• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of geneticsGeneticsCurrent IssueInformation for AuthorsEditorial BoardSubscribeSubmit a Manuscript
Genetics. May 2006; 173(1): 235–241.
PMCID: PMC1461423

Quantitative Trait Loci Controlling Refractoriness to Plasmodium falciparum in Natural Anopheles gambiae Mosquitoes From a Malaria-Endemic Region in Western Kenya

Abstract

Natural anopheline populations exhibit much variation in ability to support malaria parasite development, but the genetic mechanisms underlying this variation are not clear. Previous studies in Mali, West Africa, identified two quantitative trait loci (QTL) in Anopheles gambiae mosquitoes that confer refractoriness (failure of oocyst development in mosquito midguts) to natural Plasmodium falciparum parasites. We hypothesize that new QTL may be involved in mosquito refractoriness to malaria parasites and that the frequency of natural refractoriness genotypes may be higher in the basin region of Lake Victoria, East Africa, where malaria transmission intensity and parasite genetic diversity are among the highest in the world. Using field-derived F2 isofemale families and microsatellite marker genotyping, two loci significantly affecting oocyst density were identified: one on chromosome 2 between markers AG2H135 and AG2H603 and the second on chromosome 3 near marker AG3H93. The first locus was detected in three of the five isofemale families studied and colocalized to the same region as Pen3 and pfin1 described in other studies. The second locus was detected in two of the five isofemale families, and it appears to be a new QTL. QTL on chromosome 2 showed significant additive effects while those on chromosome 3 exhibited significant dominant effects. Identification of P. falciparum-refractoriness QTL in natural An. gambiae mosquitoes is critical to the identification of the genes involved in malaria parasite transmission in nature and for understanding the coevolution between malaria parasites and mosquito vectors.

MALARIA is the most important parasitic disease in the world. It continues to place tremendous health and economic constraints on a large percentage of the world's population despite efforts to contain it; hence, there is a tremendous need to develop new means of control (World Health Organization 1998; Breman et al. 2004; Snow et al. 2005). Among the novel approaches being pursued is the proposed genetic manipulation of the malaria vector to disrupt parasite transmission (Curtis 1994; Aultmann et al. 2001) by taking advantage of observed natural refractoriness. Although natural refractoriness is not a new observation (Vargas 1949), its underlying mechanisms remain unknown (Vernick et al. 2005). The realization that the immune system of mosquitoes has the potential to kill malaria parasites at several developmental stages (Richman and Kafatos 1996) has inspired several studies aimed at unraveling the molecular mechanisms of the mosquito's immune responses against malaria parasites. Previous studies have identified a number of molecules inhibiting malaria parasite development in mosquitoes, including antimicrobial peptides (Gwadz et al. 1989; Lambrechts et al. 2004), signaling pathways and pattern recognition peptides (Barillas-Mury et al. 2000; Dimopoulos et al. 2001), monoclonal antibodies (De Capurro et al. 2000), and SM1 (Ito et al. 2002; Jacobs-Lorena 2003).

In parallel with efforts to understand the mosquito's immune mechanisms are genetic approaches aimed at identification and characterization of refractoriness. Such approaches have been used to define melanotic encapsulation of oocysts (Collins et al. 1986) and ookinete penetration failure (Vernick et al. 1995). Several quantitative trait loci (QTL) conferring encapsulation and melanization of malaria parasites and negatively charged sephadex beads in mosquitoes have been localized (Gorman et al. 1997; Zheng et al. 1997; Thomasova et al. 2002). Natural Anopheles gambiae populations in Africa do not normally show the encapsulation/melanization phenotype to Africa-origin Plasmodium falciparum; however, a substantial proportion of An. gambiae individuals in a population do not support successful parasite development in the midguts. Identification of a genetic mechanism of natural refractoriness (i.e., failure of ookinetes to develop into oocysts in mosquito midgut) is particularly interesting. Niare et al. (2002) used linkage analysis approaches and identified two genomic regions on chromosome 2 that confer An. gambiae mosquito natural refractoriness to P. falciparum parasites in Mali, West Africa.

