Gene targeting in the rodent malaria parasite Plasmodium yoelii

Molecular & Biochemical Parasitology 113 (2001) 271– 278 www.parasitology-online.com.

Gene targeting in the rodent malaria parasite Plasmodium yoelii Maria M. Mota a,*, Vandana Thathy b, Ruth S. Nussenzweig b, Victor Nussenzweig a a

Michael Heidelberger Di6ision, Department of Pathology (MSB131), New York Uni6ersity Medical Center, 550 First A6enue, New York, NY 10016, USA b Department of Medical and Molecular Parasitology, New York Uni6ersity School of Medicine, 341 E. 25th street, New York, NY 10010, USA Received 12 October 2000; accepted 19 January 2001

Abstract It is anticipated that the sequencing of Plasmodium falciparum genome will soon be completed. Rodent models of malaria infection and stable transformation systems provide powerful means of using this information to study gene function in vivo. To date, gene targeting has only been developed for one rodent malaria species, Plasmodium berghei. Another rodent species, Plasmodium yoelii, however, is favored to study the mechanisms of protective immunity to the pre-erythrocytic stages of infection and vaccine development. In addition, it offers the opportunity to investigate unique aspects of pathogenesis of blood stage infection. Here, we report on the stable transfection and gene targeting of P. yoelii. Purified late blood stage schizonts were used as targets for electroporation with a plasmid that contains a pyrimethamine-resistant form of the P. berghei dihydrofolate reductase-thymidylate synthase (Pbdhfr-ts) fused to green fluorescent protein (gfp) gene. After drug selection, fluorescent parasites contained intact, non-rearranged plasmids that remain stable under drug-pressure. In addition, we used another dhfr-ts/gfp based plasmid to disrupt the P. yoelii trap (thrombospondin-related anonymous protein) locus by site-specific integration. The phenotype of P. yoelii TRAP knockout was identical to that previously reported for the P. berghei TRAP knockout. In the absence of TRAP, the erythrocytic cycle, gametocyte and oocyst development of the mutant parasites were indistinguishable from wild type (WT). Although the sporozoites appeared morphologically normal, they failed to glide and to invade the salivary glands of mosquitoes. © 2001 Elsevier Science B.V. All rights reserved. Keywords: Plasmodium yoelii; Transfection; Gene targeting; TRAP; Malaria

1. Introduction Malaria is caused by a parasitic protozoa of the genus Plasmodium and infection with Plasmodium falciparum is responsible for millions of deaths per annum. The sequencing of the entire genome of this parasite will soon be completed. Animal models and stable transformation systems will be very helpful in utilizing this vast amount of information to study gene function and accelerate the development of vaccines and drugs. Stable transformation and gene targeting are established for just one of the rodent malaria species, Plasmodium berghei [1,2]. This is regrettable since other Abbre6iations: DHFR-TS, dihydrofolate reductase-thymidylate synthase; GFP, green fluorescent protein; TRAP, thrombospondin-related anonymous protein; UTR, untranslated region; WT, wild type. * Corresponding author. Tel.: + 1-212-2635346; fax: +1-2122638179. E-mail address: [email protected] (M.M. Mota).

species of rodent malaria parasites are useful to study different aspects of the biology, pathology and the mechanisms of protective immunity of the human malaria parasites. For example, studies involving the biology of the Plasmodium pre-erythrocytic stages as well as the mechanisms of protective immunity elicited by immunization with irradiated sporozoites have focused preferentially on Plasmodium yoelii rather than P. berghei. The main reason for favoring P. yoelii is the striking difference in infectivity for inbred strains of mice of sporozoites from these two species of rodent malaria. Similar to P. falciparum, ten or fewer P. yoelii sporozoites are enough to infect mice, whereas hundreds of P. berghei sporozoites are required to ensure a blood stage infection [3–5]. These differences in infectivity are associated with changes in antigenicity of the pre-erythrocytic stages, and in the immune response of the host. For example, sterile immunity can be obtained in many strains of mice after a single or very few

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injections of irradiated P. berghei sporozoites [6], whereas it requires multiple doses of P. yoelii sporozoites [7]. Immunity to the pre-erythrocytic stages of human malaria parasites also develops slowly [8]. Thus, P. yoelii appear to be more ‘adapted’ to this unnatural host (its natural host being Thamnomys rutilans) and is presumed to be a better experimental model for the human infection at the pre-erythrocytic level. The erythrocytic infection by P. yoelii also provides attractive targets for investigation. For example, two P. yoelii parasite lines with distinct virulence patterns, non-lethal 17X and lethal YM [9], differ widely in their infectivity for mature or immature erythrocytes, but the underlying molecular mechanisms remain obscure. For these reasons, sequencing of the P. yoelii genome has been undertaken, and a 2X coverage is already available in the TIGR web site. Here we report, for the first time, on the stable transfection and gene targeting of P. yoelii, which now offers the possibility to search for virulence factors and to study the function of specific pre-erythrocytic parasite antigens in the context of a better model of host – parasite interactions.

