Transfection of rodent malaria parasites

Chapter 7 Transfection of Rodent Malaria Parasites Nisha Philip, Rachael Orr, and Andrew P. Waters Abstract Gene manipulation is an invaluable tool to investigate and understand the biology of an organism. Although this technology has been applied to both the human and rodent malarial parasites (RMP), Plasmodium berghei in particular offers a more robust system due to a higher and more efficient transformation rate. Here, we describe a comprehensive transfection and selection protocol using P. berghei including a variant negative selection protocol administering 5-fluorocytosine to the animals in drinking water. Additionally, we discuss and assess the latest advances in gene manipulation technologies developed in RMP to gain a better understanding of Plasmodium biology. Key words: Plasmodium berghei, Transfection, Reverse genetics, Forward genetics, Episomes, Selectable markers, Promoter swap, Artificial chromosome

1. Introduction In the last decade, large-scale sequencing of Plasmodium genomes followed by global gene expression arrays, parallel RNA sequencing, and proteomic studies have generated a wealth of information aiding a greater understanding of Plasmodium biology (1–9). Despite these advances, “hypotheticals” still encompass about 50% of the ~5,000 annotated genes. Consequently, genetic manipulation is necessary for assigning (and confirming) gene function in order to understand their role in various aspects of parasite biology, especially Plasmodium-specific attributes. In this respect the rodent malaria parasites (RMP) have been particularly useful notably Plasmodium berghei where stable genetic manipulation was first established (10), although P. yoelii (11) and P. chabaudi (12) are also somewhat amenable to genetic manipulation. Apart from relative ease of use, RMP offer a number of significant advantages:

Robert Ménard (ed.), Malaria: Methods and Protocols, Methods in Molecular Biology, vol. 923, DOI 10.1007/978-1-62703-026-7_7, © Springer Science+Business Media, LLC 2013

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their genomes are fundamentally homologous to those of the human parasites across the majority of the genome excluding the subtelomeric clusters of (typically) multigene families. Approximately 80% of the annotated genes in the genome, which typically contains ~5,500 genes, can be considered to be syntenic (13), permitting gene function to be assessed in stages that are difficult or impossible to access in human Plasmodium. They allow experimental procedures that cannot be considered with human subjects and offer experimental access to the entire life cycle. Different virulence patterns and pathologies might also be studied with RMP models. A range of reporter RMP are now available that, for example, express fluorescent proteins either constitutively or stage specifically and permit imaging and purification processes for further phenotypic or biochemical analysis (14–16). Furthermore, both host and vector may now be genetically manipulated, allowing a consideration of all aspects of parasite interactions with those it infects. RMP are limited in a number of respects, notably the inability of the blood stages to be cultivated long term in vitro, restricting study of invasion of and egress from the erythrocyte, and the relative lack of selectable markers with which to select manipulated parasites. A further general consideration is that Plasmodium parasites are haploid organisms, which is both an experimental advantage and disadvantage when considering reverse genetics. The major advantage of haploidy is that only a single allele needs to be deleted/modified, which is a substantial benefit given the paucity of positive selectable markers available. However, the problem that arises from haploidy is that genes that are essential to the vegetatively growing phase of the parasite in blood, which are used to propagate schizonts for transfection, cannot be studied since deletion or disabling through modification would be a lethal event and such transfected parasites would perish prior to selection. As noted above, asexual blood-stage growth of Plasmodium includes a series of events (parasite egress, erythrocyte invasion, and cytoadherence) that are of intense interest from therapeutic considerations, and the inability to study these processes through gene deletions mediated by conventional reverse genetics is a significant handicap requiring other approaches, some of which are described below. Nonetheless the haploid genome of the asexual blood stage of the parasite is amenable for genetic manipulation and is therefore used for transfection. However, introduction of exogenous DNA into the intracellular Plasmodium parasite is problematic because traversal of four membranes is required to reach the nucleus. In P. berghei, use of purified schizonts/merozoites partly avoids this problem, which when combined with advances in transfection technologies due to development of Amaxa nucleofection has resulted in much higher transfection frequencies (frequency of 10−3–10−4 compared to 10−6–10−9 using past methodology) than

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typically achieved with the human parasite, P. falciparum (17). In addition to two detailed transfection and selection protocols, we discuss various gene manipulation strategies the technique can be applied to. 1.1. Reverse Genetics

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Gene inactivation or modification is the most reliable and useful approach to examine gene function in parasite biology. Given that Plasmodium exclusively incorporates DNA by homologous recombination, both single and double crossover strategies can be exploited for gene manipulation (see Fig. 1). Insertion plasmids utilizing single crossover require minimally ~300 bp of homology on either side of the point of recombination (site of linearization in the plasmid) to integrate into the targeted genomic locus. Larger

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Fig. 1. Gene manipulation strategies: gene inactivation and modification can be achieved by both double and single crossover homologous recombination. (a) Gene knockouts can be generated using plasmids containing a positive selection marker (SM) flanked by arms homologous (dashed lines) to regions upstream and downstream of the gene of interest. The construct is linearized to generate a fragment with the SM and regions of homology; when transfected into the parasite, it undergoes double crossover leading to allelic exchange of the wild-type copy of the gene with the SM resulting in a gene knockout. (b) Gene disruption using insertion plasmids containing an internal fragment of the gene occurs by a single crossover event. The plasmid is linearized (gap) within the homology region, which recombines into the parasite genome to generate two truncated copies of the target gene. This process can be reversed with negative selection using 5-FC when the yFCU element is fused to the positive selectable marker. (c) Gene modification including N- and C-terminus tagging can be accomplished by constructs designed for double crossover events. The plasmid would contain an upstream homology region, followed by the SM, gene promoter, 5¢ UTR, the modified targeted gene, and finally the downstream homology region. (d) Insertion plasmids using single crossover recombination can be used to modify genes and introduce C-terminal tags. The construct contains a partial C-terminus of the modified gene of interest followed by a 3¢UTR. The linearized plasmid recombines at the targeted locus to generate a promoter-less truncated gene and a full-length functional modified gene of interest. This process can be reversed with negative selection using 5-FC when the yFCU element is fused to the positive selectable marker.

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regions of homology tend to make the recombination event more efficient/frequent. Such plasmids can be used for introducing subtle mutations and also for gene inactivation. Alternatively, linear replacement constructs containing a drug-selectable marker bordered by two arms of homology, which integrate into the genome by double homologous recombination, can be employed for gene modification and deletion. However, despite significant improvements in the technology (18) systematic functional genomic analysis is hampered by the relatively low efficiency of homologous recombination in the parasite and difficulty of producing targeting plasmids containing A/T-rich Plasmodium DNA in Escherichia coli. A recent development from the Billker group attempted to address these issues (see Chapter 8) employing a recombineering strategy that uses bacteriophage N15-derived library to stably maintain fragments of P. berghei gDNA up to 16 kb in size. Phage clones are manipulated to produce transfection-competent vectors, which through the use of long homology arms achieve over tenfold increased transfection efficiency compared to conventional methods. The approach was further scaled-up to generate P. berghei gene modification vectors in a 96-well format achieving 88% success (86/96 constructs). A genome-wide library of genetic modification vectors capable of gene deletion and tagging now makes it possible to perform large-scale functional studies of Plasmodium genes. 1.2. Selectable Markers

It was noted above that reverse genetics in P. berghei is hindered by the low number of selectable markers due to toxicity issues in the rodent host. Currently, dihydrofolate reductase-thymidylate synthase (DHFR-TS), an enzyme involved in folate metabolism, is commonly used as a drug selectable marker. The P. berghei (Pb) and Toxoplasma gondii (Tg) DHFR-TS confer resistance to pyri-methamine, a drug now administered in drinking water (19). The human DHFR gene, in addition to pyrimethamine, also confers resistance to WR99210 (20). Combination of the two markers is possible only if Pb/Tg DHFR-TS (first) and human (second) DHFR are introduced sequentially due to resistance of human DHFR to both pyrimethamine and WR99210. The scarcity of selectable markers can be somewhat mitigated by recycling them using a positive–negative selection strategy. A positive–negative selectable marker system can be applied for (1) complementing disrupted genes (generated using positive selection), followed by excision of the drug selection cassette by homologous recombination (negative selection); (2) counter-selecting episomal transformation and single crossover events to increase recovery of double crossover recombinants; (3) developing a hit-and-run strategy for generating mutations without introducing exogenous sequences into the targeted locus; and (4) recycling both the positive and negative selection markers to generate additional sequential mutations (e.g., double, triple). A negative selection system based on a yeast bifunctional enzyme (yFCU)