Because of large genetic differentiation between West Africa and East Africa An. gambiae populations (Lehman et al. 2003), different P. falciparum-refractoriness QTL may be involved in the East Africa mosquito populations. In addition, malaria transmission in the western Kenya region is perennial, and the area experiences the highest malaria transmission intensity in the world (annual entomological inoculation rate in the order of hundreds) (Beier et al, 1999), while Mali exhibits seasonal malaria transmission with much lower intensity (Kleinschmidt et al. 2001). We hypothesize that the frequency of natural refractoriness genotypes is higher, as would be expected under a higher selection pressure by the malaria parasites (Schwartz and Koella 2001). The goal of this study is to determine whether new P. falciparum-refractoriness QTL, in addition to the QTL previously identified by Niare et al. (2002), can be detected and whether P. falciparum-refractoriness QTL are common in the western Kenya An. gambiae populations. Our study identified two QTL, one detected in three of the five isofemale families studied and colocalized to the same region as Pen3 and pfin1 described previously (Zheng et al. 1997; Niare et al. 2002); the second, located on chromosome 3 and detected in two of five isofemale families, appears to be a new QTL.

MATERIALS AND METHODS

Breeding mosquito pedigrees and experimental design:

The mosquitoes used in this study were obtained from gravid female An. gambiae individuals collected from Mbita, a village on the shores of Lake Victoria in western Kenya where high malaria prevalence and high transmission intensity were reported (Mutero et al. 1998; Shililu et al. 2003). Gravid wild-caught females were kept in the insectary at the Thomas Odhiambo Campus of the International Centre of Insect Physiology and Ecology in Mbita Point. Temperature and humidity in the insectary were not regulated. The mosquitoes were supplied with a 6% glucose solution on cotton wicks for two days. They were then put in individual oviposition cups, and eggs from each individual were reared separately to generate F1 isofemale families. Individuals of each F1 family were allowed to mate within their family, and females were given a blood meal to enable them to lay eggs. Eggs from each F1 family were reared together to give rise to F2 families. Mature 5- or 6-day-old F2 females were given infectious blood meals and then dissected 8 days postinfection to obtain the quantitative phenotype data. After oviposition, legs from each wild-caught mosquito were used to identify An. gambiae spp. by species identification PCR (Scott et al. 1993). The carcasses of the wild parents and the F1 females used to generate F2 families were saved for microsatellite genotyping. Because the first mating renders An. gambiae females refractory to subsequent insemination (Craig 1967; Gwadz et al. 1971), and the majority (>97%) of female An. gambiae mosquitoes mate only once in nature (Tripet et al. 2001), each of the F2 isofemale families is likely a product of a single pair cross.

Gametocyte carrier screening:

P. falciparum carriers were recruited from 2- to 10-year old children in the rural area around Mbita using a protocol approved by the institutional review board of the State University of New York at Buffalo and the ethical review board of the Kenya Medical Research Institute. Finger-prick blood samples were collected and thick blood smears were air dried, stained with 8% Giemsa for 15 min, and microscopically examined for the presence of P. falciparum trophozoites and gametocytes. The gametocyte densities were assessed by counting against 500 leukocytes and converted to number of parasites per microliter by assuming a standard leukocyte count of 8000/μl. Asymptomatic gametocyte-positive children were recruited as donors of infectious blood meals for mosquito membrane feeding.

Experimental infections and phenotyping:

Experimental infections took place in the afternoon following the morning finger-prick examinations. A clinician from Mbita Health Center drew 2 ml of venous blood into heparinized tubes prewarmed to 37°. Prewarming was necessary to guard against a lower temperature that might mimic temperatures in the mosquito gut and thus induce exflagellation of macro-gametocytes before they could be ingested by the mosquitoes. To reduce human factors such as transmission-blocking immunity (Mulder et al. 1994) the blood was centrifuged at 37° for 3 min at 2000 × g and the serum was replaced with the same volume of nonimmune AB serum purchased from Sigma. The mixture was used to feed 5- or 6-day-old mosquitoes that had been starved for 12–16 hr using parafilm membrane feeders that were prewarmed to 37° (Tchuinkam et al. 1993). Mosquitoes were allowed to feed for 30 min, after which the fed mosquitoes were placed into cages 15 × 15 × 15 cm3 in size. These mosquitoes were then maintained in the insectary on 6% glucose in cotton wicks that were changed daily to avoid bacterial contamination. Eight days after blood feeding, mosquitoes were dissected and their mid-guts were stained with 2% mercurochrome in distilled water to determine the number of oocysts with light microscopy. The number of oocysts in the midguts of individual mosquitoes constituted targeted phenotype.

