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EMBO reports 7, 12, 1285–1289 (2006)
doi:10.1038/sj.embor.7400831
AOP Published online: 10 November 2006
Anopheles and Plasmodium: from laboratory models to natural systems in the field
Anna Cohuet1, Mike A Osta2, Isabelle Morlais3, Parfait H Awono-Ambene3, Kristin Michel2, Frederic Simard3, George K Christophides2, Didier Fontenille1 & Fotis C Kafatos2
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1 Institut de Recherche pour le Développement—Laboratoire de Lutte contre les Insectes Nuisibles, UR 016, BP 64501, 911 Avenue Agropolis, 34394 Montpellier cedex 5, France
2 Division of Cell and Molecular Biology, Imperial College London, Sir Alexander Fleming Building, South Kensington Campus, London SW7 2AZ, UK
3 Organisation de Coordination de la lutte contre les Endémies en Afrique Centrale, Laboratoire de Recherche sur le Paludisme, Yaounde BP 288, Cameroon
To whom correspondence should be addressed
Anna Cohuet Tel: +33 4 67 41 61 55; Fax: +33 67 54 20 44; E-mail: cohuet@mpl.ird.fr
Fotis C Kafatos Tel: +44 20 7594 1267; Fax: +44 20 7594 2056; E-mail: f.kafatos@imperial.ac.uk
Received 13 April 2006; Accepted 4 September 2006; Published online 10 November 2006.
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Abstract
Parasites that cause malaria must complete a complex life cycle in Anopheles vector mosquitoes in order to be transmitted from human to human. Previous gene-silencing studies have shown the influence of mosquito immunity in controlling the development of Plasmodium. Thus, parasite survival to the oocyst stage increased when the parasite antagonist gene LRIM1 (leucine-rich repeat immune protein 1) of the mosquito was silenced, but decreased when the C-type lectin agonist gene CTL4 or CTLMA2 (CTL mannose binding 2) was silenced. However, such effects were shown for infections of the human mosquito vector Anopheles gambiae with the rodent parasite Plasmodium berghei. Here, we report the first results of A. gambiae gene silencing on infection by sympatric field isolates of the principal human pathogen P. falciparum. In contrast with the results obtained with the rodent parasite, silencing of the same three genes had no effect on human parasite development. These results highlight the importance of following up discoveries in laboratory model systems with studies on natural parasite–mosquito interactions.
EMBO reports 7, 12, 1285–1289 (2006)
doi:10.1038/sj.embor.7400831
AOP Published online: 10 November 2006
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Introduction
Despite efforts in control and prevention, malaria remains one of the most devastating infectious diseases. The situation continues to worsen because of deteriorating socio-economic conditions and the increased resistance of parasites and vectors against the available antimalarial drugs and insecticides, respectively (Greenwood & Mutabingwa, 2002; Castro & Millet, 2005). The global malaria burden is mainly due to infections by Plasmodium falciparum—the most important and often deadly human malerial parasite. In much of Africa, where the malaria epidemic is most serious, most infections are transmitted by Anopheles gambiae. New approaches to control malaria include interrupting the transmission cycle in the mosquito; however, they require a better understanding of the specific interactions between human Plasmodium parasites and their natural vectors.
In life science research, model organisms greatly facilitate the identification of fundamental mechanisms that can be tested later in less tractable systems. For example, the key developmental mechanisms that were discovered in Drosophila melanogaster were later shown to be pertinent, even if not invariant, to most metazoa. The mouse model has been instrumental in biomedical research, even though its distinct physiology requires that discoveries in the mouse be validated in primates or humans before potential application to human medicine. Similarly, for complex diseases such as malaria in which three organisms—the human host, the Plasmodium parasite and the Anopheles vector—are implicated, model organisms greatly facilitate discovery. In recent years, the development of genomic and functional genomic approaches to malarial research has favoured the use of the principal human vector A. gambiae, although in combination with tractable non-human hosts and parasites. Such studies have explained important mechanisms of parasite killing in vectors. Testing their relevance to the human malarial system is now the primary goal.
Plasmodium parasites need to overcome several bottlenecks in the vector that result in most wild mosquitoes bearing few, if any, oocysts—that is, showing low infection prevalence and low parasite load (Vaughan et al, 1992; Gouagna et al, 1998; Sinden, 1999). Studies on the immune system of A. gambiae have shown that a fine balance exists between mosquito factors that affect ookinete survival to the oocyst stage, positively (agonists) or negatively (antagonists; Blandin et al, 2004; Osta et al, 2004). Such factors, and the molecular mechanisms they serve, might prove to be favourable targets for novel interventions towards blocking malaria transmission.