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consisting of cytosine deaminase and uracil phosphoribosyl transferase has been developed for both P. falciparum and P. berghei (21, 22). Negative selection pressure is applied by 5-fluorocytosine (5-FC), which is converted by yFCU into a highly toxic compound 5-fluorouracil that kills yFCU-expressing parasites within 48 h. In P. berghei, pure populations of negatively selected parasites are recovered within 5–6 days post 5-FC treatment. This system was successfully utilized to restore gene function by homologous recombination and excision of the selectable marker at the disrupted gene locus. The strategy was further developed to recycle the positive selectable marker by flanking the selection cassette by two 0.5-kb PbDHFR-TS 3¢ regions that under 5-FC pressure recombine and excise the selectable marker. Additionally, in P. falciparum the method was adapted to use site-specific recombinases such as the Flp/FRT system (23). This chapter also describes a modified detailed protocol for negative selection using 5-FC in drinking water, which offers significant improvements in labor, drug use, animal welfare, reduction in animal use, and reproducibility. 1.3. Promoter Swap

A common problem noted above is that genes that are essential in blood-stage asexual parasites cannot be deleted due to their essential nature, and in the absence of robust conditional knockout (ko) strategies this remains an issue. However, the function of these essential genes can be examined in later stages of the parasite life cycle through an approach where the wild-type promoter, which might be active in multiple stages of the life cycle, is swapped for one that is active only in intraerythrocytic asexual stages. In this manner recombinant parasites that have undergone this type of promoter swap for an essential gene are viable in the asexual blood stages where their selection and cloning take place, yet gene expression terminates once the parasites commit to sexual development and thereafter. Therefore, the function of the promoter-swapped gene can be established at its first point of phenotype appearance after (or during) gametocyte development. This approach has been taken to study the FACT (facilitates chromatin transcription) protein—four promoter swap constructs using four different promoters were introduced into the parasite and promoters from two genes (PBANKA_091420 and PBANKA_140060) gave viable parasites and were used to demonstrate the critical role of FACT in fertile male gamete formation (24).

1.4. Episomal Transfection

The tools mentioned earlier use either homologous recombination or insertion of foreign DNA into the genome to generate integrative transgenic lines. Alternatively, extrachromosomal or episomal vectors, which do not physically disrupt the genome, can be used for ectopic expression and analyzing gene function. Episomal transfection has been applied towards understanding transcriptional regulation, examining protein localization, performing

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genetic rescue experiments, as well as dissecting molecular pathways. Although episomal vectors containing bacterial derived origin of replication do multiply in the parasite, they have a major drawback of not segregating evenly into daughter cells during cell division (25, 26). Additionally, episomes are lost if not kept under constant drug pressure, which is possible only during the parasite asexual stages. Moreover, under selective drug pressure plasmids are maintained in high copy numbers, which might result in elevated gene expression causing non-physiological effects. Difficulties introduced by experimental artifacts and nonsegregation of plasmids can be overcome by designing vectors based on chromosomal elements derived from the parasite genome. The discovery of yeast centromeres, origins of replication, and telomeres led to the construction of the first eukaryotic artificial chromosome (27, 28). Artificial chromosomes have the potential to carry large fragments of genomic DNA, persist in the nucleus without selection pressure, maintain low copy numbers, and segregate evenly during mitosis and meiosis. The recent identification of the P. berghei centromere and development of a Plasmodium artificial chromosome (PAC) offers a significant advance in extrachromosomal transfection technology (29). Iwanaga et al. initially constructed a P. berghei centromere-containing circular plasmid capable of efficiently replicating and maintaining itself during cell division in the asexual blood stages, and zygote/ookinete and oocyst stages in the mosquito. The plasmid was further developed by addition of capped telomeres to generate a linear PAC (L-PAC). The L-PAC is stably maintained throughout the life cycle of the parasite in low copy number (2.1 per cell) and persists in 70% of the parasites after a complete cycle including the mosquito and liver stages. These advantages make the vector an excellent tool to (a) precisely examine transcriptional regulation during any parasite life stage, (b) complement mutant phenotypes without the use of additional selectable markers, and (c) investigate Plasmodium chromosome function. Examining chromosome function, especially telomere biology, is key to understanding regulation of Plasmodium multigene families. These gene families are enriched in sub-telomeric regions and encode variable surface antigens (e.g., var), which are targeted to the erythrocytic surface. Expression of var genes is mutually exclusive with only a single member active at a given time (30, 31). Additionally, switching of expression of the different var genes is used by the parasite to evade host immunity and promote parasite survival. Mechanisms controlling antigenic variation include histone modification, exchange of canonical histones with variant forms, and subnuclear compartmentalization (32–34). Given that L-PAC functions as natural parasite chromosome, the vector originally developed in P. berghei provides a powerful tool to better understand this complex mechanism of gene regulation in Plasmodium.

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1.5. Conditional Mutagenesis: Roles of Essential Genes?

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The problems of studying essential genes in haploid organisms through reverse genetics have been outlined already. Therefore conditional mutagenesis, usually the interruption of gene/protein function in an inducible manner, would appear to be the ideal solution to these difficulties. To date the methods that have been typically employed for induction of gene inactivation have not been applied to RMP although they have been introduced with moderate success into the human parasite P. falciparum. Tetracyclineinducible expression of transgenes (35) and selective destruction of protein fusions to the ligand-regulatable FKBP protein destabilization domain (ddFKBP—regulated by shield) (36) or to a degradation domain derived from E. coli dihydrofolate reductase (regulated by trimethoprim) (37) have all been achieved in cultured P. falciparum but not yet in RMP, in part due to the difficulties of longterm in vitro cultivation of blood stages. However, transmission of RMP through Anopheles mosquitoes is well established as is the ability to create novel genetic crosses through the in vitro fertilization of gametes derived from distinct lines of P. berghei and their subsequent passage through mosquitoes and recovery upon bite back on a mouse. This has been cleverly exploited to develop FLP recombinase-mediated stage-specific excision of blood stage-required genes in the mosquito with subsequent analysis downstream in the infected mouse. In principle, genes of interest (GoI) are flanked first by the FRT repeat sequences (substrates for the Saccharomyces cerevisiae FLP recombinase) in a fashion that upon excision brings GFP under control of the promoter of the GoI, making the parasites of interest identifiable. In early forms of this approach the feasibility was established using a strategy where the recombinase was expressed in a stage-specific manner (the late oocyst- and sporozoite-specific promoter TRAP) in an independent line, and gamete crossing performed to create a subpopulation that contained the correct combination of targeted locus and FLP recombinase (38). Latterly and building on this proof of principle, P. berghei lines have been developed that express a form of FLP recombinase (FlpL) that is maximally active at the ambient temperature of the mosquito habitat. Furthermore FLP recombinase is placed under either a midgut or a salivary gland sporozoite-specific promoter and GoI are FRT-flanked in the same line (39). The efficiency has been shown to be as high as 95%, obviating the need for the use of GFP expression to detect the experimental parasite subpopulation and allowing assignation of function downstream in liver stages. The essential and unknown role of a gene involved in merozoite formation in hepatocytes (MSP1) has been demonstrated in this fashion, expanding our knowledge of the function of this protein that was previously shown to be essential for merozoite recognition of erythrocytes. It should be noted that for all gene disruption methods, the limitation remains that if the GoI is essential at any point in the life cycle then its role in