Microsatellite genotyping and linkage analysis:

Genomic DNA was extracted individually from all the parents, F1, and F2 isofemale families following the phenol/chloroform method of Serverson (1997). DNA from wild-caught female parents used as isofemale family founders and DNA from each F1 pedigree were used to establish the segregation pattern of the markers used in molecular typing. Molecular typing used 20 polymorphic microsatellite markers selected from published linkage maps (Zheng et al. 1996; Wang et al. 1999) to cover the two autosomal chromosomes and the X chromosome for broad genome coverage. Microsatellite primers for PCR labeled with fluorescent M13 dye were obtained from Integrated DNA Technologies (Coralville, IA) and used to amplify microsatellite alleles according to the supplier's recommendations. PCR products were loaded and separated on polyacrylamide gels using a Li-Cor sequencer (Li-Cor). The size of microsatellite alleles was determined with Li-Cor GeneImagIR software and genotypes were assigned to individual mosquitoes on the basis of allele sizes.

Data analysis:

For each F2 isofemale family, mating occurred in nature and the fathers were not available for genotyping. Thus, paternity genotypes of the wild parents were inferred from allele separation patterns on polyacrylamide gels using multipoint analysis (Lander and Green 1987; Kruglyak et al. 1996; Markianos et al. 2001). Determination of paternal genotypes was necessary because linkage maps based on interval mapping require knowledge of the cis- or trans-relationships between alleles at different loci. Paternal reconstruction is straightforward for any family of >10 progeny, a condition met by all of the isofemale families in the study. Any families that were contaminated or that resulted from multiple matings were clearly evident because of the presence of aberrant nonparental genotypes, and they were discarded from the analysis. Analysis of the data to locate QTL for each family was done by least-squares interval mapping (Seaton et al. 2002) using QTL Express (http://qtl.cap.ed.ac.uk), with the thresholds for statistical significance determined by a permutation test (1000 permutations) for each chromosome. Confidence intervals for QTL location were obtained by bootstrapping (Visscher et al. 1996). For each isofemale family in which a significant QTL was detected, each of the three mosquito chromosomes was analyzed separately. Significant QTL were identified and the analyses were then repeated for each significant QTL while holding all other significant QTL as cofactors. This determines epistatic interactions between the significant QTL and any other loci on the whole genome. The data were also analyzed by maximum likelihood composite interval mapping (Wang et al. 2001–2005) using QTL Cartographer version 2.5 (http://statgen.ncsu.edu/qtlcart/WQTLCart.htm).

RESULTS

Experimental infections and phenotyping:

Five F2 isofemale families of An. gambiae were developed and successfully infected and genotyped (Table 1). The prevalence (percentage of infected mosquitoes) among the isofemale families varied from 19 to 53% whereas the mean oocyst counts varied from 4.60 to 11.26. Two families, coded 101 and 102, were developed and phenotyped during the short rainy season in October 2003. A t-test on the mean number of oocysts as a measure of parasite load in the two isofemale families gave a significant difference (P < 0.05). Because these two families were reared under identical environmental conditions and exposed to an infectious blood meal from the same gametocyte donor, the difference in oocyst density was caused by genetic variations between mosquito families. Three families, coded 201, 202, and 203, were developed and phenotyped during the long rainy season, May through June 2004. Of these three families, 202 and 203 were reared under identical environmental conditions and given an infectious blood meal from the same gametocyte donor. Similarly, t-test on the mean number of oocysts between the families gave a significant difference (P < 0.05), indicating significant genetic variation in mosquito susceptibility to malaria parasite infection.

TABLE 1
Prevalence and intensity of oocyst infections in F2 isofemale Anopheles gambiae families

Microsatellite genotyping and linkage analysis:

In all isofemale families, six loci were genotyped on the X chromosome and seven loci were genotyped on each of the second and third chromosomes. Given that the genome size of An. gambiae is 215 cM (Zheng et al. 1996), the resolution of our map scan is 10.75 cM. This marker density is appropriate for detecting and mapping of An. gambiae QTL at F2 recombination (Darvasi et al. 1993; Broman 2001). Genotypes of the F2 isofemale progeny were tested for Mendelian segregation distortion at each locus within each family. Significant segregation distortions were found for some loci, but the affected loci were not clustered on any particular chromosomal region and thus could not affect QTL analysis. Significant QTL for refractoriness were detected on chromosomes 2 and 3 in some isofemale families. None were detected on chromosome X. The two methods of QTL analysis gave similar results for QTL position and effects. Table 2 shows a summary of LOD score peaks and their locations, additive and dominance effects, and the variance explained by respective QTL. Figure 1 shows the QTL profiles for all studied isofemale families on chromosomes 2 and 3. Markers flanking peak LOD scores and 95% confidence intervals for QTL-containing regions are given in Table 3. The results given in Tables 2 and and33 and Figure 1 are from least-squares analysis by QTL Express, with the exception of the variance explained by the QTL that were derived from the maximum likelihood analysis with QTL Cartographer.