2. Materials and methods

2.1. Plasmid constructions The construction of plasmid pMD205-GFP, originally named pPyrFlu, has been described [10]. Briefly, it consists of a pyrimethamine-resistant form of P. berghei dihydrofolate reductase-thymidylate synthase (dhfr-ts) gene fused to the gfpmut2 gene [11] under the control of 2.2 kb of 5%- and 0.5 kb of 3%-UTRs from Pbdhfr-ts. Plasmid pPyTRAPint was constructed by directionally cloning an internal truncated fragment of the P. yoelii trap gene into plasmid pMD205-GFP digested with the restriction enzymes BamHI and NotI (Fig. 3A). The PyTRAP fragment was cloned by PCR using P. yoelii 17X genomic DNA and oligonucleotides PyTRAP-F (sense, 5%-ggc ggatcc gcgtgttccttaatggtcaggaaactctt-3%) and PyTRAP-R (antisense, 5%-ggc gcggccgc atatccattattagatttagactggtttt-3%). The underlined sequences represent BamHI and NotI restriction enzyme sites, respectively. The truncated TRAP fragment lacks nucleotides 1–56 and the last 196 nucleotides of the TRAP open reading frame.

2.2. P. yoelii pyrimethamine sensiti6ity To test the sensitivity of P. yoelii 17X to pyrimethamine, experiments were performed in which groups of 3 P. yoelii infected mice were treated for different periods of time with different concentrations of drug. To assess the level of sensitivity to the drug,

parasitemia was monitored by light microscopy of air dried, methanol fixed, blood smears stained with 10% Giemsa stain.

2.3. Parasite manipulations P. yoelii 17X was obtained as a cloned line from the WHO Registry of Standard Malaria Parasites, University of Edinburgh and was a gift of Professor David Walliker. Parasite reference populations (stabilates) were prepared from Balb/c mice infected by bite of Anopheles stephensi mosquitoes. Cryopreserved parasite stabilates were thawed at 37°C and promptly syringe passaged into 16–18 g Balb/c mice intraperitoneally (i.p.). Parasites were then used directly from these mice or first syringe passaged to as many mice as desired. In order to obtain a preparation containing \ 90–95% schizonts for transformation, the blood was collected by cardiac puncture from one to five mice, depending on the parasitemia (corresponding to approximately 5–10×107 infected erythrocytes). The collected blood was initially placed in 10 ml of complete culture medium (RPMI 1640 medium containing heat-inactivated fetal calf serum (20%), neomycin (50 IU ml − 1) and Hepes (25 mM)), containing 40 IU of heparin, and washed once (300 g for 8 min). The isolated blood sample was subsequently incubated in 150 ml of complete culture medium at 10% O2, 5% CO2, 85% N2 gas mixture at 37°C for different periods of time. After different times of culture, each 35 ml of culture suspension was layered on top of 10 ml of different Nycodenz/ phosphate buffered saline (PBS) gradients and centrifuged for 25 min at 300× g at room temperature. The schizonts were collected on the interface and washed once for 8 min at 200×g with culture medium. The schizont pellet (approximately 100 ml) was then resuspended in 300 ml of PBS containing 0.2 mM CaCl2 for assessing the numbers of schizonts obtained by each culture and/or gradient condition for transfection.

2.4. Transfection and selection of transformants For each electroporation, 10–40 mg of plasmid DNA was added to 5–10× 107 schizonts in a 0.4 cm electroporation cuvette and electroporated as for P. berghei [1]. The electroporated suspension was then injected intravenously into a 16–18 g mouse that in some experiments had been treated 5 days earlier with a single dose of phenylhydrazine hydrochloride (100 mg kg − 1 body weight; dissolved in a 0.9% (w/v) solution of NaCl) to induce reticulocytosis. Starting 27–30 h postinjection, mice were treated with varying concentrations of pyrimethamine until parasites were no longer detected (B 0.001% parasitemia). When the parasitemia increased again to detectable levels, mice were treated with pyrimethamine and resistant parasites transferred

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to new mice to produce parasite numbers sufficient for parasite DNA isolation and analysis.

