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Dissecting the Genetic Basis of Resistance to Malaria Parasites in Anopheles gambiae

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Science  02 Oct 2009:
Vol. 326, Issue 5949, pp. 147-150
DOI: 10.1126/science.1175241

Variable Defenses

Recent mapping of resistance alleles in the mosquito Anopheles gambiae that provide protection against the human malaria parasite Plasmodium falciparum revealed a major Plasmodium resistance island (PRI), comprising allelic versions of two leucine-rich repeat-containing proteins, LRIM1 and APL1, which form a complex with the complement C3-like protein TEP1. Using RNA interference inactivation of heterozygous allelic versions of TEP1 genes, Blandin et al. (p. 147) show that TEP1 heterogeneity reflects phenotypic variation among mosquito strains parasitized with the rodent malaria parasite Plasmodium berghei. It remains unclear whether the observed differences are the outcomes of different selection regimes, because of differing mechanisms, or because the complex is also used in other contexts.

Abstract

The ability of Anopheles gambiae mosquitoes to transmit Plasmodium parasites is highly variable between individuals. However, the genetic basis of this variability has remained unknown. We combined genome-wide mapping and reciprocal allele-specific RNA interference (rasRNAi) to identify the genomic locus that confers resistance to malaria parasites and demonstrated that polymorphisms in a single gene encoding the antiparasitic thioester-containing protein 1 (TEP1) explain a substantial part of the variability in parasite killing. The link between TEP1 alleles and resistance to malaria may offer new tools for controlling malaria transmission. The successful application of rasRNAi in Anopheles suggests that it could also be applied to other organisms where RNAi is feasible to dissect complex phenotypes to the level of individual quantitative trait alleles.

Anopheles gambiae mosquitoes are major vectors of Plasmodium falciparum, a protozoan parasite that causes the most severe form of human malaria in Africa. The fact that mosquito strains that are completely resistant to malaria parasites can be selected (1, 2) indicates that genetic factors in mosquitoes control the level of parasite transmission. Understanding the genetic basis of this resistance has been a long-standing question. The L3-5 resistant strain kills and melanizes a wide variety of parasite species (1). Previous genetic analyses of crosses between this strain and the susceptible 4Arr strain infected with two simian parasite species focused on the melanotic encapsulation phenotype and identified several quantitative trait loci (QTLs), whose relative contributions varied with parasite species and between F2 generation families (3, 4). Recently, it became clear that melanization occurs after parasite killing as a means to dispose of dead parasites in some strains, whereas in others, killed parasites are only cleared by lysis (fig. S2A) (57). In this study, we aimed at mapping the genomic regions and identifying genes that control resistance (the absence of live parasites) of mosquitoes to the rodent malaria parasite Plasmodium berghei.

We set up reciprocal crosses of the resistant L3-5 and susceptible 4Arr strains. F1 mosquitoes were intercrossed, and individual females were isolated to lay eggs, yielding 10 F2 families. Females were blood-fed on mice infected with PbGFPcon, a transgenic clone of P. berghei expressing GFP constitutively (8). Fluorescent live and dead melanized parasites were counted on dissected midguts 7 to 9 days post infection [Fig. 1A and supporting online material (SOM) text]. As expected, parental L3-5 females displayed only melanized parasites (with the exception of one that bore one live parasite), and 4Arr mosquitoes displayed only live parasites. Most of the 111 F1 mosquitoes exhibited an intermediate phenotype (mix of live and melanized parasites). Both parental and F1 phenotypes were present in the 402 F2 females. Percentages of resistant (devoid of live parasite) and melanizing (bearing at least one melanized parasite) mosquitoes in each generation (Fig. 1B) did not follow the segregation pattern of simple Mendelian traits [P < 0.001 in both cases (9)], indicating that the killing of P. berghei and the mode of clearance of dead parasites are complex traits that are each likely to result from the segregation of several alleles.

