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Liverpool School of Tropical Medicine, Pembroke Place, Liverpool, L3 5QA, UK (G.L.)
Faculty of Life Sciences, University of Copenhagen, Thorvaldsensvej 40, 1870 Frederiksberg C, Denmark (Y.N.)
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 demonstrate 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 RNA interference 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 causing the most severe form of human malaria in Africa. The fact that mosquito strains completely 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 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 melanises a 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 melanisation occurs after parasite killing, as a means to dispose of dead parasites in some strains, while in others, killed parasites are only cleared by lysis (Fig. S2A) (5–7). In this study, we aimed at mapping the genomic regions and identifying genes that control resistance (i.e., 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 melanised parasites were counted on dissected midguts 7 to 9 days post infection (Fig. 1A and supporting online text). As expected, parental L3-5 females displayed only melanised 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 melanised parasites). Both parental and F1 phenotypes were present in the 402 F2 females. Percentages of resistant (i.e. devoid of live parasite) and melanising (bearing at least one melanised parasite) mosquitoes in each generation (Fig. 1B) did not follow the segregation pattern of simple Mendelian traits (p-values < 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.
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 supporting online text). Linkage analysis comparing resistant and non-resistant mosquitoes identified a single region on chromosome 3L (Fig. 1C). We interpreted this region, covering approximately 19Mb, 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 melanising and non-melanising 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 melanisation (Fig. 1C): a major QTL on chromosome 2R, which we named Pbmel1 (5Mb), for P. berghei melanisation locus 1, and a minor pericentromeric QTL on chromosome 3, Pbmel2 (17Mb), which partially overlaps with Pbres1. Linkage mapping using the actual counts of live or melanised parasites identified the same loci as above (Fig. S2B and supporting online text). Interestingly, the newly identified QTLs overlap with regions previously identified as controlling melanisation 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 melanisation 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: it binds to, and promotes the killing of midgut stages of the rodent parasite P. berghei, and it is highly polymorphic (5). To examine TEP1 polymorphism in the L3-5 and 4Arr strains, we cloned and sequenced the full ORF (Figs. 2 and S3). We renamed the previously known TEP1r (or TEP16) from the L3-5 strain (5, 13), TEP1*R1, and TEP1s (or TEP1) from the PEST strain (14), 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, 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.
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 instead of chromosomal deletions to silence each allele separately in F1 mosquitoes, enabling 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. We also note that G3 and 4Arr mosquitoes share the same susceptible phenotype.
We designed three pairs (a–c) of short dsRNAs (dsR/dsS) targeting specifically *R1 and *S3, and tested them in the parental L3-5 and G3 strains (Fig. S4 and supporting online 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: 3–4 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 to specifically study the function of each allele (Figs. 3B, S4B and supporting online text).
DsRNA-treated F1 mosquitoes were infected on mice carrying PbGFPcon (Figs. 3C and S4C). Control dsLacZ-treated F1 mosquitoes bore a mixture of live and melanised parasites. *R1-depleted mosquitoes (dsRa) were significantly more susceptible than *S3-depleted (dsSa) mosquitoes, and were completely unable to melanise. 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 melanisation of dead parasites. The reversal of the F1 phenotype towards the susceptible parent phenotype upon depletion of TEP1*R1, or towards 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 supporting online 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 required to precisely identify the essential SNP(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 and4A):4A): e.g. 50–70% of *R1/R2 and *R1/S2 mosquitoes were resistant in F2, while <7% were resistant in F1. Thus, this additional locus/loci appear to be unlinked to TEP1, and 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).
Importantly, 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, not with the major Plasmodium resistance island (PRI) (16–18) (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 regulates its binding to parasites and their subsequent killing (19, 20). Polymorphisms in proteins that control TEP1 function, or in TEP1 itself, 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, and how the determinant polymorphisms are maintained and selected in field populations, will be of tremendous importance for the control of malaria transmission.
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 the NIH and the Deutsche Forschungsgemeinschaft (L.M.S.), CNRS and INSERM (E.A.L. and S.A.B), the European Molecular Biology Organization Young Investigator 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 HHMI research scholar. EMBL Nucleotide Sequence Database accession numbers for TEP1 alleles sequenced in this report: FN431782 – FN431785.
Summary sentence: Polymorphisms in a mosquito complement-like factor confer resistance to malaria parasites.