2.5. Fluorescent confocal microsocopy and FACS analysis of resistant parasite populations Blood from mice infected with pyrimethamine-resistant P. yoelii populations was collected by tail bleeding in PBS-heparin solution. Parasite DNA in infected cells was stained with ethidium bromide to allow distinction between infected and uninfected cells. Pellets of cells (approximately 105 – 106 cells) were incubated for 3–5 min with 50 ml of PBS containing ethidium bromide (1 mg ml − 1), washed twice in PBS and further analyzed by confocal microscopy. For FACS analysis, cells were diluted to an approximate concentration of 106 cells per ml and analyzed in a Becton – Dickinson FACScan machine equipped with an Argon laser tuned at 488 nm.

2.6. Southern hybridization analysis of resistant parasite populations Genomic DNA of parasite populations was prepared as previously described [1]. Southern blotting was performed with the entire TRAP or GFP coding sequence as probes. The probes were labeled with DIG-ddUTP by random priming, and the chemiluminescence was detected by using CSPD (Boehringer Mannheim).

2.7. Analysis of parasite de6elopment and infecti6ity A. stephensi mosquitoes were fed on infected mice and dissected at day 14 post-feeding. Midgut or salivary glands associated sporozoites were obtained as described [12]. To test sporozoite infectivity, 103, 104 and

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105 salivary glands associated sporozoites were injected intravenously into mice and their parasitemia was checked daily by Giemsa-stained blood smears. To analyze sporozoite motility, sporozoites were maintained in 3% BSA-RPMI medium for up to 3 h at 4°C, followed by microscopic observation. To determine the proportion of internalized sporozoites among salivary gland-associated sporozoites, intact salivary glands were dissected out at day 14 post-feeding and incubated in trypsin (1.25 mg ml − 1) in RPMI medium for 10–15 min at 37°C. Cells were then centrifuged for 3 min, the supernatant containing the attached sporozoites was collected, and the salivary glands were then ground to free internalized sporozoites. The numbers of sporozoites in each fraction was counted using a hemocytometer.

3. Results and discussion

3.1. P. yoelii 17X sensiti6ity to pyrimethamine Transformation of asexual blood stages of Plasmodium spp. has been successfully achieved using the selectable marker DHFR-TS bifunctional enzyme in a pyrimethamine-resistant form. In order to study the in vivo sensitivity of P. yoelii 17X to pyrimethamine, groups of 3 Balb/c mice infected with P. yoelii 17X were treated with different concentrations of the drug and their parasitemia followed for 3–4 weeks post-infection. As shown in Fig. 1, P. yoelii 17X is highly sensitive to pyrimethamine. At doses higher than 1 mg kg − 1 body weight, two consecutive treatments were sufficient to eradicate the infection, and at doses lower than 1 mg kg − 1 body weight, treatment had to be prolonged for 2 or 3 more days.

Fig. 1. Pyrimethamine sensitivity of P. yoelii 17X blood stages. Groups of three mice were treated with different concentrations of the drug (0.2–10 mg kg − 1 body weight). All data are expressed as the arithmetic means of the three mice in each group. Arrows represent the days of treatment for each group.

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Fig. 2. Stable transfection in P. yoelii 17X blood stages under pyrimethamine treatment. (A – C) Fluorescent confocal microscopy analysis of PyEPI-GFP population stained with ethidium bromide. GFP fluorescence is represented in green (A and C) and ethidium bromide in red (B and C). (D – F) Flow cytometry analysis of an original PyEPI-GFP population (D) and a population transferred into new mice (E and F) stained with ethidium bromide. The recipient mice were not treated with pyrimethamine. E and F represent 2 and 4 days after transfer, respectively.

