hanekamp, T., Tsoka, S., Karp, P. D. & Altman, R. B. Computational analysis of Plasmodium falciparum metabolism: organizing genomic information to facilitate drug discovery. Genome Res. 14, 917–924 (2004).
Carlton, J., Silva, J. & Hall, N. The genome of model malaria parasites, and comparative genomics. Curr. Issues Mol. Biol. 7, 23–37 (2005).
Hall, N. Et al. A comprehensive survey of the Plasmodium life cycle by genomic, transcriptomic, and proteomic analyses. Science 307, 82–86 (2005).
LaCount, D. J. Et al. A protein interaction network of the malaria parasite Plasmodium falciparum. Nature 438, 103–107 (2005).
Stoeckert, C. J. Jr et al. PlasmoDB v5: new looks, new genomes. Trends Parasitol. 22, 543–546 (2006).
Winzeler, E. A. Applied systems biology and malaria. Nature Rev. Microbiol. 4, 145–151 (2006).
Walliker, D., Carter, R. & Morgan, S. Genetic recombination in malaria parasites. Nature 232, 561–562 (1971).
Carlton, J., Mackinnon, M. & Walliker, D. A chloroquine resistance locus in the rodent malaria parasite Plasmodium chabaudi. Mol. Biochem. Parasitol. 93, 57–72 (1998).
Hayton, K., Ranford-Cartwright, L. C. & Walliker, D. Sulfadoxine-pyrimethamine resistance in the rodent malaria parasite Plasmodium chabaudi. Antimicrob. Agents Chemother. 46, 2482–2489 (2002).
Cravo, P. V. Et al. Genetics of mefloquine resistance in the rodent malaria parasite Plasmodium chabaudi. Antimicrob. Agents Chemother. 47, 709–718 (2003).
Culleton, R., Martinelli, A., Hunt, P. & Carter, R. Linkage group selection: rapid gene discovery in malaria parasites. Genome Res. 15, 92–97 (2005).
Martinelli, A. Et al. A genetic approach to the de novo identification of targets of strain-specific immunity in malaria parasites. Proc. Natl Acad. Sci. USA 102, 814–819 (2005). This paper describes application of linkage group selection to identify targets of strain-specific immunity.
van Dijk, M. R., Waters, A. P. & Janse, C. J. Stable transfection of malaria parasite blood stages. Science 268, 1358–1362 (1995).
de Koning-Ward, T. F., Janse, C. J. & Waters, A. P. The development of genetic tools for dissecting the biology of malaria parasites. Annu. Rev. Microbiol. 54, 157–185 (2000).
Balu, B. & Adams, J. H. Advancements in transfection technologies for Plasmodium. Int. J. Parasitol. 37, 1–10 (2007).
Carlton, J. M., Hayton, K., Cravo, P. V. & Walliker, D. Of mice and malaria mutants: unravelling the genetics of drug resistance using rodent malaria models. Trends Parasitol. 17, 236–242 (2001).
Carter, R., Hunt, P. & Cheesman, S. Linkage group selection — a fast approach to the genetic analysis of malaria parasites. Int. J. Parasitol. 37, 285–293 (2007).
Sinden, R. E. & Hartley, R. H. Identification of the meiotic division of malarial parasites. J. Protozool. 32, 742–744 (1985).
Babiker, H. A. Et al. Random mating in a natural population of the malarial parasite Plasmodium falciparum. Parasitology 109, 413–421 (1994).
Paul, R. E. L. Et al. Mating patterns in malaria parasite populations of Papua New Guinea. Science 269, 1709–1711 (1995).
Wellems, T. E. & Plowe, C. V. Chloroquine-resistant malaria. J. Infect. Dis. 184, 770–776 (2001).
Carter, R. & Mendis, K. N. Evolutionary and historical aspects of the burden of malaria. Clin. Microbiol. Rev. 15, 564–594 (2002).
Chou, A. C., Chevli, R. & Fitch, C. D. Ferriprotoporphyrin IX fulfills the criteria for identification as the chloroquine receptor of malaria parasites. Biochemistry 19, 1543–1549 (1980).
Fidock, D. A. Et al. Allelic modification of the cg2 and cg1genes do not alter the chloroquine response of drug-resistant Plasmodium falciparum. Mol. Biochem. Parasitol. 110, 1–10 (2000).
Cooper, R. A. Et al. Alternative mutations at position 76 of the vacuolar transmembrane protein PfCRT are associated with chloroquine resistance and unique stereospecific quinine and quinidine responses in Plasmodium falciparum. Mol. Pharmacol. 61, 35–42 (2002).