Fig. 1

Loci associated with resistance and clearance of dead parasites. (A) Numbers of melanized (x axis) and live (y axis) parasites per mosquito in reciprocal crosses of the resistant L3-5 and susceptible 4Arr strains. (B) Percentages of resistant (devoid of live parasites) and melanizing (bearing at least one melanized parasite) mosquitoes in each generation. (C) Linkage mapping for the resistant (red) and melanizing (black) traits, with estimated logarithm of the odds ratio for linkage (LOD score) thresholds represented as dotted lines (3.00 and 2.88, respectively). Genetic markers; centromere positions (C); chromosome arms; and the TEP1, LRIM1, and APL1 loci (light blue) are indicated below the axes. Previously identified QTLs for resistance against simian parasites (light purple) or P. falciparum (dark purple) are positioned below the chromosomes. The PRI corresponds to the region covered by the QTLs Pfin1, Pfin4, Pfin5, and Pfmel2.

To map loci controlling resistance to parasites, we genotyped 39 informative markers spanning the entire genome in 206 selected F2 individuals with extreme phenotypes (fig. S1 and SOM text). Linkage analysis comparing resistant and nonresistant mosquitoes identified a single region on chromosome 3L (Fig. 1C). We interpreted this region, covering ~19 Mb, as a major locus responsible for resistance to P. berghei and named it Pbres1 for P. berghei resistance locus 1. We further compared the genotypes of melanizing and nonmelanizing mosquitoes and detected two intervals that are likely to contain regulators affecting the mode of clearance of dead parasites (i.e., the balance between lysis and melanization) (Fig. 1C): (i) a major QTL on chromosome 2R, which we named Pbmel1 (5 Mb) for P. berghei melanization locus 1, and (ii) a minor pericentromeric QTL on chromosome 3, Pbmel2 (17 Mb), that partially overlaps with Pbres1. Linkage mapping using the actual counts of live or melanized parasites identified the same loci as above (fig. S2B and SOM text). The newly identified QTLs overlap with regions previously identified as controlling melanization of P. cynomolgi and P. berghei in L3-5 mosquitoes (Fig. 1C) (3, 4, 10), indicating that the major mechanisms underlying parasite elimination in L3-5 are likely partially conserved and independent of parasite species. Nevertheless, clear quantitative differences exist between the four studies, probably, at least in part, because previous studies did not consider resistance and melanization as distinct traits.

Because of its major role in parasite transmission, we investigated the resistance QTL on chromosome 3L in more detail. Pbres1 contains ~975 genes, among which 35 can be classified as “immune-related” (11). This category includes the gene encoding the thioester-containing protein 1 (TEP1), a complement-like molecule circulating in the hemolymph with key antiparasitic activity (12). Two features make it an attractive candidate: (i) it binds to and promotes the killing of midgut stages of the rodent parasite P. berghei, and (ii) it is highly polymorphic (5). To examine TEP1 polymorphism in the L3-5 and 4Arr strains, we cloned and sequenced the full open reading frame (Fig. 2 and fig. S3). We renamed the previously known TEP1r (or TEP16) from the L3-5 strain (5, 13) as TEP1*R1, and we renamed TEP1s (or TEP1) from the PEST strain (14) as TEP1*S1. All TEP1 sequences in L3-5 mosquitoes were identical to *R1. Sequences from the 4Arr strain appeared to be chimeras of TEP1*S and TEP1*R: One was closer to *S1, thus we named it TEP1*S2; the second allele clustered with *R1 in the phylogenetic tree and was therefore named TEP1*R2. We also sequenced TEP1 from our G3 colony and confirmed that it was closely related, although not identical, to TEP1*S1. We named this allele TEP1*S3.

Fig. 2

TEP1 polymorphism. (A) Phylogenetic tree built from the global alignment of complete amino acid sequences of TEP1 alleles from L3-5 (*R1), 4Arr (*R2 and *S2), and G3 (*S3) mosquitoes and the previously described *S1 allele from the PEST strain. Scale bar indicates estimated amino acid substitutions per site. (B) Schematic representation of TEP1 sequences. Amino acid sequences of *S1 and *S3 are represented by orange horizontal bars, *R1 by a blue bar. The 4Arr alleles are combinations of *S1/S3 and *R1, as illustrated by short stretches of aligned sequences. The short horizontal lines below *R1 and *S3 indicate the regions targeted by dsRa and dsSa, respectively.