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Plasmodium life cycle events downstream of the first critical point of function cannot be studied. Analysis of genes essential to blood stages can be facilitated by this FLP-mediated stage-specific approach to excision of GoI. However, some genes that are essential in blood stages are also critical at other stages as they play a conserved role in the invasive organelles shared by merozoite and sporozoite, for example AMA1. Although the recent developments described above and elsewhere have made it feasible to perform larger scale gene-by-gene disruption analysis, the large number of hypotheticals, some of which have already been shown to be responsible for parasite-specific processes (40), provides a challenge to making educated predictions of gene function. A complementary unbiased phenotype-driven approach might provide insights into critical aspects of parasite biology. 1.6. Forward Genetics

Forward genetics has the potential to be a powerful medium/ high-throughput approach to identify genes responsible for observable alterations in phenotype. The forward genetics approach, unlike reverse genetics, offers a valuable unbiased tool that is able to identify multiple genes as well as noncoding genomic regions responsible for a given phenotype. Variations in phenotype can be achieved either by random (chemical, radiation, and insertional) or site-specific mutagenesis. The first study using transposable elements in Plasmodium followed a random mutagenesis approach and employed the Class II transposable element mariner, which moves in the genome using a cut and paste mechanism, in the presence of a transposase enzyme. Unfortunately in P. falciparum mariner proved to be quite inefficient and its use has not been reported in other species of Plasmodium. Other transposable elements including P-element, Tc1, Tol2, and piggyBac have been extensively used to study gene function in model organisms (41–45). Site-specific insertional mutagenesis employing Tn5 shuttle mutagenesis has also been attempted (46). However, this approach involved cloning every targeted gene into E. coli rendering it a long and cumbersome procedure. Recently, the piggyBac transposable element, which inserts at TTAA sites in the genome, was adapted for both human and RMP (47, 48). The piggyBac system was first developed in P. falciparum and applied to a phenotypic screen to identify genes required for the intraerythrocytic development cycle of the parasite (49). The system was adapted to a dual-plasmid system by separating the gene encoding the transposase enzyme and the integrating asymmetric repeats that normally flank the transposase. The asymmetric repeats flanked a positive selectable marker, which allowed selection for stably transformed parasites when integrated through the activity of the transposase, whose gene was transiently introduced

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on the co-transfected helper plasmid. Eighty-one independent transfections yielded about 180 unique insertion sites (all characterized by the AATT recognition site) randomly distributed across the 14 chromosomes but with a preferential bias towards 5¢ UTR regions. Although the strategy has a high efficiency of random insertion, P. falciparum genome manipulation is however plagued by very low transfection efficiency, making such mutagenesis unlikely to achieve saturation levels. Taking advantage of the higher transfection efficiency in P. berghei, Fonager et al. further developed the piggyBac system in the RMP (48). Compared to P. falciparum, the piggyBac system applied to P. berghei system yielded 16–18-fold more insertion sites at AATT repeats in the genome. Additionally no 5¢UTR bias was observed and insertions were randomly spread throughout the genome. Stable integration of the transposase gene into the genome was also tested for persistent expression of the enzyme, anticipating that this would promote further transposon mobility. This mobility can potentially be exploited for reversion of the mutant phenotype to establish a causal relationship between the mutation and the observed phenotype. Remobilization can also be used for transposon excision and reinsertion into other genomic loci, increasing the potential coverage of the transposon from a single transfection experiment. A further advance of the system was to identify promoters active during asexual stages. A “promoter trap” plasmid construct was designed consisting of a piggyBac 5¢ element followed by a promoter-less GFP cassette, where GFP expression was observed only when the construct inserted into transcription units downstream of active promoters. This approach can be further employed to identify promoters active during other developmental stages. In the last decade the malaria research community has achieved significant technological advances in understanding gene function in the malarial parasite. Altogether, improved transfection frequency, increased site-specific recombination efficiency, conditional mutagenesis, transposon mutagenesis, and centromeric plasmids have provided the means to considerably facilitate examination of parasite biology. Table 1 summarizes the available gene manipulation tools and their uses. In spite of these technical advances, the Plasmodium molecular genetic toolbox misses some key pieces. Development of robust conditional knockout/inducible strategies is critical to understand functions of genes essential during bloodstage growth. The very essentiality of these genes makes them attractive targets for vaccine and drug development. Further development of strategies to regulate gene expression both post-transcriptionally and post-translationally would provide valuable tools to understand the druggable parasite genome and develop new intervention tactics.

Allows identification of genes essential for blood-stage forms Can be used to set up medium-throughput phenotypic screens Generates stable mutations

Allows high-throughput phenotypic screens Laborious to achieve saturation mutagenesis levels Can be used for efficient promoter trap screens Generates stable mutations

Shuttle mutagenesis

PiggyBac

Stable episomal expression of transgene Evenly segregates at low copy number into daughter parasites allowing for precise examination of biological processes

Conditional mutagenesis allowing functional Precise expression of recombinase examination of essential blood-stage genes essential in other parasite life cycle stages

Similar advantages to temporal recombinase Choice of promoter is crucial to but requires only a single plasmid avoid leakiness and to maintain construct gene expression close to endogenous levels during other life cycle stages

Centromeric plasmids and Plasmodium artificial chromosomes (PAC)

Temporal recombinase expression

Promoter swap

Transgene expression only Stability of large inserts in current formulations untested

Involves complex cloning procedures in Escherichia coli

Low throughput Relatively low efficiency in the absence of long homology arms

Allows for precise replacement or modification of gene Generates stable mutations

Double crossover homologous recombination with linear fragment containing two flanking regions of homology

Variety of promoters available to avoid artifacts or lethality through overexpression

Combination with recombineering could create PAC libraries Potential for complementation/ dominant negative studies

Current standard approach

Low throughput Reselect with drug if worried Unstable, integration event can revert to give wt + original episome

Allows for gene silencing and modification

Comments

Single crossover homologous recombination with linearized episome

Disadvantages

Advantages

Tool

Table 1 Plasmodium berghei transfection toolbox

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2. Materials All experimental procedures using animals should conform to the relevant local, national, and European legislations. Procedures must have been given ethical clearance. Consult your local animal welfare officer for more information. 2.1. Phenylhydrazine Treatment of Schizont Donor

1. Phenylhydrazine hydrochloride (Sigma-Aldrich): 12.5 mg/ml phenylhydrazine in 0.9% NaCl solution (see Note 1). 2. 25 or 26G needle. 3. 1-ml Syringe. 4. Theiler’s original (TO) or NIH Swiss outbred mouse, female, 25 g, aged 6 weeks.

2.2. In Vitro Culture of Schizonts

1. Microscope slides with frosted end. 2. Methanol. 3. Giemsa stain. 4. Sörensen staining buffer: KH2PO4 2.541 g, Na2HPO4·2 H2 0.5507 g per 5 L of dH2O, pH to 7.2 with NaOH (see Note 2). 5. Heparin: 200 U/ml prepared in PBS, filtered with a 0.22-mm filter device. 6. Fetal bovine serum (FBS) (heat inactivated). 7. Complete culture medium RPMI1640 with l-glutamine and 25 mM HEPES without NaHCO3: weigh 0.85 g NaHCO3, add 5 U/ml penicillin and 5 mg/ml streptomycin, pH to 7.3, filter with a 0.22-mm filter device, and supplement with 25% fetal calf serum. 8. Gas mixture: 5% CO2, 5% O2, and 90% N2. 9. Cell culture 150-cm2 plug seal flask. 10. Orbital shaking incubator. 11. Light microscope with 100× lens plus immersion oil.

2.3. Purification of Mature Schizonts

1. Nycodenz stock solution: 138 g of Nycodenz powder (Axisshield) dissolved in 500 ml buffered medium, autoclaved at 120°C for 20 min (see Note 3). 2. Buffered medium: 5 mmol/l Tris–HCl at pH 7.5, 3 mmol/ l KCl, 0.3 mmol/l CaNa2EDTA. 3. PBS. 4. Table-top centrifuge with swing out rotor and no brake capability.