Putative components of the A. gambiae immune system were identified by using comparative genomic analysis (Christophides et al, 2002; Zdobnov et al, 2002) combined with transcriptomic studies of the A. gambiae response to Plasmodium berghei infections (Oduol et al, 2000; Christophides et al, 2002; Dimopoulos et al, 2002; Vlachou et al, 2005). Functional analysis by RNA interference-mediated gene silencing showed that certain genes, which are upregulated in the mosquito during ookinete invasion of the midgut, participate in humoral mechanisms of P. berghei killing in the vector. Thus, silencing of the antagonist LRIM1 (leucine-rich repeat immune protein 1) gene markedly increases the number of developing oocysts (Osta et al, 2004), as does the silencing of the complement-like gene thioester-containing protein (gene) 1 (TEP1; Blandin et al, 2004). Both LRIM1 and TEP1 encode circulating haemolymph (blood) proteins that favour the killing of ookinetes by lysis and melanization after midgut invasion. By contrast, two circulating C-type lectins—C-type lectin 4 (CTL4) and CTL mannose binding 2 (CTLMA2)—inhibit the melanization response and protect the development of ookinetes to the oocyst stage (Osta et al, 2004). More recent studies have established that antagonistic immune reactions are controlled, at least in part, by immune signalling pathways such as the Relish/Imd pathway (Meister et al, 2005). The number of P. berghei oocysts in A. gambiae is also regulated by local midgut epithelial responses that engage diverse—both negative and positive—factors (Vlachou & Kafatos, 2005; Vlachou et al, 2005).
The interplay of several humoral and cellular regulators of ookinete survival offers potential targets for novel malaria control strategies, aiming to support interactions that are either fatal for the parasite or to disrupt others that are protective. However, the interactions that have been identified in the laboratory combination A. gambiae–P. berghei might not fully represent those in the natural A. gambiae–P. falciparum systems (Boëte, 2005). Recently, a laboratory study has shown that transcriptional A. gambiae immune responses to P. falciparum and P. berghei are not identical, and might have different effects depending on the parasite species (Dong et al, 2006). Therefore, studies of interactions between A. gambiae and P. falciparum carried out under natural conditions are essential for understanding the specific molecular mechanisms that are pertinent to successful development of the principal human parasite, and to identify proper targets for interrupting human malaria transmission. Here, we have used gene silencing to investigate the role of CTL4, CTLMA2 and LRIM1 in the development of P. falciparum in A. gambiae, and the transcriptional responses of these genes to infection by this parasite species. These experiments were carried out under nearly natural conditions near Yaoundé in Cameroon, by membrane-feeding mosquitoes representative of the local vector population with field isolates of P. falciparum.
Results And Discussion
Gene silencing mediated by double-stranded RNA (dsRNA) injection was carried out in female mosquitoes of the Yaoundé strain and was confirmed by reverse transcription–PCR (RT–PCR). The knockdown efficiency (% kd) was determined by comparing, for each gene of interest, the transcript levels in mosquitoes injected 4 days earlier with either dsRNA for that gene (experimental mosquitoes) or with dsRNA for green fluorescent protein (GFP) (controls). The percentage knockdown values were reproducibly high and averaged 81.5% for CTL4, 96.9% for CTLMA2 and 96.8% for LRIM1. In previous experiments with P. berghei, such kd values resulted in strong effects on parasite prevalence and parasite load (Osta et al, 2004). Control and experimental mosquito groups were infected in parallel by feeding on four samples of human blood carrying field isolates of P. falciparum, including 20, 16, 16 or 32 gametocytes per microlitre. The respective oocyst prevalence in control midguts was 48.6%, 50.0%, 56.3% and 90.9%. The four replicates showed comparable kd values and were pooled (Table 1).