Sidhu, A. B., Verdier-Pinard, D. & Fidock, D. A. Chloroquine resistance in Plasmodium falciparum malaria parasites conferred by pfcrt mutations. Science 298, 210–213 (2002).
Djimde, A. Et al. A molecular marker for chloroquine-resistant falciparum malaria. N. Engl. J. Med. 344, 257–263 (2001).
Wootton, J. C. Et al. Genetic diversity and chloroquine selective sweeps in Plasmodium falciparum. Nature 418, 320–323 (2002). Description of CQ-resistance founder events and subsequent selective sweeps across the major malarious regions of the world.
Mehlotra, R. K. Et al. Evolution of a unique Plasmodium falciparum chloroquine-resistance phenotype in association with PfCRT polymorphism in Papua New Guinea and South America. Proc. Natl Acad. Sci. USA 98, 12689–12694 (2001).
Bray, P. G. Et al. PfCRT and the trans-vacuolar proton electrochemical gradient: regulating the access of chloroquine to ferriprotoporphyrin IX. Mol. Microbiol. 62, 238–251 (2006).
Cooper, R. A. Et al. Mutations in transmembrane domains 1, 4 and 9 of the Plasmodium falciparum chloroquine resistance transporter alter susceptibility to chloroquine, quinine and quinidine. Mol. Microbiol. 63, 270–282 (2007).
Mu, J. Et al. Multiple transporters associated with malaria parasite responses to chloroquine and quinine. Mol. Microbiol. 49, 977–989 (2003).
Wongsrichanalai, C. Et al. In vitro susceptibility of Plasmodium falciparum isolates in Vietnam to artemisinin derivatives and other antimalarials. Acta Trop. 63, 151–158 (1997).
Reed, M. B., Saliba, K. J., Caruana, S. R., Kirk, K. & Cowman, A. F. Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum. Nature 403, 906–909 (2000).
Kidgell, C. Et al. A systematic map of genetic variation in Plasmodium falciparum. PLoS. Pathog. 2, e57 (2006). A primary report of allelic variability in the P. Falciparum genome detected by high-density microarrays.
Farrall, M. Quantitative genetic variation: a post-modern view. Hum. Mol. Genet. 13, R1–R7 (2004).
Suthram, S., Sittler, T. & Ideker, T. The Plasmodium protein network diverges from those of other eukaryotes. Nature 438, 108–112 (2005).
Nair, S. Et al. A selective sweep driven by pyrimethamine treatment in Southeast Asian malaria parasites. Mol. Biol. Evol. 20, 1526–1536 (2003).
Roper, C. Et al. Antifolate antimalarial resistance in southeast Africa: a population-based analysis. Lancet 361, 1174–1181 (2003).
Nair, S. Et al. Recurrent gene amplification and soft selective sweeps during evolution of multidrug resistance in malaria parasites. Mol. Biol. Evol. 24, 562–573 (2007).
Al-Olayan, E. M., Beetsma, A. L., Butcher, G. A., Sinden, R. E. & Hurd, H. Complete development of mosquito phases of the malaria parasite in vitro. Science 295, 677–679 (2002).
Hollingdale, M. R. In In vitro Methods for Parasite Cultivation (eds Taylor, A. E. R. & Baker, J. R.) 180–198 (Academic, New York, 1987).
Udomsangpetch, R. Et al. Short-term in vitro culture of field isolates of Plasmodium vivax using umbilical cord blood. Parasitol. Int. 56, 65–69 (2006).
Kocken, C. H. Et al. Plasmodium knowlesi provides a rapid in vitro and in vivo transfection system that enables double-crossover gene knockout studies. Infect. Immun. 70, 655–660 (2002).
Rich, S. M., Licht, M. C., Hudson, R. R. & Ayala, F. J. Malaria's Eve: evidence of a recent population bottleneck throughout the world populations of Plasmodium falciparum. Proc. Natl Acad. Sci. USA 95, 4425–4430 (1998).
Volkman, S. K. Et al. Recent origin of Plasmodium falciparum from a single progenitor. Science 293, 482–484 (2001).
Hughes, A. L. & Verra, F. Very large long-term effective population size in the virulent human malaria parasite Plasmodium falciparum. Proc. Biol. Sci. 268, 1855–1860 (2001).
Mu, J. Et al. Chromosome-wide SNPs reveal an ancient origin for Plasmodium falciparum. Nature 418, 323–326 (2002).
Joy, D. A. Et al. Early origin and recent expansion of Plasmodium falciparum. Science 300, 318–321 (2003).
Su, X. Z., Mu, J. & Joy, D. A. The 'Malaria's Eve' hypothesis and the debate concerning the origin of the human malaria parasite Plasmodium falciparum. Microbes. Infect. 5, 891–896 (2003).