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2.4. Electroporation of Schizonts

1. Basic parasite Nucleofector Kit 2: solution 2, supplement 3, cuvettes, pipettes (Lonza). 2. Amaxa Nucleofector II Device (Lonza). 3. Purified schizonts obtained from Subheading 3.3. 4. Culture media from the top of Nycodenz centrifugation gradient tube. 5. 5–10 mg of construct in 10 ml of dH2O or TE.

2.5. Intravenous Injection of Transfected Schizonts

1. 0.3-ml Insulin syringe with 30 G needle (BD biosciences).

2.6. Selection of Transfected Parasites

1. Pyrimethamine (Sigma-Aldrich): stock (100×) is 7 mg/ml dissolved in DMSO (see Note 4).

2. Hot box or infrared heat lamp. 3. TO or NIH Swiss outbred mouse, female, 25 g, aged 6 weeks, one per transfection reaction.

2. WR99210 in DMSO (gift from Jacobus Pharmaceuticals, Princeton, NJ. Drugs). 3. Darkened or opaque drinking bottles. 2.7. Monitoring the Growth of Transfected Parasites

1. Microscope slides with frosted end. 2. Methanol. 3. Giemsa stain. 4. Sörensen staining buffer, as above in Subheading 2.2, item 4.

2.8. Collection of Parasites for Cryopreservation and for Isolation of DNA for Analysis

1. Cryotubes. 2. PBS. 3. Heparin: 200 U/ml prepared in PBS, filter with a 0.22-mm filter device. 4. Plasmodipur filter (EuroProxima). 5. Zeba spin column, empty 10-ml (Fisher) (see Note 5). 6. Whatman CF-11 cellulose powder (Fisher) (see Note 5). 7. Erythrocyte lysis buffer: 10× stock, 1.5 M NH4Cl, 0.1 M KHCO3, 0.01 M EDTA. This should be prepared to 1× working stock in prechilled dH2O. 8. 30% glycerol/PBS (v/v) with heparin 10 U/ml.

2.9. Preparation of Agarose Blocks Containing Chromosomes

1. 50-Well disposable plug molds (BioRad). 2. TNE buffer: 50 mM Tris, pH 8.0, 100 mM NaCl, 5 mM EDTA. 3. Low-melting agarose (Sigma-Aldrich): 15 mg/ml TNE buffer. 4. Proteinase K: 20 mg/ml. 5. SE buffer: 0.5 M EDTA, 1% sarcosyl lauroyl sulphate, pH 8.0.

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2.10. Cloning of Transfected Parasites

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1. Giemsa-stained thin blood film. 2. PBS. 3. Hemocytometer. 4. Heparin capillaries. 5. Light microscope. 6. 0.3-ml Insulin syringe, 30 G needle (BD biosciences).

2.11. In Vivo Maintenance and Production of Transgenic Parasites Containing Episomes

1. Pyrimethamine (Sigma-Aldrich): stock 100× is 7 mg/ml dissolved in DMSO (see Note 4).

2.12. Negative Selection Using 5-Fluorocytosine In Vivo Provided in the Drinking Water

1. 5-FC (Sigma-Aldrich): 1 mg/ml in water (see Note 6).

3. Methods 3.1. Phenylhydrazine Treatment of Schizont Donor (Day 0)

A mouse is infected with P. berghei to serve as a source of bloodstage parasites for the culture and purification of schizonts. The schizont is the developmental stage that is used for introduction of foreign DNA by electroporation. Phenylhydrazine treatment results in the induction of reticulocytosis. 1. A TO outbred mouse of 25 g is intraperitoneally (i.p.) injected on day 0, typically a Wednesday, with 0.1 ml of 12.5 mg/ml phenylhydrazine hydrochloride (see Note 1). 2. On day 2 the phenylhydrazine-treated mouse is i.p. infected with 0.2 ml of a thawed suspension of a cryopreserved stabilate (see Note 7). 3. On day 4 (Monday) or 5 (Tuesday), Giemsa-stained thin blood films are prepared from a droplet of tail blood to detect the level of parasitemia, generally between 0.5% and 10% at this stage (see Note 2).

3.2. In Vitro Culture of Schizonts (Day 5)

Blood stages of P. berghei are cultured in RPMI1640 medium (pH 7.3) containing FBS. In general, parasites are maintained in vitro for only one developmental cycle: ring forms/young trophozoites are allowed to develop into mature schizonts during a period of 16–23 h (see Note 8).

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1. Collect 1–2 ml of infected blood by cardiac puncture using a 2-ml syringe from a mouse with a parasitemia of 1–10% (see Note 9). Usually the blood is collected between 10 a.m. and 4 p.m. on day 4 or 5. 2. Transfer the blood to 40 ml complete culture medium per ml of blood collected, doubling this volume if infection is above 3%. It is important, however, to ensure that there are no multiply infected erythrocytes if parasitemia is above 3% (see Note 10). 3. Cultures can be maintained in closed, plastic culture flasks of 150 cm2 that have been gassed at the start of the culture period using the gas mixture 5% CO2, 5% O2, and 90% N2. There should be no more than 80 ml of culture per flask; therefore any volume greater than this should be split equally between multiple flasks (see Note 11). 4. Put the flask on a shaker in a 37°C water bath, incubator, or climate room. 5. Switch on the orbital shaker after 5 min at a minimal speed (20–30 rpm) to keep the cells in suspension. 6. Leave the parasites in culture at 37°C until the next morning 9.00 a.m. (see Note 12). 7. Take 0.5 ml from the culture in an Eppendorf tube at 9.00 a.m. to determine the “quality” of the parasites. 8. Pellet the cells (maximum speed, 5 s) and discard the supernatant. 9. Make a thin blood smear of the cells and Giemsa-stain the slide. 10. Examine schizont development using a light microscope at a 1,000× magnification (100× objective, immersion oil) (see Note 13). 11. Start the purification of the schizonts. 3.3. Purification of Mature Schizonts (Day 6)

Prior to electroporation, the schizonts (1–3% of the total cell population) are separated from the uninfected erythrocytes that are present in the culture. 1. Prepare a 55% Nycodenz/PBS solution (v/v). In general a total volume of 30 ml, with 16.5 ml Nycodenz stock solution and 13.5 ml PBS, is required for a culture suspension of 80 ml (see Note 14). 2. Transfer the culture suspension containing the schizonts to two 50-ml tubes (35–40 ml per tube). 3. Using a 10-ml pipette, gently add 10 ml of the Nycodenz solution in each tube under the culture suspension so that a sharp contrasting division is visible between the two suspensions (see Note 15). 4. Centrifuge for 20–30 min 450 ´ g at 1,500 rpm in a swing out rotor at room temperature (RT) with no brake.

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5. Collect carefully the “brown” (grey) layer at the interface between the two suspensions using a plastic Pasteur pipette. 6. In general a total volume of about 15–30 ml is collected from the two tubes (see Note 16). Do not discard the gradient tube as the media from the top is reused later in this protocol. 7. Pellet the schizonts by centrifugation at 1,500 rpm for 8 min. For this “washing” step, add to the schizont suspension ~20 ml culture medium obtained from the top of the gradients. 8. Discard the supernatant from the purified schizonts and retain the media from step 6. 3.4. Electroporation of Schizonts (Day 6)

The Amaxa Nucleofector Device II, sourced from Lonza, is used in this protocol. 1. The schizont pellet obtained from one mouse is used for five to eight transfections (see Note 17). 2. Resuspend the schizont pellet in 5–8 ml of culture medium. 3. Transfer the suspension to five to eight Eppendorf tubes (1 ml/ tube). The parasites, 1–3 × 107 schizonts, from one tube are used for one transfection. 4. Pellet the cells by centrifugation (5 s, maximum speed). 5. Discard the supernatant. 6. Add 100 ml of the Human T-cell Nucleofector solution 2 with supplement 3¢-buffer to 5–10 ml of the DNA solution (5–10 mg DNA construct in water or TE buffer), and resuspend the parasites. 7. Transfer the parasite/DNA/buffer solution to a cuvette provided, dispersing any air bubbles with a gentle tap. 8. Transfect using the Amaxa Nucleofector Device with protocol U-033 to electroporate. 9. Add 50–100 ml of culture medium (retained from the earlier gradient tube) to the cuvette immediately after transfection. 10. The 150-ml suspension containing parasites transfected with the Amaxa Nucleofector Device is injected immediately into one mouse.