Figure 1.
Profiles of quantitative trait loci mapping on chromosome 2 (A) and 3 (B) for different isofemale families. The markers listed on the x-axis are microsatellite markers. The F-values were derived from least-squares analyses and the horizontal line indicates ...
TABLE 2
Summary of location and effects of detected QTL for refractoriness against Plasmodium falciparum infection in Anopheles gambiae isofemale families
TABLE 3
Location, confidence intervals, and flanking markers for significant refractoriness QTLs to Plasmodium falciparum infection in Anopheles gambiae isofemale families

For chromosome 2, three of five isofemale families studied showed significant QTL for the number of oocysts in F2 populations, whereas two isofemale families showed significant QTL on chromosome 3. The significant QTL detected on chromosome 2 in this study exhibited a significant additive genetic effect, but the respective dominance effects were not significant. In addition, these QTL consistently showed negative additive effects, suggesting that the QTL detected in the three isofemale families on chromosome 2 are likely to be the same loci and affect malaria parasites in a similar manner. Conversely, the significant QTL detected in two isofemale families for chromosome 3 appear to be acting dominant as indicated by their respective standard errors. The dominance effect measures the degree to which heterozygous individuals express the trait above or below the average for homozygous individuals. A positive sign will indicate that heterozygous mosquitoes exhibit a higher value than the average for homozygous mosquitoes, while a negative value will indicate that heterozygous mosquitoes have a lower average number of oocysts. In this study, the significant QTL on chromosome 3 had a negative dominance effect implying that they are associated with refractoriness to P. falciparum infection. From the genetic effects of the detected QTL and their relative positions on the two chromosomes, the most parsimonious interpretation of these results is that there is one additive QTL on chromosome 2 and one dominant QTL on chromosome 3 and that these control refractoriness in families in which significant QTL were detected, rather than multiple QTL occurring in different families.

DISCUSSION

The identification of chromosomal regions associated with resistance against infection by P. falciparum in An. gambiae is the starting point for efforts to clone and characterize the actual genes that confer refractoriness. Identification of such genes is important not only for understanding the biological mechanisms of refractoriness but also for insights into the coevolutionary relationships between mosquitoes and malaria parasites in different ecological regions where malaria is endemic. A number of studies have been undertaken to map chromosomal regions associated with different mechanisms of refractoriness to malaria parasites by mosquito vectors. Zheng et al. (1997) identified one major QTL named Pen1 and two minor QTL named Pen2 and Pen3 for encapsulation of P. cynomolgi B by An. gambiae. Similarly, Gorman et al. (1997) identified a QTL, which appeared to be identical to Pen1, for melanin deposition on charged sephadex beads and encapsulation of P. berghei by An. gambiae. In further analyses, Pen1 has been fine mapped to a region spanning 1.5 Mb of DNA, allowing comparison of An. gambiae and Drosophila melanogaster at the sequence level. This has revealed unusually high polymorphism at this locus (Thomasova et al. 2002). A study on P. cynomolgi Ceylon encapsulation by An. gambiae identified two chromosomal regions similar to Pen2 and Pen3 on chromosome 2 and another new QTL that mapped to chromosome 3; this implied variation of QTL for the encapsulation mechanism that varies with parasite species (Zheng et al. 2003). Despite the identification of major QTL for encapsulation using laboratory and field experiments, no single causative locus has yet been isolated. In addition, encapsulation of African-origin P. falciparum is not efficient in its natural vector, An. gambiae. Studies with An. gambiae mosquitoes from Tanzania have shown that <1% of the mosquitoes could encapsulate P. falciparum, but 90% of them could melanize sephadex beads (Schwartz and Koella 2002). More studies using natural vector–parasite interactions with the tools and knowledge acquired from studies with inbred strains are needed. Continued dissection of the mosquito genome, finer-scale mapping, increased functional characterization of genes and their protein products, and continued candidate gene analysis of mosquitoes' innate immunity genes will lead to more informed efforts to identify malaria refractory genes.