Jeffares, D. C. Et al. Genome variation and evolution of the malaria parasite Plasmodium falciparum. Nature Genet. 39, 120–125 (2007). Description of genome-wide polymorphism within and between species, and evolutionary implications of the findings.
Mu, J. Et al. Genome-wide variation and identification of vaccine targets in the Plasmodium falciparum genome. Nature Genet. 39, 126–130 (2007). Selection signatures from a survey of ∼3,500 predicted genes identify sequences that are likely to be under immune pressure.
Volkman, S. K. Et al. A genome-wide map of diversity in Plasmodium falciparum. Nature Genet. 39, 113–119 (2007). SNPs identified by large-scale sequencing reveal genome-wide LD, selection and recombination
Hughes, A. L. & Verra, F. Extensive polymorphism and ancient origin of Plasmodium falciparum. Trends Parasitol. 18, 348–351 (2002).
Anderson, T. J. Et al. Microsatellite markers reveal a spectrum of population structures in the malaria parasite Plasmodium falciparum. Mol. Biol. Evol. 17, 1467–1482 (2000).
Mu, J. Et al. Recombination hotspots and population structure in Plasmodium falciparum. PLoS Biol. 3, e335 (2005).
Hayward, R. E. Et al. Shotgun DNA microarrays and stage-specific gene expression in Plasmodium falciparum malaria. Mol. Microbiol. 35, 6–14 (2000).
Llinas, M., Bozdech, Z., Wong, E. D., Adai, A. T. & DeRisi, J. L. Comparative whole genome transcriptome analysis of three Plasmodium falciparum strains. Nucleic Acids Res. 34, 1166–1173 (2006).
Young, J. A. Et al. The Plasmodium falciparum sexual development transcriptome: a microarray analysis using ontology-based pattern identification. Mol. Biochem. Parasitol. 143, 67–79 (2005).
Mok, B. W. Et al. Comparative transcriptomal analysis of isogenic Plasmodium falciparum clones of distinct antigenic and adhesive phenotypes. Mol. Biochem. Parasitol. 151, 184–192 (2007).
Lovegrove, F. E. Et al. Simultaneous host and parasite expression profiling identifies tissue-specific transcriptional programs associated with susceptibility or resistance to experimental cerebral malaria. BMC Genomics 7, 295 (2006).
Mockler, T. C. Et al. Applications of DNA tiling arrays for whole-genome analysis. Genomics 85, 1–15 (2005).
Price, R. N. Et al. Mefloquine resistance in Plasmodium falciparum and increased pfmdr1 gene copy number. Lancet 364, 438–447 (2004).
Anderson, T. J. Et al. Are transporter genes other than the chloroquine resistance locus (pfcrt) and multidrug resistance gene (pfmdr) associated with antimalarial drug resistance? Antimicrob. Agents Chemother. 49, 2180–2188 (2005).
Hill, W. G., Babiker, H. A., Ranford-Cartwright, L. C. & Walliker, D. Estimation of inbreeding coefficients from genotypic data on multiple alleles, and application to estimation of clonality in malaria parasites. Genet. Res. 65, 53–61 (1995).
Walliker, D., Babiker, H. A. & Ranford-Cartwright, L. C. The genetic structure of malaria parasite populations in Malaria: Parasite Biology, Pathogenesis and Protection (ed. Sherman, I. W.) 235–255 (ASM, Washington DC, 1998).
Conway, D. J. Et al. High recombination rate in natural populations of Plasmodium falciparum. Proc. Natl Acad. Sci. USA 96, 4506–4511 (1999). Description and discussion of high recombination frequencies in parasite populations of five African countries.
Laserson, K. F. Et al. Genetic characterization of an epidemic of Plasmodium falciparum malaria among Yanomami Amerindians. J. Infect. Dis. 180, 2081–2085 (1999).
Florens, L. Et al. A proteomic view of the Plasmodium falciparum life cycle. Nature 419, 520–526 (2002).
Lasonder, E. Et al. Analysis of the Plasmodium falciparum proteome by high-accuracy mass spectrometry. Nature 419, 537–542 (2002).
Khan, S. M. Et al. Proteome analysis of separated male and female gametocytes reveals novel sex-specific Plasmodium biology. Cell 121, 675–687 (2005).
Sam-Yellowe, T. Y. Et al. Proteome analysis of rhoptry-enriched fractions isolated from Plasmodium merozoites. J. Proteome. Res. 3, 995–1001 (2004).
Ginsburg, H. Progress in in silico functional genomics: the malaria Metabolic Pathways database. Trends Parasitol. 22, 238–240 (2006).