3.5. Intravenous Injection of Transfected Schizonts (Day 6)

1. Leave the mice at 37 °C for 15–20 min before electroporation of the parasites. The tail veins swell at 37°C, simplifying the intravenous injection procedure with 0.3-ml insulin syringes with 30G needles. 2. Inject the transfected parasites as quickly as possible into the tail veins of mice that have been anesthetized with isofluorane or restrained using a tunnel (consult local guidelines). It is best to carry out this procedure in a timely manner since the veins constrict rapidly after the mouse exits the hot box. Usually the animal is injected between 10.30 and 11.00 am.

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3.6. Selection of Transfected Parasites (Day 7)

Selection of transfected parasites can be carried out using either pyrimethamine for the PbDHFR/TS selectable marker or WR99210 for both the hDHFR and PbDHFR/TS. Where possible, it is preferable to use pyrimethamine due to its ease of application and efficiency in drinking water (see Note 18).

3.6.1. Selection with Pyrimethamine, Provided in the Drinking Water

1. Provide the animals with drinking water containing pyrimethamine (see Note 4) one day after transfected parasites have been injected. 2. Provide the drug for a period of 4–7 days.

3.6.2. Selection with WR99210 in DMSO, Injected Subcutaneously

1. Inject a single dose of 0.1 ml WR99210 solution (ranging from 6 to 20 mg/kg bodyweight) subcutaneously (in the neck) into mice (20 g) (see Note 19). 2. Repeat the treatment on the following 2–3 days (for a total of three to four treatments).

3.7. Monitoring the Growth of Transfected Parasites (Day 14)

3.8. Collection of Parasites for Cryopreservation and for Isolation of DNA for Analysis (Days 14–18)

1. Parasitemia is checked in Giemsa-stained blood films from day 7 after electroporated parasites have been injected (see Note 20). 2. Parasites are collected from mice when parasitemia is 1–5% and are stored in liquid nitrogen (cryopreservation) or used for genotype analysis. 1. Cryopreservation is carried out by collecting 0.5 ml of heart blood by cardiac puncture in 0.5 ml of a 30% (v/v) glycerol/ PBS solution with 10 U Heparin/ml. This suspension is transferred to 2 cryotubes, 0.5 ml per tube, kept for 5 min at 4 °C or in an isopropanol cryobox stored at −80°C, and then frozen directly in liquid nitrogen for storage. 2. The rest of the heart blood is collected in 5 ml of PBS to extract DNA for genotype analysis. 3. “Prewash” a Plasmodipur filter (EuroProxima) with 10 ml of culture medium or PBS using a 20-ml syringe placed on top of the filter. Always remove the filter before withdrawing the plunger. 4. The infected blood suspension is passed through the filter using a 20-ml syringe to remove leukocytes (see Note 5). 5. Elute with 15–20 ml of culture medium or PBS. 6. Pellet the infected erythrocytes by centrifugation at 1,500 rpm for 8 min. 7. Remove the supernatant. 8. Lyse the uninfected red blood cell (RBC) by resuspending the RBC pellet in 50 ml of cold (4°C) 1× erythrocyte lysis buffer. 9. Incubate the suspension on ice for 3–5 min. The clarity of the suspension is an indicator of the state of the lysis procedure, i.e., when fully clear the lysis is complete.

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10. Pellet the parasites by centrifugation for 8 min at 1,500 rpm. 11. Remove the supernatant. 12. Resuspend the small parasite pellet in PBS to wash once more by pelleting and remove the supernatant. 13. Remove 15–20 ml of the pellet to mix with 20 ml low-melting agarose for the preparation of an agarose block (see Subheading 3.9) for pulsed-field gel electrophoresis. 14. Store the rest of the pellet in an Eppendorf tube at −20°C for further genomic DNA analysis by PCR, Southern analysis of restricted DNA, and plasmid rescue (see Notes 21 and 22). 3.9. Preparation of Agarose Blocks Containing Chromosomes (Days 14–18)

1. Pellet the parasites by centrifugation at 1,500 rpm for 8 min after erythrocyte lysis as described in Subheading 3.8. 2. Mix part of the parasite pellet 1:1 with 1.5–2% low-melting agarose stored at 37°C in a water bath or a hot block. 3. Prepare small blocks using a plug mold or by carefully pouring the agarose suspension on a microscope slide in the area that is marked with waterproof marker-pen. 4. Let the agarose set at RT for a few minutes. 5. Cut the blocks (5 mm/5 mm/2 mm; length/width/height). 6. Place the blocks in 5 ml of SE buffer in a 15-ml tube. 7. Add 50 ml of 20 mg/ml proteinase K to the SE buffer and incubate overnight at 37°C. 8. Store the blocks in SE buffer at 4°C. Blocks can be stored for several years without loss of quality. 9. Southern analysis of pulse-field gel electrophoresis or field inversion gel electrophoresis performed with the agarose chromosome should confirm correct integration of the transfected DNA construct into the target chromosome or the presence of circular plasmid and wild-type parasites (see Note 22).

3.10. Cloning of Transfected Parasites

Start the cloning procedure after confirmation of correct integration of the constructs by Southern blot analysis of separated chromosomes or of restricted genomic DNA (see Note 23). 1. Inject a mouse i.p. with phenylhydrazine (see Subheading 3.1) on day 0, Friday. 2. Infect a mouse i.p. with 0.3 ml of a blood suspension from cryopreserved parasites on day 3 (Monday). Take one cryotube from the liquid nitrogen stocks to thaw at RT. 3. Check the parasitemia on day 4 (Tuesday) by examining Giemsa-stained blood films (see Note 24). 4. Use a mouse with 0.3–1% parasitemia calculated accurately by counting ³6,000 total cells (with no multiply infected erythrocytes, which is essential).

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5. Collect 5 ml of infected blood in a heparinized capillary tube. 6. Suspend the blood in 1 ml of complete culture medium or PBS in an Eppendorf tube. 7. Take a small sample (10 ml) and count the RBC in a hemocytometer. 8. Calculate the number of RBC per ml of blood cell suspension (usually one grid containing 16 of the smallest squares on the hemocytometer holds a 4-nl volume of cell suspension). By combining the parasitemia and number of RBC per ml, the number of infected RBC per ml can be calculated. 9. Dilute the suspension with culture medium or PBS by serial dilution in such a way that 0.5–2 parasites are present per 0.2 ml of culture medium or PBS (see Notes 24 and 25). 10. Place mice at 37°C for 10–20 min before the injection of the infected RBC. 11. Inject 0.2 ml of the suspension intravenously into the tail veins of ten mice using 0.3-ml insulin syringes with 30G needles. 12. Check the parasitemia of these mice in Giemsa-stained smears at day 8 after infection. In a successful experiment 20–50% of the mice become positive and typically have a parasitemia of 0.3–1% at day 8. Collect the blood from the positive mouse for cryopreservation and for collection of DNA for PCR analysis (see Subheading 3.8). 3.11. In Vivo Maintenance and Production of Transgenic Parasites Containing Episomes

The growth characteristics of blood stages from transgenic parasites containing episomes are different from transgenic parasites containing DNA integrated into their genome (see Notes 26 and 27). The initial infection to maintain production of transgenic parasites can be established by two methods: via passage from an existing infection, or i.p. from a cryopreserved parasite stock.

3.11.1. Infection of Mice via Mechanical Passage

1. Collect on day 0, usually Thursday, 15 ml of tail blood (three droplets from a heparinized capillary) in 0.2 ml PBS of a mouse with a parasitemia of 5–15% (collect this tail blood from a mouse that has been infected the previous week by mechanical passage, either from mouse to mouse on a Thursday or directly from liquid nitrogen storage on Friday). 2. Inject the 0.2 ml blood/PBS suspension i.p. immediately into a mouse. 3. Proceed with the drug treatment.