This study identified a QTL on chromosome 2 associated with microsatellite markers AG2H603 and AG2H135 in three isofemale families. The second locus detected in this study is near the proximal end of chromosome 3 near microsatellite marker AG3H93, and it appears to be a novel locus. The QTL detected on chromosome 2 was significant in three isofemale families, although these families were exposed to infectious blood meals from different gametocyte donors. Further, these QTL shared the same chromosomal regions and had similar genetic effects, suggesting the same gene or genes on chromosome 2 may be used against different parasite isolates by different mosquitoes. A previous linkage analysis study using F2 isofemale families of An. gambiae developed from wild-caught females was done in Mali, West Africa, and identified two QTL named pfin1 near AG2H603 and pfin2 near AG2H290 (Niare et al. 2002). In comparison to the West Africa study, our results suggest that the locus on chromosome 2 described here could be the same as pfin1. Interestingly, the region also encompasses the Pen3 locus, a locus associated with the parasite encapsulation mechanism (Gorman et al. 1997; Zheng et al. 1997). This suggests that a common genetic factor determining mosquito vector response to malaria parasites is located in this chromosomal region. The second QTL, pfin2, detected by Niare et al. (2002), was not detected in this study. The marker associated with pfin2, AG2H290, was consistently monomorphic in all the isofemale families used in this study. Failure to detect pfin2 and detection of a new QTL on chromosome 3 in our study suggests that additional genes and different gene combinations are involved in refractoriness to malaria parasites in western Kenya. This raises the possibility that different QTL may be responsible for refractoriness in different Plasmodium–mosquito interactions. This observation is not unique, as Zheng et al. (2003) have demonstrated that different QTL are responsible for encapsulation of different strains of P. cynomolgi. Such observations have also been made in D. melanogaster, where a fly strain that encapsulates eggs of the parasitic wasp Leptopilina boulardi L104 from Congo is susceptible to L. boulardi G317 from Tunisia and L. heterotoma, implying that expression of resistance to parasites varies among host–parasite systems (Carton and Nappi, 1997; Lambrechts et al. 2006). The design of the present study does not, however, allow the evaluation of whether mosquitoes from East and West Africa exhibit different refractoriness QTL or whether the genetic difference in the parasites between East and West Africa is dealt with by different QTL in mosquitoes. Genetic characterizations of factorial combinations between mosquito genotypes and parasite genotypes from the two regions are needed to answer this question.

Since the QTL detected in this study were based on the phenotype of infection intensities, as established by number of oocysts on day 8 postinfection, the loci could be associated with several biological mechanisms that reduce the number of oocysts. Such mechanisms include destruction of oocysts in ways similar to intracellular lytic responses in the midgut (Vernick et al. 1995), humoral responses against oocysts (Paskewitz et al. 1988), and mosquito defensin, an antibacterial peptide that destroys malaria parasites if a concurrent bacteria infection primes the mosquito's immune system (Lowenberger et al. 1999; Lambrechts et al. 2004). Several functional candidate gene analyses have implicated a number of genes and mechanisms that affect the ookinete and oocyst stages of development. Examples include thio-ester-containing proteins, leucin-rich protein, and c-type lectins (CTL4 and CTLMA2) that have been shown to influence refractoriness (Blandin et al. 2004; Osta et al. 2004). The knowledge and tools developed in such studies are useful in analyzing natural vector–parasite systems.

In summary, we identified two QTL conferring An. gambiae refractoriness to the P. falciparum parasite. These two QTL exhibit different genetic effects. The QTL on chromosome 2 is additive, while the one on chromosome 3 shows a dominant effect. The QTL on chromosome 2 maps to a similar region as Pen3 and pfin1, reported in previous studies. The QTL on chromosome 3 is novel. The two QTL explain ~20% of the variation in parasite density, suggesting that other not-yet-identified loci may be involved. Characterization of the natural mosquito populations in responses to infection and development of natural malaria parasites will provide valuable insights into the molecular basis of host resistance to malaria parasites and coevolution between mosquito vectors and malaria parasites. This will have important implications for the development of novel strategies for controlling malaria transmission.