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Table 1
Plasmodium falciparum oocyst distribution in the midguts of knockdown Anopheles gambiae of the Yaoundé strain
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The mean number of oocysts per midgut in all control group mosquitoes (even when lacking oocysts) was 3.02 (s.e.=0.37); the oocyst load (mean oocyst number among mosquitoes showing at least one oocyst) was 4.98 (s.e.=0.47). These values are consistent with previous data on natural infections with P. falciparum in human blood (Tahar et al, 2002; Gouagna et al, 2004; Boudin et al, 2005; Lambrechts et al, 2005). By contrast, infections of the same vector with P. berghei frequently show mean oocyst numbers higher than 50 (Blandin et al, 2004; Osta et al, 2004; Abraham et al, 2005). This difference, between the laboratory model and the human malaria system in the field, might be inherent to the parasite itself or the host–parasite interactions.
Kolmogorov–Smirnov tests showed that at day 7 after infection, the distributions of developing P. falciparum oocysts in control and CTL4, CTLMA2 or LRIM1 knockdown mosquitoes are statistically similar (Table 1). Additionally, melanized ookinetes were not detected in the CTL knockdowns. These results are in contrast with our previous results from gene-silencing laboratory experiments on P. berghei infections (Osta et al, 2004). These previous experiments were carried out in the G3 strain, which originated from a mix of various populations of M and S molecular forms of A. gambiae and is polymorphic for chromosomal inversions. The G3 strain differs considerably from the Yaoundé strain used here, which is the offspring of a pure, standard chromosomal and molecular M form population from Cameroon. As an appropriate control, we used P. berghei to infect Yaoundé mosquitoes and confirmed both their high susceptibility to P. berghei and pronounced effects of gene silencing (Table 2); the results were comparable with those reported previously for infections of the G3 strain (Osta et al, 2004). Therefore, the effects of gene silencing on P. berghei and P. falciparum infections differ because of the parasite species and not the mosquito strain.
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Table 2
Plasmodium berghei oocyst distribution and melanized ookinetes in the midguts of knockdown Anopheles gambiae of the Yaoundé strain
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We compared the expression levels of CTL4, CTLMA2 and LRIM1 24 h after feeding Yaoundé mosquitoes with human blood, which was either non-infected or infected with P. falciparum. At this crucial time, parasites emerge at the basal surface of the midgut, come into contact with haemolymph components and either begin the transformation into oocysts or die. The data from two independent experiments showed an average of 2.4-, 1.7- and 1.5-fold upregulation of these genes in the carcasses of infected mosquitoes (Fig 1). A comparable level of upregulation in carcasses—where these genes are predominantly expressed—was detected in previous infections with P. berghei (Osta et al, 2004). Carcasses are enriched in haemocytes (mosquito blood cells) that predominantly expresses these genes (M.A.O., unpublished data). The midguts showed low expression levels of these genes and no significant upregulation on infection. It should be noted that, at least with respect to P. berghei, the upregulated genes include both a parasite antagonist (LRIM1) and two agonists (CTL4, CTLMA2). Thus, transcriptional upregulation does not correlate directly with the outcome of infections, although further studies are warranted to test this conclusion more rigorously.
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Figure 1
Plasmodium falciparum infection activates Anopheles gambiae immune response. Transcript levels of CTL4 (A), CTLMA2 (B) and LRIM1 (C) genes in midguts (Mg) and carcasses (Ca) of mosquitoes 24 h after feeding on blood from gametocyte carriers (black bars) or uninfected persons (grey bars). The ordinate represents relative expression values. Error bars represent s.d. in the pooled data set of two independent biological experiments.