3.11.2. Infection of Mice from a Cryopreserved Parasite Stock

1. Infect a mouse i.p. at day 0 (usually a Friday) with 0.3 ml of a blood suspension from cryopreserved parasites (take one cryotube from the liquid nitrogen, thaw at RT). 2. Proceed with the drug treatment.

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Mice are treated 1 or 2 days after infection. Selection with pyrimethamine in vivo, provided in the drinking water: 1. Provide the animals with drinking water containing pyrimethamine 1 day after parasites have been injected and for a period of 3–4 days. 2. Parasites are collected at a parasitemia of 3–15%, not later than 2 days after the final treatment of pyrimethamine to prevent the accumulation of parasites in the population that have lost the episomes. Selection with WR99210 in DMSO injected subcutaneously: 1. Inject a single dose of 0.1 ml WR99210 solution as in Subheading 3.6.2. 2. Repeat this treatment on the following 2–3 days for a total of three to four treatments. 3. Parasites are collected at a parasitemia of 3–15%, not later than 2 days after the final treatment of pyrimethamine to prevent the accumulation of parasites in the population that have lost the episomes.

3.12. Negative Selection Using 5-Fluorocytosine In Vivo Provided in the Drinking Water

This protocol is based upon the use of outbred mice weighing 20–30 g, presuming that the animal consumes 5–10 ml of drinking water daily, to receive an equivalent dose of 5–10 mg of 5-FC per day (see Notes 28 and 29). 1. A TO or an NIH Swiss outbred mouse at 25 g is infected i.p. on day 0, typically a Friday, with 0.1 ml of a thawed suspension of cryopreserved infected erythrocytes. 2. Monitor the parasitemia daily from day 3 until the negative selection process has reached a close. 3. Collect the blood by cardiac puncture when the parasitemia is 0.5–5%, typically a Monday, day 3, enabling passage of parasitized blood into two mice in such a fashion that parasitemia is observed on day 5 (see Notes 30 and 31). 4. Drinking water with 1 mg/ml 5-FC pro-drug should be applied once a parasitemia of £0.1% is achieved; this can be from day 4 to 6 (see Note 32). The drug water should be replenished after 4 days of treatment if the experiment is still running. 5. The infection levels should drop off to zero after 48 h of 5-FC treatment. Parasite clearance is generally around days 6–9, with the parasitemia rising again at days 8–10. 6. Once the parasitemia is greater than 3%, typically a Monday (day 10), collection of the parasites should be carried out with a cardiac puncture. Prepare cryopreserved stabilates and a parasite

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pellet following Subheading 3.8 to prepare DNA and an agarose chromosome block. 7. Cloning is recommended prior to further manipulation of the line.

4. Notes 1. Phenylhydrazine hydrochloride is a suspected carcinogen. A mask must be worn when handling the powder form and gloves used at all times as it can be absorbed through the skin. Take care when disposing of this substance to flush with copious amounts of water due to its toxic effect on aquatic organisms. 2. Giemsa solution is prepared at 12% (v/v) with Sörensen staining buffer. Once the thin blood film smears have dried, they are fixed with methanol for 2 s, air-dried, and left to stain for a minimum of 10 min before rinsing with tap water. 3. Because Nycodenz powder becomes very viscose when dissolving, it is not necessary to fully dissolve the powder into the buffered medium before autoclaving as this assists the process. Autoclave at 120°C for 20 min. 4. Pyrimethamine stock needs to be vortexed to dissolve into DMSO. Working stock should be diluted from 100× stock with tap water and then pH adjusted to pH 3.5–5 with 1 M HCl before a clear solution is obtained. Pyrimethamine should then be dispensed from an opaque or darkened bottle due to it being light sensitive. 5. Where Plasmodipur filters are not available or a cheaper alternative is required, an empty 10-ml Zeba spin column can be filled with 3–4 cm of Whatman CF-11 powder. Continue to equilibrate with a wash as per the method in Subheading 3.8. 6. Drinking water containing 5-FC should be prepared using tap water and requires vigorous shaking to dissolve. Additionally, vortexing and the use of warmed water can help to force the crystals into solution. 5-FC can be dissolved up to 15 mg/ml solution. The solution is light sensitive; therefore any transparent drinking bottles should be covered to prevent exposure to light. As with any other drug in water it should be replenished after 4 days. 7. Blood is collected at 1–10% parasitemia to be mixed 1:1 with 30% glycerol/PBS (v/v) with heparin 10 U/ml. The blood/ glycerol suspension is aliquoted into 500 ml volumes to be stored in 1-ml cryotubes at −80°C initially, before being transferred to liquid nitrogen for long-term storage.

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8. Blood is collected from the positive mouse (1–2 ml) or rat (5–7 ml) (infected at day 2) at a parasitemia of 1–3% at day 5 (Monday) or 6 (Tuesday) between 10.00 a.m. and 4.00 p.m. Higher parasitemia is suboptimal since many erythrocytes will become multiply infected. In animals that are kept under the normal day/night light regime, the development of the parasites is relatively synchronous. In these animals rupturing of the schizonts and invasion by merozoites occur in the early morning between 4.00 and 6.00 a.m. Therefore, most parasites are in the ring form/young trophozoite stage when the infected blood is collected. The infected blood is then cultured overnight at 36.5–37°C. By 9.00 am the next day, all parasites have developed into mature schizonts, which do not rupture under in vitro conditions. Schizonts of P. berghei containing mature merozoites can survive for several hours and can be manipulated without rupturing and loss of viability. There is no need to remove the leucocytes from the blood when the schizonts are used for transfection. 9. Alternatively when requiring large-scale culture to accommodate over ten transfections a Wistar rat (180–220 g) is injected i.p. with 0.5 ml of phenylhydrazine 12.5 mg/ml. 10. RBC should be singly infected in order to produce healthy mature schizonts. If multiple infected cells are present a passage to another mouse should be performed to continue the experiment with optimal infection conditions. Larger scale culture of schizonts can be carried out using a rat to provide infected erythrocytes, which usually generates enough schizonts for up to 30 transfections. Follow the method in Subheading 3.2 collecting 5–7 ml of blood using a 10-ml syringe at step 1 of the protocol. 11. Continuous gassing systems are also available to incubate cultures, but are not necessary. If gassing once at the start of culturing, ensure that the flask has a non-vented, plug seal cap to prevent gas escaping. This is crucial since it has been observed that without gas, as long as the flask is sealed and not vented, the culture will still develop viable schizonts. 12. The culture temperature is critical since the parasite developmental rate is dependent on the temperature. Above 38.5°C, parasites will degenerate, and at temperatures lower than 37°C, the parasites will develop into healthy parasites but the developmental time is extended. Even at a temperature of 30°C, the parasites will reach the mature schizont stage, but the development of ring forms into schizonts will take longer than 48 h. 13. Healthy, viable schizonts are distinguished by the presence of 12–24 “free” merozoites within one RBC and one cluster of pigment (hemozoin). Smearing the cells on the microscope