Acknowledgments

We thank Ken Vernick from the University of Minnesota, George Seaton from the Roslyn Institute, Shengchu Wang from North Carolina State University, Dave Severson from the University of Notre Dame, and an anonymous reviewer for valuable discussions. This research was supported by National Institutes of Health grant D43 TW-01505 and United Nations Development Programme/WORLD BANK/World Health Organization Special Programme for Research and Training in Tropical Diseases (TDR) grant A10429.

References

  • Aultmann, K. S., B. J. Beaty and E. D. Walker, 2001. Genetically manipulated vectors of human disease: a practical overview. Trends Parasitol. 17: 507–509. [PubMed]
  • Barillas-Mury, C., B. Wizel and Y. S. Han, 2000. Mosquito immune responses and implications in malaria transmission: lessons from insect model systems and implications in vertebrate innate immunity and vaccine development. Insect Biochem. Mol. Biol. 30: 429–442. [PubMed]
  • Beier, J. C., G. F. Killeen and J. I. Githure, 1999. Entomologic inoculation rates and Plasmodium falciparum malaria prevalence in Africa. Am. J. Trop. Med. Hyg. 61: 109–113. [PubMed]
  • Blandin, S., S. H. Shiao, L. F. Moita, C. J. Janse, A. P. Waters et al., 2004. Complement-like protein TEP1 is a determinant of vectorial capacity in malaria vector Anopheles gambiae. Cell 116: 661–670. [PubMed]
  • Breman, J. G., M. S. Alilio and A. Mills, 2004. Conquering the intolerable burden of malaria: what's new, what's needed: a summary. Am. J. Trop. Med. Hyg. 71: 1–15. [PubMed]
  • Broman, K. W., 2001. Review of statistical methods for QTL mapping in experimental crosses. Lab. Anim. 30: 44–52. [PubMed]
  • Carton, Y., and A. J. Nappi, 1997. Drosophila cellular immunity against parasitoids. Parasitol. Today 13: 218–227. [PubMed]
  • Collins, F. H., R. K. Sakai, K. D. Vernick, S. Paskewitz, D. C. Seeley et al., 1986. Genetic selection of a Plasmodium-refractory strain of the malaria vector Anopheles gambiae. Science 234: 607–610. [PubMed]
  • Craig, Jr., G. B., 1967. Mosquitoes: female monogamy induced by male accessory gland substance. Science. 156: 1499–1501. [PubMed]
  • Curtis, C. F., 1994. The case for malaria control by genetic manipulation of its vectors. Parasitol. Today 10: 371–374. [PubMed]
  • Darvasi, A., A. Weinreb, V. Minke, J. I. Weller and M. Soller, 1993. Detecting marker-QTL linkage and estimating QTL gene effect and map location using a saturated genetic map. Genetics 134: 943–951. [PMC free article] [PubMed]
  • De Capurro, L., J. Coleman, B. T. Beernstsen, K. M. Myles, K. E. Olson et al., 2000. Virus expressed, recombinant single-chain antibody blocks sporozoite infection of salivary glands in Plasmodium gallinaceum-infected Aedes aegypti. Am. J. Trop. Med. Hyg. 4: 427–433. [PubMed]
  • Dimopoulos, G., H. M. Muller, E. A. Levashina and F. C. Kafatos, 2001. Innate immune defense against malaria infection in the mosquito. Curr. Opin. Immunol. 13: 79–88. [PubMed]
  • Gorman, J. M., D. W. Severson, A. J. Cornel, F. H. Collins and S. M. Paskewitz, 1997. Mapping quantitative trait locus involved in melanotic encapsulation of foreign bodies in malaria vector Anopheles gambiae. Genetics 146: 965–971. [PMC free article] [PubMed]
  • Gwadz, R. W., G. B Craig, Jr. and W. A. Hickey, 1971. Female sexual behavior as a mechanism rendering Aedes aegypti refractory to insemination. Biol. Bull. 140: 201. [PubMed]
  • Gwadz, R. W., D. Kaslow, J.-Y. Lee, W. L. Maloy, M. Zasloff et al., 1989. Effects of magainins and cercropins on the sporogonic development of malaria parasites in mosquitoes. Infect. Immun. 57: 2628–2633. [PMC free article] [PubMed]