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In conclusion, we have observed an important difference in the impact of silencing immunity genes on infection levels between the laboratory model (rodent) parasite and the sympatric natural (human) parasite isolates. Three A. gambiae genes encoding systemically circulating proteins, which strongly influence P. berghei development (positively or negatively), did not show comparable effects on P. falciparum infections in the same mosquito species and strain. It could be argued that the latter result might be due to the low infection levels of that parasite in A. gambiae. Future experiments to test this hypothesis definitively, by manipulating infection levels for both parasites, are conceivable but difficult—especially in the field—and are required to address this hypothesis. At least in terms of transcription, P. falciparum indeed affects the expression of some immune genes (Fig 1). Furthermore, an extensive series of studies on CTL4 failed to show a correlation between the degree of P. berghei ookinete melanization and infection levels (M.A.O., unpublished data). At present, we favour the hypothesis that the observed contrasting phenotypes might reflect co-evolution of the human parasite and its natural vector. Indeed, P. falciparum is naturally transmitted, but P. berghei might encounter A. gambiae only accidentally. This rodent parasite is naturally transmitted by A. dureni in Central Africa (reviewed by Gillies & De Meillon, 1968), and is unlikely to encounter the highly anthropophilic A. gambiae. The opposite is true for the human parasite P. falciparum, which shares an evolutionary history with the strongly anthropophilic A. gambiae, potentially resulting in unique co-adaptations. This supposition is consistent with the tantalizing observation that the A. gambiae refractory strain L3-5 is able to melanize numerous species of Plasmodium, including non-African P. falciparum isolates, but not—or only poorly—African P. falciparum isolates that are sympatric with A. gambiae (Collins et al, 1986). Co-adaptation would have favoured low infection levels if, as proposed by Boëte (2005), both parasite and host benefit by limiting the parasite numbers; infection is costly for the vector (Anderson et al, 2000; Ferguson & Read, 2002) and reducing the burden might favour transmission to the vertebrate. The bottleneck that natural parasites suffer at the ookinete-to-oocyst transition (Gouagna et al, 1998) might at first be interpreted as evidence for selection of host responses that reduce the parasite load. However, such selection is unlikely to be unconstrained: an excessive immune response could be detrimental not only for the parasites but also for the host itself, as immune system overactivation might have costly side effects (Moret & Schmid-Hempel, 2000; Schmid-Hempel, 2003). Thus, co-adaptation might have benefited both parties by favouring a robust equilibrium on the basis of limited, but sufficient to keep the parasite load low, responses of several A. gambiae genes (not just the ones we tested) to P. falciparum. Our data suggest that, despite the lower infection levels compared with P. berghei, P. falciparum induces an immune response of A. gambiae genes including, but probably not limited to, upregulation of LRIM1, CTL4 and CTLMA2 24 h after blood feeding. In fact, a recent genome-wide comparison of A. gambiae expression profiles after infection with P. berghei and P. falciparum parasites (Dong et al, 2006) showed that the former induces a wide and profound impact on the transcriptome, whereas the latter elicits a more specific immune response, possibly suggesting co-adaptation of the immune system to P. falciparum infection.
It remains unclear whether natural parasite–mosquito interactions are sufficiently distinct to preclude the possibility of extrapolating results obtained in P. berghei–A. gambiae to the natural transmission system. Future work will need to test whether other potent circulating immune-active proteins of the mosquito, such as TEP1 (Blandin et al, 2004) or RFABG (Vlachou et al, 2005), also show disparate effects on the rodent model compared with field isolates of the human parasite. In addition, proteins that are synthesized and locally active within the invaded epithelium, such as cytoskeleton-modifying components (Vlachou et al, 2005), also need to be tested. Genes in quantitative trait loci, shown by investigations in field conditions, such as Anopheles Plasmodium-responsive leucine-rich repeat 1 (APL1; Riehle et al, 2006) are also excellent candidates for affecting P. falciparum in natural transmission systems.
Although the possibility of parasite–vector genotypic co-adaptation is largely conjectural, a recent study has reported local effects of parasite genotype on the vector competence of specific mosquito genotypes (Lambrechts et al, 2005). To evaluate the possibility of parasite–vector co-adaptation and mosquito immune response in detail, this intriguing initial evidence needs to be followed up by studies on the infection and transcriptional regulation in diverse populations of A. gambiae by various Plasmodium species and P. falciparum genotypes. Ultimately, such studies will need to be combined with the current analysis of the role of human genotypes in malaria prevalence (Ruwende et al, 1995; Modiano et al, 2001; Fowkes et al, 2006). Although the complexity of genetic interactions in the host–parasite–vector trio might seem daunting, the availability of genomic and functional tools in all three organisms will make such studies feasible. In turn, such studies might lead to a deeper understanding of the patterns of malaria transmission and potentially provide new approaches to controlling this devastating global disease.
Methods
Mosquito strain and gene silencing. The sympatric mosquitoes used in this study were A. gambiae of the Yaoundé strain. The Yaoundé mosquito strain was established in 1988 from field specimens caught in the suburbs of Yaoundé city, in the south of Cameroon (Tchuinkam et al, 1993). Mosquitoes that fed through a parafilm membrane were used and then maintained under standard insectary conditions. The colony belongs to the M molecular and Forest chromosomal forms (standard chromosomal arrangements). The Yaoundé strain is representative of the local natural transmission system in the south of Cameroon, where malaria transmission is mainly due to A. gambiae (M and S molecular forms, Forest chromosomal form), although A. funestus, A. nili and A. moucheti can also be locally important (Antonio-Nkondjio et al, 2002).