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slides often damages the red cell membrane and the merozoites are visible as more or less clustered yet free parasites. A purple (red) defined compact nucleus and a dot of blue cytoplasm are characteristics of viable merozoites. About 15–25% of the parasites in these smears are single-nucleated (young) gametocytes. Degenerate schizonts often show a compact morphology in which the separate merozoites are difficult to recognize. Be careful not to mistake developing schizonts (which are still in the process of nuclear division prior to budding off of the merozoites) for degenerated schizonts (17). 14. Start the purification procedure between 9.00 am and 10.00 am. Starting later in the morning results in a higher percentage of degenerated schizonts. For the density gradients, use Nycodenz instead of Percoll. Percoll is used by many workers to separate parasite stages. However, in contrast to Percoll, Nycodenz does not affect the viability of parasites. Collect about 0.6 × 108 to 0.16 × 109 schizonts (i.e., 0.6 × 109 to 1.6 × 109 merozoites) from 1 to 2 ml of heart blood from a mouse and from a rat about 3 × 108 to 1 × 109 schizonts (i.e., 3 × 109 to 1 × 1010 merozoites) from 5 to 7 ml of heart blood. 15. When performing a density gradient, it is possible to practice using colored water or RPMI1640 to determine the speed and pressure to release the Nycodenz solution to create a sharp line. It is useful to note that when loading a 10-ml pipette the Nycodenz should be taken to 14 ml to more easily visualize when 10 ml have been dispensed into the falcon. If a sharp line is not seen, the culture can be pelleted at 1,800 rpm for 8 min and resuspended in medium to start the gradient process again. 16. A total volume of about 30–40 ml is collected from the four tubes aliquoted from 160 ml of culture when using a rat. The schizonts (and leucocytes, gametocytes, and old trophozoites if present) will collect at the interface of the two suspensions, while the uninfected cells will pellet on the bottom of the tubes. 17. Thirty transfections can be set up from the schizont pellet of one rat, resuspending in 1 ml of culture medium per transfection intended. 18. To date three selectable markers exist for the transformation of Plasmodium: the pyrimethamine-resistant form of the DHFR/ TS gene of Plasmodium and of T. gondii, and the human DHFR gene. The latter gene confers not only resistance to pyrimethamine but also to the antimalarial drug WR99210. Introduction of all three genes into pyrimethamine-sensitive P. berghei parasites gives rise to a large increase (~1,000×) in pyrimethamine resistance that allows for a relatively simple in vivo selection procedure. The TgDHFR/TS gene is preferable to the Plasmodium gene as a selectable marker because it

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reduces the likelihood of unwanted recombination with the endogenous PbDHFR/TS gene and it may confer higher levels of resistance to pyrimethamine. Pyrimethamine selection in vivo is preferred over WR99210 selection, since the latter drug generates more side effects in rodents. The WR99210/hDHFR selection system can be used in conjunction with the pyrimethamine/TgDHFR-TS selection system, allowing for multiple manipulation of the genome such as the knockout of two genes in the same parasite clone or for complementation of knockout parasites. Since hDHFR also confers resistance to pyrimethamine, the hDHFR marker can only be used as the second selectable marker when both selectable markers are required. Treatment of the animals starts 1 day after transfected parasites have been injected into the mice (to allow the parasites to complete one full developmental cycle in the absence of drug pressure) and treatment is performed on 3–4 consecutive days. Since inoculation of the transfected schizonts occurs between 10.30 and 11.00 a.m. and one cycle takes 22–24 h, start treating the animals in the afternoon. Treatment with pyrimethamine is now performed by providing the pyrimethamine in the drinking water and not by drug injection. 19. The concentration of WR99210 is dependent on the selectable markers present in the parasite line and the episomal/integrated nature of the constructs. If only one copy of the hDHFR gene is present, select with lower concentrations of 6–8 mg; if both a resistant DHFR-TS and the hDHFR genes are present, select with 12–16 mg. 20. One day after injection of the transfected parasites, parasitemia is usually between 0.05 and 3%. After the first two drug treatments parasitemia rapidly becomes undetectable, indicating that most of the parasites do not contain the constructs. In unsuccessful experiments, parasites are often detected between day 13 and 15 after the injection of transfected parasites. These parasites are usually wild-type parasites that survived the drug treatment protocol. 21. Any material that is intended for use as a chromosome agarose block must be processed as such on that day and not stored at −20 °C since freezing fractures the chromosomes, therefore damaging the block. 22. Characterize the genotype of the uncloned, transfected parasites as follows: (a) PCR to show the presence of the selectable marker (for example TgDHFR-TS, hDHFR) and correct integration. These PCR are performed to rapidly have an indication whether the transfection was successful.

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(b) Separation of the chromosomes using Pulsed-Field Gel Electrophoresis (FIGE or CHEF) followed by hybridization of the separated chromosomes using the 3¢UTR of the PbDHFR-TS gene. This DNA fragment is present in most available constructs and is also present at chromosome 7 as endogenous gene in P. berghei. This is a standard analysis performed in all transfection experiments. The hybridization pattern and intensity give information on (1) the integration event in the target chromosome, (2) the presence of episomes (that migrate in a different pattern compared to the chromosomes), and (3) the ratio between wild-type and transfected parasites in the population. (c) Southern analysis of restricted genomic DNA. This is only done if the PCR and the chromosome hybridizations give confusing results and confirmation of correct integration is needed. Usually Southern analysis is performed on genomic DNA from cloned parasites. Southern analysis is also performed as a means to determine the ratio of wildtype to recombinant parasites in primary populations of transfected parasites. (d) Plasmid rescue: plasmid rescue provides a means to confirm the integrity of introduced episomes and to recover genomic DNA flanking the insertion site (single crossover only). 23. Do not clone parasites that are transfected with a single, circular construct unless more than one construct is present in the population. The demonstration of the presence of the unaltered construct by southern analysis and plasmid rescue is normally sufficient for these parasite populations. 24. The limiting dilution cloning procedure should be started within 1–2 h after preparation of the Giemsa-stained slides. If there is a subsequent delay longer than 2 h, parasitemia might increase as a result of invasion of new parasites. A higher parasitemia in the starting material will of course affect the cloning procedure. 25. Parasites are cloned using the method of limiting dilution of blood-stage parasites, i.e., injection of a single parasite into animals. The method described here is empirically adapted to specific laboratory conditions and it is possible that other laboratories have to make small changes in order to get reliable results with this cloning procedure. In the Leiden laboratory the inoculum is 2 parasites per mouse, whereas in the Glasgow’s laboratory we observe that 0.5 parasite per mouse is optimal resulting in an infection rate of 20–50% of the mice. It is considered that a cloning experiment is successful if less than 50% of the mice become positive. 26. Under drug pressure transformed parasites that contain episomes grow slower than transgenic parasites containing DNA

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integrated into their genome. The slower growth rate is due to the unstable segregation of the episomes during schizogony, resulting in the production of merozoites in each cycle, which do not contain episomes. These latter parasites are therefore sensitive to pyrimethamine during their development into the next schizont generation. Up to 50–60% of merozoites produced from episome-containing parasites lack the episomes. The episome-positive parasites may contain up to 20–40 copies of the episome per nucleus. 27. It is important to maintain episomally transfected parasites under drug pressure during in vivo multiplication to prevent the loss of the episomes from the parasite population. Wildtype parasites show about a ten times multiplication rate per 24 h in mice, up to a parasitemia of 3–5%. At higher parasitemia the multiplication rate is slower as a result of a shortage of suitable host cells and multiple infected cells, which does not support optimal growth of the parasites. Transgenic parasites containing episomes grow much more slowly under drug pressure. It is preferable to start infections in mice with relatively high numbers of these transgenic parasites to obtain the required parasites in a relatively short period (4–8 days) reducing the period of pyrimethamine treatment of the mice to 3–4 days. 28. 5-FC has dissolving capabilities of up to 15 mg/ml solution and should be dissolved in tap water when used for negative selection provided in drinking water. Dissolving it requires vigorous shaking, although vortexing and the use of warmed water can help force the crystals into solution. The solution is light sensitive; therefore any transparent drinking bottles should be covered to darken them. As with any other drug water it should be replenished after 4 days if required. 29. Previously, negative selection was carried out over 4 days during which 10 mg of 5-FC was i.p. injected every day. The 5-FC stock was at 10 mg/ml in 0.9% NaCl solution, a concentration at the limit of the solubility of the compound. Negative selection via drinking water is less laborious, using a lower dose of 5-FC with cage mates using the same drinking water, thereby allowing for a reduced and refined animal usage. 30. If the parasitemia is between 0.2 and 1% then 50 ml of blood should be mixed with 0.2 ml of PBS to be injected ip, producing a parasitemia between 0.2 and 1.5% after 48 h, typically a Wednesday. 31. Alternatively, two mice can be infected directly with 0.1– 0.25 ml of a cryopreserved stabilate on a Monday, reducing animal numbers required. 32. If clearance of parasites is not visible after 48 h the concentration can be increased to 1.5 mg/ml.