3.2. In 6itro culti6ation and gradient separation of P. yoelii blood stages leads to purified schizont preparation for transformation In the P. berghei transfection system schizonts are the targets of the electroporation process. P. berghei schizonts are easily obtained after overnight culture [13]. However, when P. yoelii 17X parasites were cultured for 16 h (as previously described for P. berghei ), a significant proportion of parasites consisted of extracellular merozoites, or rings/young trophozoites probably resulting from a recent invasion. It is well known that, unlike P. berghei schizonts, P. yoelii schizonts rapidly rupture in vitro and the new merozoites are capable of invading available reticulocytes. To determine the optimal time to obtain maximum numbers of P. yoelii schizonts, smears were prepared every other hour between a time window of 8 – 14 h after the start of the culture. In three independent experiments, the highest numbers of schizonts were obtained after 12 h in vitro (data not shown). Next we determined the Nycodenz/PBS gradient conditions for purifying schizonts. Consistently, a 55% Nycodenz/PBS gradient in PBS led to the loss of up to 90% schizonts; a 65% Nycodenz/PBS gradient yielded a high number of schizonts, but contained a large pro-

portion (\ 10%) of mononuclear cells and of other P. yoelii stages. As a compromise, we chose a 60% Nycodenz/PBS gradient, the yields were somewhat smaller, but contamination less significant (data not shown).

3.3. Stable transformation of P. yoelii P. yoelii 17X merozoites were transfected with plasmid pMD205-gfp that contains a pyrimethamine-resistant form of P. berghei dhfr-ts gene fused to gfp-mut2 gene [10]. Newly invaded parasites were readily detectable 4 h after injection of the electroporated parasites into mice. Two days after pyrimethamine treatment (5 mg kg − 1 body weight, starting 27 h postinoculation), parasitemias were undetectable. A pyrimethamine-resistant and fluorescent population (PyEPI-GFP) emerged 5 days post-transfection, and expanded under drug pressure of 5 mg kg − 1 body weight of pyrimethamine. These parasite populations were transferred to new mice and the pyrimethamineresistant parasites were collected for further analysis. We found that resistant parasites contained autonomously replicating plasmids that were maintained under drug pressure. The presence of intact autonomously replicating plasmids was confirmed by rescuing them from resistant parasites, transforming

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Escherichia coli with the DNA and subjecting the plasmid DNA to restriction analysis (not shown). In P. berghei, the plasmid pMD205-gfp is maintained in the absence of pyrimethamine selection [10]. However, this is not the case in P. yoelii. In the experiment shown in Fig. 2, blood infected with resistant parasites was stained with ethidium bromide (EB) and analyzed by flow cytometry to distinguish between infected and non-infected erythrocytes (Fig. 2). In the initial pyrimethamine-resistant population, 95% of infected erythrocytes (EB positive) were also GFP positive (Fig. 2D). This parasite population was

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then transferred into naive mice that had not received drug treatment. Two days later, only approximately 55% of the infected population was GFP positive (Fig. 2E) and 4 days later, less than 1% of the infected cells were GFP positive (Fig. 2F). As a control, mice infected with an identical parasite population were treated with pyrimethamine daily. Flow cytometry analysis showed that all infected erythrocytes were fluorescent. Therefore, this plasmid is stable in P. yoelii only under drug pressure, as has also been reported for other autonomously replicating plasmids in P. berghei and P. falciparum [14 –16].

Fig. 3. Gene targeting at the trap locus of P. yoelii and generation of clones PyINT-GFP1 and PyINT-GFP2. (A) WT trap genomic locus and integration generated by integration of plasmid pPyTRAPINT via a single cross-over between the trap homologous sequences. Plasmid pPyTRAPINT was linearized at the unique I site of the trap coding sequence. (B) Genomic Southern hybridization of WT P. yoelii 17X and clones PyINT-GFP1 (C1) and PyINT-GFP2 (C2). Digestions with AflII and HindIII, both cutting once in pPyTRAPINT but not cutting in trap coding sequence, show that the unique band from WT is transformed in two, when hybridized with trap probe. The addition of the size of the two bands, is equal to the size of WT added to pPyTRAPINT, as expected. The same blot hybridized with GFP probe demonstrates the positions of AflII and HindIII restriction sites in the locus.

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Table 1 Phenotype of WT and P. yoelii TRAP null mutant A. stephensi mosquitoesa Parasite population

WT PyINT-GFP1 a

Number of sporozoites/infected mosquito

Typical gliding

MG

SG total

SG external

SG internal

SG (%)

41 000 42 000

25 700 5400

4900 5800

20 100 250

73 0

Each value represents the mean of three independent dissections. MG, midgut; SG, salivary gland.