To determine whether the diverse alleles of TEP1 have a phenotypic effect, we compared the degree of resistance of mosquitoes that differed solely in the expression of TEP1 alleles. For this, we developed an assay similar to reciprocal hemizygosity analysis in yeast (15): We used reciprocal allele-specific RNA interference (rasRNAi) instead of chromosomal deletions to silence each allele separately in F1 mosquitoes, enabling us to compare the function of each allele in the same genetic background (Fig. 3A). We crossed resistant L3-5 and susceptible G3 mosquitoes, as these strains are homozygous for TEP1 and bear representative alleles of the TEP1*R and TEP1*S classes. Additionally, G3 and 4Arr mosquitoes share the same susceptible phenotype.

Fig. 3

TEP1*R1 is more efficient than TEP1*S3 in parasite killing. (A) rasRNAi. Each box represents a gene. With the use of short dsRNA probes specifically directed against *R1 (dsR) or *S3 (dsS), each TEP1 allele is silenced separately in F1 mosquitoes (open box), allowing us to compare the function of each allele in the same genetic background. (B) TEP1 expression in the F1 progeny of crosses between L3-5 females and G3 males (L3-5 × G3) was measured by allele-specific quantitative real-time polymerase chain reaction 3 days after dsRNA treatment. Expression levels of TEP1*R1 and *S3 were normalized to their levels in the dsLacZ control. Mean (central bar) ± SEM (error bar) of three independent experiments. (C) Parasite counts in the F1 progeny of L3-5 × G3. The results of three independent experiments were pooled, and sample sizes are shown in brackets. Mean (central bar) ± SEM (error bar). Significance for differences between groups are indicated (Mann-Whitney on key comparisons): **P < 0.001; ns, not significant.

We designed three pairs (a to c) of short double-stranded RNAs (dsRNAs; dsR/dsS) specifically targeting *R1 and *S3 and tested them in the parental L3-5 and G3 strains (fig. S4 and SOM text). We used dsLacZ as a negative control and dsTEP1 that targets both alleles as a positive control (5). Pair a (dsRa and dsSa) was selected for further experiments: 3 to 4 days after dsRNA treatment, TEP1*R1 was depleted from L3-5 mosquitoes upon treatment with dsRa, but not dsSa, and reciprocally, injection of dsSa, but not dsRa, in G3 mosquitoes reduced TEP1*S3 levels. In the F1 progeny of reciprocal crosses between L3-5 and G3 mosquitoes, both alleles were silenced to a similar level by allele-specific RNAi, allowing us to specifically study the function of each allele (Fig. 3B, fig. S4B, and SOM text).

dsRNA-treated F1 mosquitoes were infected on mice carrying PbGFPcon (Fig. 3C and fig. S4C). Control dsLacZ-treated F1 mosquitoes bore a mixture of live and melanized parasites. *R1-depleted mosquitoes (dsRa) were significantly more susceptible than *S3-depleted (dsSa) mosquitoes and were completely unable to melanize. Moreover, *S3-depleted mosquitoes were consistently more resistant than dsLacZ controls, containing fewer live parasites. Thus, TEP1*R1 is more efficient than TEP1*S3 in promoting parasite killing and melanization of dead parasites. The reversal of the F1 phenotype toward the susceptible parent phenotype upon depletion of TEP1*R1, or toward the resistant parent phenotype upon depletion of TEP1*S3, indicates that polymorphisms in TEP1 are a major determinant of resistance to P. berghei in these mosquito strains.