  • Ito, J., A. Ghosh, L. A. Moreira, E. A. Wilmer and M. Jacobs-Lorena, 2002. Transgenic anopheline mosquitoes impaired in transmission of malaria parasite. Nature 417: 452–455. [PubMed]
  • Jacobs-Lorena, M., 2003. Interrupting malaria transmission by genetic manipulation of anopheline mosquitoes. J. Vect. Borne Dis. 40: 73–77. [PubMed]
  • Kleinschmidt, I., J. Omumbo, O. Briet, N. Giesen, N. Sogoba et al., 2001. An empirical malaria distribution map for West Africa. Trop. Med. Intern. Health 6: 779–786. [PubMed]
  • Kruglyak, L., J. M. Daly, M. P. Reeve-Daly and E. S. Lander, 1996. Parametric and nonparametric linkage analysis: a unified multipoint approach. Am. J. Hum. Gen. 58: 1347–1363. [PMC free article] [PubMed]
  • Lambrechts, L., J. M. Vulule and J. C. Koella, 2004. Genetic correlation between melanization and antibacterial immune responses in a natural population of the malaria vector Anopheles gambiae. Evolution Int. J. Org. Evolution 58: 2377–2381. [PubMed]
  • Lambrechts, L., S. Fellous and J. C. Koella, 2006. Coevolutionary interactions between host and parasite genotypes. Trends Parasitol. 22: 12–16. [PubMed]
  • Lander, E., and P. Green, 1987. Construction of multilocus genetic linkage maps in humans Proc. Natl. Acad. Sci. USA 84: 2363–2367. [PMC free article] [PubMed]
  • Lehman, T., M. Light, N. Lissa, B. T. A. Maega, J. M. Chimumbwa et al., 2003. Population genetic structure of Anopheles gambiae in Africa. J. Hered. 94: 133–137. [PubMed]
  • Lowenberger, C. A., S. Kamal, J. Chiles, S. Paskewitz, P. Bulet et al., 1999. Mosquito-Plasmodium interactions in response to immune activation of the vector. Exp. Parasitol. 91: 59–69. [PubMed]
  • Markianos, K., M. J. Daly and L. Kruglyak, 2001. Efficient multipoint linkage analysis through reduction of inheritance space. Am. J. Hum. Gen. 68: 963–977. [PMC free article] [PubMed]
  • Mulder, B., T. Tchuikam, K. Dechering, J. P. Verhave, P. Carnevale et al., 1994. Malaria transmission-blocking activity in experimental infections of Anopheles gambiae from naturally infected Plasmodium falciparum gametocyte carriers. Trans. R. Soc. Trop. Med. Hyg. 88: 121–125. [PubMed]
  • Mutero, C. M., J. H. Ouma, B. K. Agak, J. A. Wanderi and R. S. Copeland, 1998. Malaria prevalence and use of self-protection measures against mosquitoes in Suba District, Kenya. East Afr. Med. J. 75: 11–15. [PubMed]
  • Niare, O., K. Markianos, J. Volz, F. Oduol, A. Toure, et al., 2002. Genetic loci affecting resistance to human malaria parasites in a West African mosquito vector population. Science 298: 213–216. [PubMed]
  • Osta, M. A., G. K. Christophides and F. C. Kafatos, 2004. Effects of mosquito genes on Plasmodium development. Science 303: 2030–2032. [PubMed]
  • Paskewitz, S. M., M. R. Brown, A. O. Lea and F. H. Collins 1988. Ultrastructure of the encapsulation of Plasmodium cynomolgi (B strain) on the midgut of a refractory strain of Anopheles gambiae. J. Parasitol. 74: 432–439 [PubMed]
  • Richman, A., and F. C. Kafatos, 1996. Immunity to eukaryotic parasites in vector insects. Curr. Opin. Immunol. 8: 14–19. [PubMed]
  • Schwartz, A., and J. C. Koella, 2001. Trade-offs, conflict of interest and manipulation in Plasmodium-mosquito interactions. Trends Parasitol. 39: 84–88. [PubMed]
  • Schwartz, A., and J. C. Koella, 2002. Melanization of Plasmodium falciparum and C-25 sephadex beads by field-caught Anopheles gambiae (Diptera: Culicidae) from southern Tanzania. J. Med. Entomol. 39: 84–88. [PubMed]
  • Scott, J. A., W. G. Brogdon and F. H. Collins, 1993. Identification of single specimens of Anopheles gambiae complex by the polymerase chain reaction. Am. J. Trop. Med. Hyg. 49: 520–529. [PubMed]