3.4. Transformation of P. yoelii with linear plasmids leads to site-specific integration of DNA into the genome by homologous recombination We targeted the P. yoelii 17X trap locus (PyTRAP, thrombospondin-related anonymous protein) using an insertion plasmid pPyTRAPINT (Fig. 3A) linearized at a unique SpeI site present in the PyTRAP open reading frame. In two independent experiments, the construct was electroporated into 1 – 5×107 schizonts and injected into phenylhydrazine-treated mice, followed by selection for pyrimethamine resistance with 0.4 mg kg − 1 body weight. Phenylhydrazine-treatment induces reticulocytosis and enhances the invasion rate of the P. yoelii 17X merozoites. A drug-resistant parasite population (PyINT-GFP) appeared on day 6 post-electroporation and was recovered in both experiments. When parasitemias reached 3–5%, parasites from both populations were collected for genomic DNA analysis, and also cloned by limiting dilution. By Southern blot analysis the parental populations contained a very large proportion (approximately 80%) of integrants at the TRAP locus (not shown). This was confirmed by the analysis of five different clones obtained by limiting dilution of one of the parental populations. Four of the five clones consisted exclusively of parasites that had undergone integration while the fifth clone was a mix of wild type (WT) and integrant parasites. Two of the four clones were analyzed further. Southern blot analysis with TRAP and GFP probes showed that the clones (PyINT-GFP1 and PyINT-GFP2) contained a single copy of the entire pPyTRAPint integrated at the genomic trap locus (Fig. 3B). As expected, these clones contained two truncated copies of the trap gene, the first lacking the last 196 nucleotides of the gene and the downstream untranslated region, and the second lacking the 5% promoter sequences and the first 56 nucleotides of the gene (Fig. 3).

3.5. Phenotype of the P. yoelii 17X TRAP null mutant lines The phenotype of the P. yoelii TRAP knockout was identical to that of the P. berghei knockout [12]. In short, since P. yoelii mutants were selected during the erythrocytic cycle, TRAP is not necessary for development of

these stages of the parasite. The numbers of mature gametocytes were also similar in the WT and in TRAP null mutant clones (not shown). To study their sporogonic cycle, A. stephensi mosquitoes were fed on mice infected with WT or PyINT-GFP1 populations bearing high levels of gametocytes. The percentage of infected mosquitoes (\ 95%) as well as the number of oocysts per infected mosquito (between 50 and 70) were identical in both populations. At day 8 post-feeding, the number of midgut sporozoites was compared in mosquitoes infected with WT P. yoelii and PyINT-GFP1. Similar numbers were found in both populations (Table 1). In contrast, on days 13–15 post-feeding, the number of salivary gland-associated sporozoites per infected mosquito was approximately six to eight times smaller in PyINT-GFP1 than in WT populations (Table 1). In P. berghei this difference was even greater (approximately 60 times; [12]). Salivary glands from both populations were then treated with trypsin to distinguish between sporozoites only attached to the salivary glands from those that invaded the glands. After trypsin-treatment, the glands were washed and then ruptured to free the internal sporozoites. In mosquitoes infected with the WT population, the proportion of total salivary gland-associated sporozoites released from within the glands was 80.49 3.2%. However, only 4.190.4% of the total salivary gland-associated sporozoites were released from within the glands in mosquitoes infected with the PyINT-GFP1 populations. Thus, as in P. berghei, P. yoelii TRAP null mutants fail to invade the salivary glands of mosquitoes. The higher number of salivary gland-associated sporozoites in P. yoelii null mutants as compared with P. berghei TRAP null mutants [12] is probably due to the much higher numbers of sporozoites produced in mosquitoes infected with P. yoelii. Motility assays were also performed, and confirmed that TRAP null mutants do not exhibit gliding motility (Table 1). Salivary glandassociated sporozoites were isolated from mosquitoes at day 15 post-feeding and injected into mice in order to evaluate the infectivity of PyINT-GFP1 sporozoites. As shown (Table 2), TRAP null mutants sporozoites were less infective than the corresponding WT sporozoites. This was assessed by the difference observed in prepatent periods. A pre-patent period is defined as the number of days between injection with sporozoites and the first appearance of parasites in the peripheral blood.