To examine whether the two allelic variants of the 4Arr strain (*S2 and *R2, which belong to the TEP1*S and TEP1*R classes) also differ in their efficiency in parasite killing, we further refined our association analysis of the F2 progeny of the QTL mapping crosses and genotyped all F2 progeny for TEP1 (Fig. 4 and SOM text). Most *R1/R1 F2 mosquitoes (81%) were fully resistant, and those that were not carried only a few live parasites. In contrast, 90% of *S2/S2 mosquitoes were susceptible, containing high parasite loads. *R2/R2 mosquitoes had an intermediate phenotype, suggesting that although *R2 is closely related to *R1, the few polymorphisms between these two alleles affect its efficiency in parasite killing. Further studies are needed to precisely identify the essential single-nucleotide polymorphism(s) and the molecular mechanisms that underlie this resistance. In addition, *R1/R2 mosquitoes were more resistant than *R1/S2 mosquitoes, indicating that the two 4Arr alleles confer different degrees of resistance, with *R2 > *S2. Thus, the complexity of the resistance inheritance in our crosses is partially explained by the segregation of the three TEP1 alleles. Still, other genes besides TEP1 must contribute. This is apparent from comparing phenotypes of groups from different generations with the same TEP1 genotypes (Figs. 1B and 4A): For instance, 50 to 70% of *R1/R2 and *R1/S2 mosquitoes were resistant in F2, whereas <7% were resistant in F1. Thus, this additional locus (or loci) appears to be unlinked to TEP1 and also to have a limited impact in mosquitoes homozygous for the extreme alleles *R1 and *S2, which have similar resistance as the parental strains but are essential to support resistance in heterozygotes. Future work may focus on identifying secondary QTL(s) and potential candidate TEP1 suppressor gene(s).

Fig. 4

Correlation between TEP1 genotype and phenotype upon P. berghei infection in the F2 generation. (A) Percentages of resistant and melanizing mosquitoes for each genotype. Sample sizes are shown in brackets. Significance for differences between groups were calculated taking into account F2-family structure (9): **P < 0.001; *P < 0.05; ns, not significant. (B) Parasite counts in F2 mosquitoes as shown in Fig. 1A.

The single locus identified here that controls resistance to P. berghei and includes TEP1 does not overlap with previously reported QTLs controlling the intensity of infection of natural populations by the human malaria parasite P. falciparum, and in particular, does not overlap with the major Plasmodium resistance island (PRI) (1618) (Fig. 1C). Two leucine-rich repeat proteins encoded in the PRI, APL1 and LRIM1, form a complex with TEP1. These proteins maintain mature TEP1 in circulation and regulate its binding to parasites and their subsequent killing (19, 20). Polymorphisms in TEP1 itself or in proteins that control TEP1 function might both contribute to the efficiency of TEP1 antiparasitic activity. The differences between the QTLs identified in laboratory strains and in field mosquitoes might thus reflect the sampling of determinant polymorphism(s) in various players of the same pathway, rather than different mechanisms employed to limit development of human and rodent malaria parasite species. Consistently, silencing of TEP1 increases A. gambiae susceptibility to both murine and human Plasmodia (5, 21). Haplotypes of the susceptible and resistant alleles of TEP1, as well as recombinants between these forms, exist in field populations from East and West Africa (22). Understanding the genetic basis of resistance to malaria parasites, as well as how the determinant polymorphisms are maintained and selected in field populations, will be of tremendous importance for the control of malaria transmission.

Supporting Online Material

www.sciencemag.org/cgi/content/full/326/5949/147/DC1

Materials and Methods

SOM Text

Figs. S1 to S4

Tables S1 to S3

References

  • * These authors contributed equally to this work.

  • Present address: Liverpool School of Tropical Medicine, Pembroke Place, Liverpool, L3 5QA, UK.

  • Present address: Faculty of Life Sciences, University of Copenhagen, Thorvaldsensvej 40, 1870 Frederiksberg C, Denmark.

References and Notes

  1. Materials and methods are available as supporting material on Science Online.
  2. We thank A. Cohuet for sharing sequencing information, R. Carmouche and J. Luis from the EMBL Genecore facility for support with genotyping, R. Bourgon for advice on statistical analyses, A. Budd for suggestions on the phylogenetic analysis, D. Doherty and J. Soichot for help with breeding mosquitoes, and E. Marois and C. Ramakrishnan for comments on the manuscript. We acknowledge the support of F. Kafatos, in whose laboratory this work was initially carried out. This work was supported by grants from NIH and the Deutsche Forschungsgemeinschaft (L.M.S.), CNRS and INSERM (E.A.L. and S.A.B), the European Molecular Biology Organization (EMBO) Young Investigators Program (E.A.L.), and the European Commission Network of Excellence “BioMalPar” (E.A.L.). S.A.B. received a postdoctoral fellowship from EMBO. E.A.L. is an international Howard Hughes Medical Institute research scholar. EMBL Nucleotide Sequence Database accession numbers for TEP1 alleles sequenced in this report are as follows: FN431782 to FN431785.
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