Sinden, R. E. A proteomic analysis of malaria biology: integration of old literature and new technologies. Int. J. Parasitol. 34, 1441–1450 (2004).
Carlton, J. M. Et al. Genome sequence and comparative analysis of the model rodent malaria parasite Plasmodium yoelii yoelii. Nature 419, 512–519 (2002).
Aravind, L., Iyer, L. M., Wellems, T. E. & Miller, L. H. Plasmodium biology: genomic gleanings. Cell 115, 771–785 (2003).
Trager, W. & Williams, J. Extracellular (axenic) development in vitro of the erythrocytic cycle of Plasmodium falciparum. Proc. Natl Acad. Sci. USA 89, 5351–5355 (1992).
Wu, Y., Sifri, C. D., Lei, H. H., Su, X. Z. & Wellems, T. E. Transfection of Plasmodium falciparum within human red blood cells. Proc. Natl Acad. Sci. USA 92, 973–977 (1995).
Duraisingh, M. T., Triglia, T. & Cowman, A. F. Negative selection of Plasmodium falciparum reveals targeted gene deletion by double crossover recombination. Int. J. Parasitol. 32, 81–89 (2002).
Deitsch, K., Driskill, C. & Wellems, T. Transformation of malaria parasites by the spontaneous uptake and expression of DNA from human erythrocytes. Nucleic Acids Res. 29, 850–853 (2001).
Carvalho, T. G. & Menard, R. Manipulating the Plasmodium genome. Curr. Issues Mol. Biol. 7, 39–55 (2005).
Meissner, M. Et al. Tetracycline analogue-regulated transgene expression in Plasmodium falciparum blood stages using Toxoplasma gondii transactivators. Proc. Natl Acad. Sci. USA 102, 2980–2985 (2005).
Mamoun, C. B., Gluzman, I. Y., Goyard, S., Beverley, S. M. & Goldberg, D. E. A set of independent selectable markers for transfection of the human malaria parasite Plasmodium falciparum. Proc. Natl Acad. Sci. USA 96, 8716–8720 (1999).
Desjardins, R. E., Canfield, C. J., Haynes, J. D. & Chulay, J. D. Quantitative assessment of antimalarial activity in vitro by a semiautomated technique. Antimicrobial Agents Chemother. 16, 710–718 (1979).
White, N. J. Assessment of the pharmacodynamic properties of antimalarial drugs in vivo. Antimicrob. Agents Chemother. 41, 1413–1422 (1997).
Miller, L. H., Baruch, D. I., Marsh, K. & Doumbo, O. K. The pathogenic basis of malaria. Nature 415, 673–679 (2002).
Gaur, D., Mayer, D. C. & Miller, L. H. Parasite ligand-host receptor interactions during invasion of erythrocytes by Plasmodium merozoites. Int. J. Parasitol. 34, 1413–1429 (2004).
Cowman, A. F. & Crabb, B. S. Invasion of red blood cells by malaria parasites. Cell 124, 755–766 (2006).
Miller, L. H., Mason, S. J., Dvorak, J. A., McGinniss, M. H. & Rothman, I. K. Erythrocyte receptors for (Plasmodium knowlesi) malaria: Duffy blood group determinants. Science 189, 561–563 (1975).
al-Khedery, B., Barnwell, J. W. & Galinski, M. R. Antigenic variation in malaria: a 3′ genomic alteration associated with the expression of a P. Knowlesi variant antigen. Mol. Cell 3, 131–141 (1999).
Dzikowski, R., Frank, M. & Deitsch, K. Mutually exclusive expression of virulence genes by malaria parasites is regulated independently of antigen production. PLoS. Pathog. 2, e22 (2006).
Kaestli, M. Et al. Virulence of malaria is associated with differential expression of Plasmodium falciparum var gene subgroups in a case-control study. J. Infect. Dis. 193, 1567–1574 (2006).
Trimnell, A. R. Et al. Global genetic diversity and evolution of var genes associated with placental and severe childhood malaria. Mol. Biochem. Parasitol. 148, 169–180 (2006).
Shelburne, S. A. & Musser, J. M. Virulence gene expression in vivo. Curr. Opin. Microbiol. 7, 283–289 (2004).
Shortt, H. E., Fairley, N. H., Covell, G., Shute, P. G. & Garnham, P. C. C. The pre-erythrocytic stage of Plasmodium falciparum. Trans. R. Soc. Trop. Med. Hyg. 44, 405–419 (1951).
Shi, Q. Et al. Alteration in host cell tropism limits the efficacy of immunization with a surface protein of malaria merozoites. Infect. Immun. 73, 6363–6371 (2005).
Popular Posts
UKHSA Advisory Board: preparedness for infectious disease threats
Comments
Post a Comment