Gene silencing was achieved by injecting 207 ng of dsRNA into the thorax of 1-day-old females, as described previously (Blandin et al, 2002). In each of five replicates carried out, 60–100 females were injected with dsRNA for each of the genes tested. DsRNA for GFP was used as reference. The efficiency of gene silencing was determined as described by Osta et al (2004).
Plasmodium falciparum gametocyte carriers-recruitment and experimental infection. P. falciparum gametocyte carriers were recruited among 5- to 12-year-old children at schools in Mfou, a town located 30 km from Yaoundé city in an area endemic for malaria. Blood samples were collected by finger-prick from each volunteer and thick blood smears were stained with Giemsa and examined by microscopy for the presence of P. falciparum. Sexual and asexual stages were counted in observed fields that cumulatively contained at least 500 leukocytes; an estimate of parasite density was obtained by assuming a standard number of 8,000 leukocytes/ l of blood. Children with asexual parasitaemia exceeding 1,000 parasites/l were immediately treated with the amodiaquine and artesunate drug combination according to national guidelines. Asymptomatic gametocyte-positive children were enrolled as volunteers. The recruitment procedures were approved by the Cameroonian and WHO ethical review committees.
For each experimental replicate, mosquitoes were injected with dsRNA and 4 days later were fed on blood from different P. falciparum gametocyte carriers. Five millilitres of venous blood was collected in heparinized vacutainers. To limit the effect of human factors, such as transmission-blocking immunity (Boudin et al, 2005), the blood was centrifuged for 3 min at 2,000 r.p.m. and the serum was replaced with the same volume of AB serum from a French donor with no exposure to malaria. Mosquitoes were starved for 16 h and allowed to feed for 15 min on this mixture using standard membrane-feeding (Tchuinkam et al, 1993). Unfed and partially fed mosquitoes were discarded. After 7 days, mosquitoes were dissected and midguts were stained with 0.4% mercurochrome in distilled water to count oocyst numbers by light microscopy. Experiments were considered successful when the infection prevalence of control mosquitoes (GFP dsRNA) was at least 30%. Mosquito infections with the P. berghei GFP-CON strain (Franke-Fayard et al, 2004) were carried out as described by Osta et al (2004).
Transcript level analysis in infected mosquitoes compared with non-infected mosquitoes. Females of the Yaoundé A. gambiae strain were fed on either gametocyte-positive blood or non-infected human blood. Transcript level analysis of CTL4, CTLMA2 and LRIM1 was carried out by using QRT–PCR. Briefly, midguts and carcasses of Yaoundé mosquitoes, fed on uninfected human blood or P. falciparum-infected human blood, were dissected 24 h after blood feeding, and were kept separately in RNAlater (Ambion, Austin, TX, USA) before RNA isolation. A batch of 30 control mosquitoes was dissected 7 days after blood feeding to determine the level of infection. Total RNA was isolated by using the TRIzol Reagent (Invitrogen, Paisley, UK) according to the supplier's instructions, and contaminant genomic DNA was removed by DNase I (Invitrogen) treatment. Complementary DNA was synthesized from total RNA (2–3 g) by using the SuperScript II RNase H- Reverse Transcriptase and oligo(dT)12–18, as described by the supplier (Life Technologies Inc., Paisley, UK). QRT–PCR was carried out with the ABI Prism 7700 Sequence Detection System, by using the SYBR Green PCR Master Mix kit (Applied Biosystems, Foster City, CA, USA) according to the manufacturer's instructions. Primer sequences are described by Osta et al (2004). Relative gene expression values were calculated by using the comparative CT method after checking for efficiency of target amplification as described in the User Bulletin #2. The S7 ribosomal protein gene was used as an internal reference.
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Acknowledgements
We thank Dr M. Engelbert, C. Efemba and E. Bozewan of the Mfou hospital for assistance; A.M. Mendes and Dr D. Vlachou for sharing materials; and A.C. Koutsos and Dr C. Boëte for stimulating discussions. We are grateful to the inhabitants of Mfou for their cooperation. M.A.O. was supported by a Marie Curie Intra-European fellowship. This investigation received financial support from the UNICEF/UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases (TDR), grant A50241.
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