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A delay of a single day in patency implies at least ten times difference in innoculum [17]. Since infection with 104 PyINT-GFP1 sporozoites resulted in a delay of 2.5 days (when compared with WT), we must conclude that TRAP null mutant sporozoites are at least 100 times less infective than WT sporozoites. Polymerase chain reaction (PCR) analysis showed that the blood stage parasites induced by PyINT-GFP1 sporozoites contained only WT trap, with no trace of disrupted trap (not shown), most likely resulting from excision of the plasmid pPyTRAPint by intralocus recombination via the TRAP duplicated sequence. This has been shown to occur in P. berghei when insertion plasmids are used [12]. Stable genetic transformation has been achieved in a human malaria parasite, P. falciparum, but only in vitro studies can be performed in this model. Stable transfection has also been demonstrated in two non-human primate malaria parasites, Plasmodium knowlesi [18] and Plasmodium cynomolgi [19], and two rodent malaria parasites, P. berghei [14,20] and now P. yoelii. While recognizing the value of transfection for the study of P. knowlesi and P. cynomolgi, the ethical, methodological and economical problems involved in using non-human primates prevent their widespread utilization. Given that the maintenance of the entire rodent malaria’s life cycle in vivo is not difficult, rodents continue to be the first choice for studies on the biology of Plasmodium and its interaction with its host. Although there are differences in many aspects of the biology and the pathology of human and rodent malarias, conservation of function of several proteins and domains, of the regulation of gene expression and even of the higher order of chromosome structure has been described (for review see [21]). In particular, and of relevance to vaccine development, no differences have been found to date between biological properties of human and rodent malarias, during the pre-erythrocytic stages of infection. Thus, studies in rodent models can provide useful information about the human malaria parasites. There are several attractive features for using the P. yoelii model. A preliminary annotation at 2X coverage

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of the P. yoelii genome is already available. Since mice are more susceptible to infection with P. yoelii sporozoites than with P. berghei sporozoites, P. yoelii has always been the favored model for studying the mechanisms of immunity to the pre-erythrocytic parasite stages and the development of vaccines [4]. In this context, the major shortcoming of the P. yoelii model has been the inability to perform in vitro studies of the liver stages, since P. yoelii sporozoites do not complete the exoerythrocytic cycle in hepatocytes or any other cell line tested. Recently, however, this problem has been surmounted when it was found that a mouse hepatoma cell line is equally well infected by P. yoelii and P. berghei sporozoites leading to the complete development of exo-erythrocytic forms of both parasites [22]. A transfection system in P. yoelii is also highly desirable for studying the biology of the blood stages. One major question in malaria biology is the molecular mechanism that determines the preference of the different Plasmodium spp. for young or mature erythrocytes, and its contribution to the virulence of the parasites. Some Plasmodium spp. can invade erythrocytes of all ages (P. falciparum and Plasmodium chabaudi ), others prefer mature erythrocytes (P. knowlesi and Plasmodium malariae) and yet others preferentially or exclusively invade reticulocytes (Plasmodium 6i6ax, P. cynomolgi and P. berghei ). P. yoelii is an attractive model for studying the molecular basis of these differences since there are two different parasite lines of P. yoelii, a non-lethal 17X line which prefers reticulocytes and a lethal YM line that invades erythrocytes of all stages of development [9]. A practical advantage of the P. yoelii model is that larger numbers of P. yoelii (25 000 –40 000) than of P. berghei sporozoites (10 000 –20 000) can be isolated from the salivary glands of mosquitoes (A. stephensi ). In addition, P. yoelii 17X is much more susceptible to pyrimethamine than P. berghei NK65 parasite line, that has been used in previous studies, thus simplifying selection of recombinants and shortening the time required to obtain clonal populations.

Table 2 Infectivity to mice of WT and P. yoelii TRAP null mutant Parasite population

WT

PyINT-GFP1

a

Salivary gland-associated sporozoites Number of injected sporozoites

Number of infected animalsa

Pre-patent periodb

103 104 105 103 104 105

3/3 3/3 3/3 0/3 3/3 2/2

2.5 2 2 − 4.5 4

Number of infected animals/number of animals injected with the sporozoite suspension. Number of days between sporozoite injection and detection of at least one erythrocytic stage on 100 microscopic fields of Giemsa-stained blood smear negative, parasitemia undetectable up to day 10 after sporozoite injection. b

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In short, we have demonstrated for the first time the feasibility and ease of stable transfection and gene targeting in P. yoelii using available dhfr-ts based selectable markers. This will permit functional analysis of genes expressed in the exoerythrocytic stages in the context of a favored model used for vaccine development and applicable to human malaria infections.

Acknowledgements

[9]

[10]

[11]

[12]

We thank John Hirst for the FACS analysis and Ivette Caro for the mosquito dissections. We dedicate this work to Dr Michael Gottlieb. The work was supported by a grant from the National Institutes of Health (AI43052).

[13]

[14] [15]

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