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References 1. Bártfai R et al (2010) H2A.Z demarcates intergenic regions of the Plasmodium falciparum epigenome that are dynamically marked by H3K9ac and H3K4me3. PLoS Pathog 6:e1001223 2. Bozdech Z et al (2003) The transcriptome of the intraerythrocytic developmental cycle of Plasmodium falciparum. PLoS Biol 1:e5 3. Florens L et al (2002) A proteomic view of the Plasmodium falciparum life cycle. Nature 419:520–526 4. Gardner MJ et al (2002) Genome sequence of the human malaria parasite Plasmodium falciparum. Nature 419:498–511 5. Khan SM et al (2005) Proteome analysis of separated male and female gametocytes reveals novel sex-specific Plasmodium biology. Cell 121:675–687 6. Lasonder E et al (2002) Analysis of the Plasmodium falciparum proteome by highaccuracy mass spectrometry. Nature 419:537–542 7. Le Roch KG et al (2003) Discovery of gene function by expression profiling of the malaria parasite life cycle. Science 301:1503–1508 8. Otto TD et al (2010) New insights into the blood-stage transcriptome of Plasmodium falciparum using RNA-Seq. Mol Microbiol 76:12–24 9. Salcedo-Amaya AM et al (2009) Dynamic histone H3 epigenome marking during the intraerythrocytic cycle of Plasmodium falciparum. Proc Natl Acad Sci USA 106:9655–9660 10. van Dijk MR et al (1995) Stable transfection of malaria parasite blood stages. Science 268:1358–1362 11. Mota MM et al (2001) Gene targeting in the rodent malaria parasite Plasmodium yoelii. Mol Biochem Parasitol 113:271–278 12. Reece SE, Thompson J (2008) Transformation of the rodent malaria parasite Plasmodium chabaudi and generation of a stable fluorescent line PcGFPCON. Malar J 7:183 13. Kooij TW et al (2005) A Plasmodium wholegenome synteny map: indels and synteny breakpoints as foci for species-specific genes. PLoS Pathog 1:e44 14. Franke-Fayard B et al (2004) A Plasmodium berghei reference line that constitutively expresses GFP at a high level throughout the complete life cycle. Mol Biochem Parasitol 137:23–33 15. Mair GR et al (2010) Universal features of post-transcriptional gene regulation are critical for Plasmodium zygote development. PLoS Pathog 6:e1000767

16. Ponzi M et al (2009) Egress of Plasmodium berghei gametes from their host erythrocyte is mediated by the MDV-1/PEG3 protein. Cell Microbiol 11:1272–1288 17. Janse CJ et al (2006) High-efficiency transfection and drug selection of genetically transformed blood stages of the rodent malaria parasite Plasmodium berghei. Nat Protoc 1:346–356 18. Janse CJ et al (2006) High efficiency transfection of Plasmodium berghei facilitates novel selection procedures. Mol Biochem Parasitol 145:60–70 19. van Dijk MR et al (1994) Mechanisms of pyrimethamine resistance in two different strains of Plasmodium berghei. Mol Biochem Parasitol 68:167–171 20. Fidock DA, Wellems TE (1997) Transformation with human dihydrofolate reductase renders malaria parasites insensitive to WR99210 but does not affect the intrinsic activity of proguanil. Proc Natl Acad Sci USA 94:10931–10936 21. Braks JA et al (2006) Development and application of a positive-negative selectable marker system for use in reverse genetics in Plasmodium. Nucleic Acids Res 34:e39 22. Maier AG et al (2006) Negative selection using yeast cytosine deaminase/uracil phosphoribosyl transferase in Plasmodium falciparum for targeted gene deletion by double crossover recombination. Mol Biochem Parasitol 150:118–121 23. van Schaijk BC et al (2010) Removal of heterologous sequences from Plasmodium falciparum mutants using FLPe-recombinase. PLoS One 5:e15121 24. Laurentino EC et al (2011) Experimentally controlled downregulation of the histone chaperone FACT in Plasmodium berghei reveals that it is critical to male gamete fertility. Cell Microbiol 13:1956–1974 25. O’Donnell RA et al (2001) An alteration in concatameric structure is associated with efficient segregation of plasmids in transfected Plasmodium falciparum parasites. Nucleic Acids Res 29:716–724 26. van Dijk MR et al (1997) Replication, expression and segregation of plasmid-borne DNA in genetically transformed malaria parasites. Mol Biochem Parasitol 86:155–162 27. Murray AW, Szostak JW (1983) Construction of artificial chromosomes in yeast. Nature 305:189–193 28. Stinchcomb DT et al (1979) Isolation and characterisation of a yeast chromosomal replicator. Nature 282:39–43

7 29. Iwanaga S et al (2010) Functional identification of the Plasmodium centromere and generation of a Plasmodium artificial chromosome. Cell Host Microbe 7:245–255 30. Chookajorn T et al (2007) Epigenetic memory at malaria virulence genes. Proc Natl Acad Sci USA 104:899–902 31. Freitas-Junior LH et al (2005) Telomeric heterochromatin propagation and histone acetylation control mutually exclusive expression of antigenic variation genes in malaria parasites. Cell 121:25–36 32. Hernandez-Rivas R et al (2010) Telomeric heterochromatin in Plasmodium falciparum. J Biomed Biotechnol. doi:10.1155/2010/ 290501 33. Lopez-Rubio JJ et al (2007) 5¢ flanking region of var genes nucleate histone modification patterns linked to phenotypic inheritance of virulence traits in malaria parasites. Mol Microbiol 66:1296–1305 34. Petter M et al (2011) Expression of P. falciparum var genes involves exchange of the histone variant H2A.Z at the promoter. PLoS Pathog 7:e1001292 35. Meissner M et al (2005) Tetracycline analogueregulated transgene expression in Plasmodium falciparum blood stages using Toxoplasma gondii transactivators. Proc Natl Acad Sci USA 102:2980–2985 36. Armstrong CM, Goldberg DE (2007) An FKBP destabilization domain modulates protein levels in Plasmodium falciparum. Nat Methods 4:1007–1009 37. Muralidharan V et al (2011) Asparagine repeat function in a Plasmodium falciparum protein assessed via a regulatable fluorescent affinity tag. Proc Natl Acad Sci USA 108:4411–4416 38. Carvalho TG et al (2004) Conditional mutagenesis using site-specific recombination in

Transfection of Rodent Malaria Parasites

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

125

Plasmodium berghei. Proc Natl Acad Sci USA 101:14931–14936 Combe A et al (2009) Clonal conditional mutagenesis in malaria parasites. Cell Host Microbe 5:386–396 de Koning-Ward TF et al (2009) A newly discovered protein export machine in malaria parasites. Nature 459:945–949 Ivics Z et al (2009) Transposon-mediated genome manipulation in vertebrates. Nat Methods 6:415–422 Mátés L et al (2009) Molecular evolution of a novel hyperactive Sleeping Beauty transposase enables robust stable gene transfer in vertebrates. Nat Genet 41:753–761 Patton EE, Zon LI (2001) The art and design of genetic screens: zebrafish. Nat Rev Genet 2:956–966 Schneider A, Leister D (2006) Forward genetic screening of insertional mutants. Methods Mol Biol 323:147–161 Yergeau DA, Mead PE (2007) Manipulating the Xenopus genome with transposable elements. Genome Biol 8(Suppl 1):S11 Sakamoto H et al (2005) Towards systematic identification of Plasmodium essential genes by transposon shuttle mutagenesis. Nucleic Acids Res 33:e174 Balu B et al (2005) High-efficiency transformation of Plasmodium falciparum by the lepidopteran transposable element piggyBac. Proc Natl Acad Sci USA 102:16391–16396 Fonager J et al (2011) Development of the piggyBac transposable system for Plasmodium berghei and its application for random mutagenesis in malaria parasites. BMC Genomics 12:155 Balu B, Adams JH (2006) Functional genomics of Plasmodium falciparum through transposon-mediated mutagenesis. Cell Microbiol 8:1529–1536