  • Seaton, G., C. S. Haley, S. A. Knott, M. Kearsey and P. M. Visscher, 2002. QTL Express: mapping quantitative trait loci in simple and complex pedigrees. Bioinformatics 18: 339–340. [PubMed]
  • Serverson, D. W., 1997. RFLP analysis of insect genomes, pp. 309–320 in The Molecular Biology of Insect Disease Vectors, edited by J. M. Crampton, C. B. Beard and C. Louis. Chapman & Hall, London.
  • Shililu, J., C. Mbogo, C. Mutero, J. Gunter, C. Swalm et al., 2003. Spatial distribution of Anopheles gambiae and Anopheles funestus and malaria transmission in Suba District, western Kenya. Insect Sci. Appl. 23: 187–196.
  • Snow, R. W., C. A. Guerra, A. M. Noor, H. Y. Myint and S. I. Hay, 2005. The global distribution of clinical episodes of Plasmodium falciparum malaria. Nature 434: 214–217. [PMC free article] [PubMed]
  • Tchuinkam, T., B. Mulder, K. Dechering, H. Stoffels, J. P. Verhave et al., 1993. Experimental infections of Anopheles gambiae with Plasmodium falciparum of naturally infected gametocyte carriers in Cameroon: factors influencing the inefectivity of mosquitoes. Trop. Med. Parasitol. 44: 271–276. [PubMed]
  • Thomasova, D., L. Q. Ton, R. R. Copley, E. M. Zdobnov, X. Wang et al., 2002. Comparative genomic analysis in the region of a major Plasmodium-refractoriness locus of Anopheles gambiae. Proc. Natl. Acad. Sci. USA 99: 8179–8184. [PMC free article] [PubMed]
  • Tripet, F., Y. T. Toure, C. E. Taylor, D. E. Norris, G. Dolo et al., 2001. DNA analysis of transferred sperm reveals significant levels of gene flow between molecular forms of Anopheles gambiae. Mol. Ecol. 10: 1725–1732. [PubMed]
  • Vargas, L., 1949. Culicine and aedine mosquitoes and the malaria infections of lower animals, pp. 526–538 in Malariology, edited by M. F. Boyd. W. B. Saunders, Philadelphia.
  • Vernick, K. D., H. Fujioka, D. C. Seeley, B. Tandler, M. Aikawa et al., 1995. Plasmodium gallinaecium: A refractory mechanism of ookinete killing in the mosquito Anopheles gambiae. Exp. Parasitol. 80: 583–595. [PubMed]
  • Vernick, K. D., F. Oduol, B. P. Lazzaro, J. Glazebrook, J. Xu, et al., 2005. Molecular genetics of mosquito resistance to malaria parasites. CTMI 295: 383–415. [PubMed]
  • Visscher, P. M., R. Thompson and C. S. Haley, 1996. Confidence intervals in QTL mapping by bootstrapping. Genetics 143: 1013–1020. [PMC free article] [PubMed]
  • Wang, R., F. C. Kafatos and L. Zheng, 1999. Microsatellite markers and genotyping procedures for Anopheles gambiae. Parasitol. Today 15: 33–37. [PubMed]
  • Wang, S., C. J. Basten and Z-B. Zeng, 2001–2005 Windows QTL Cartographer 2.5. Department of Statistics, North Carolina State University, Raleigh, NC (http://statgen.ncsu.edu/qtlcart.wQTLcart.html/).
  • World Health Organization, 1998. Fact Sheet 94. World Health Organization, Geneva.
  • Zheng, L., M. Q. Benedict, A. J. Cornel, F. H. Collins and F. C. Kafatos, 1996. An integrated map of the African human malaria parasite mosquito, Anopheles gambiae. Genetics 143: 941–952. [PMC free article] [PubMed]
  • Zheng, L., A. J. Cornel, R. Wang, H. Erfle, H. Voss et al., 1997. Quantitative trait loci for refractoriness of Anopheles gambiae to Plasmodium cynomolgi B. Science 276: 425–428. [PubMed]
  • Zheng, L., S. Wang, P. Romans, H. Zhao, C. Luna et al., 2003. Quantitative trait loci in Anopheles gambiae controlling the encapsulation response against Plasmodium cynomolgi Ceylon. BMC Genet. 24: 16. [PMC free article] [PubMed]

Articles from Genetics are provided here courtesy of Genetics Society of America
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...