We thank Drs Randy Elkins, Robin Kastenmayer, Ted Torrey, Dan Pare and Tovi Lehman for critical reading of manuscripts. This work was supported by the Intramural Research Program of the Division of Intramural Research, National Institute of Allergy and Infectious Diseases, National Institutes of Health, and by the 973 National Basic Research Program of China, #2007CB513103. We thank NIAID intramural editor Brenda Rae Marshall for assistance.

Aseptic techniques must be applied to all materials that will be administrated into animals to avoid inadvertent introduction of exogenous infectious agents into mice that can confound experimental outcomes.

1. 	 Infection of Laboratory Mice with Blood-stage Malaria Parasites
<ol>
<li>At room temperature, thaw two vials containing the frozen blood stage malaria parasites to be crossed. In this example, two rodent malaria parasite strains used are Plasmodium yoelii yoelii and Plasmodium yoelii nigeriensis.</li>
<li>Fit a syringe with a 22-30 gauge (G) needle and draw up 400 &mu;L of sterile, pharmaceutical grade PBS (pH 7.4).</li>
<li>Using the same syringe, draw up 100 &mu;L of the thawed infected blood; invert the syringe up and down until the content is homogenously mixed. (Optional: a pipette may be used to transfer PBS and infected blood into a collection tube prior to transferring into a syringe for injection).</li>
<li>Inject 200 &mu;L of parasite containing solution directly into the peritoneum of a mouse. When finished, discard the syringe in a sharps bin. (See also Supplementary Materials for maintenance of laboratory mice). In general, the standard size of the inoculum is between 100-500 &mu;L, depending on the levels of parasite infections in the original frozen materials.</li>
<li>Inject the two parasite strains to be crossed into two mice each. Mice will become positive microscopically 3 to 5 days after the injections. When parasitemias reach between 1-5%, continue to the next step.</li>
</ol>
2. 	Mixed Clone Infection of Mice
<ol>
<li>Prior to mixed clone infection, collect tail blood from the two donor mice infected with the different parasite strains for microscopic examination by Giemsa stained thin blood smears (Fig 3), and for measurement of red blood cell density (see Supplementary Materials for detailed procedures).</li>
<li>Calculate the volume of a physiological saline (1.5% w/v trisodium citrate dihydrate, 0.85% w/v sodium chloride) and mouse blood required to produce an inoculum mixture, using the parasitemia and red blood cell density measurements obtained earlier.</li>
<li>Adjust the parasite solution to a final concentration of one million infected red blood cells per 100&mu;l.</li>
<li>Combine the infected blood from the two donors in a 1:1 ratio to obtain a mixed-clone inoculum. When the two parental clones differ in growth rate, adjust the proportions of a faster-growing parental to a slower-growing parental clone to 1:2 or 1:5.</li>
<li>Inject 100 &mu;L of the mixed sample into the peritoneum of each mouse to be infected. (see also step 3.9 to determine the number of mice to be injected)</li>
<li>Using this method, infect two mice each with the single parental strains. The single-clone infections will allow determination of successful transmission of the two parental strains.</li>
</ol>
3. 	Feeding Mosquitoes
<ol>
<li>Prepare three cages containing 300-500 adult (male and females) Anopheles stephensi mosquitoes each, age 5-7 days post emergence from pupae. The containers are covered with netting secure by a plastic or metal lid. Two cages are for feeding on parental clones; the third cage is for preparation of a genetic cross.</li>
<li>Starting four days after the mixed-clone infection, prepare Giemsa-stained thin blood tail smears from the infected mice.</li>
<li>Examine the smears under a microscope by scanning at least 50 microscopic fields at 1000x magnification and determine whether male and female gametocytes are present.</li>
<li>Male and female gametocytes are recognizable due to the presence of large vacuole and large granules (Fig 3). Male and female gametocytes have pink and blue cytoplasm, respectively. If gametocytes are present, follow Step 3.6.</li>
<li>If gametocytes are absent, keep monitoring daily until they appear. If gametocytes are present, continue the following step.</li>
<li>Inject 50 &mu;L of anesthetic drug cocktail containing 33 mg/mL of ketamine and 3 mg/mL of xylazine into the peritoneum of a mouse to anesthetize each infected mouse (per 20 gram mouse body weight). Final dose of ketamine and xylazine is 82.5 and 7.5 mg/Kg, respectively.</li>
<li>Place the mice face down on the top of a cage containing adult Anopheles stephensi mosquitoes that have been starved of sugar supply for 24 hrs and allow the mosquitoes to feed for 30 minutes without interruption. </li>
<li>Maintain the mosquitoes in an insectary under standard conditions (temperature is maintained between 24-26&deg;C; also see Supplementary Methods for maintenance of laboratory mosquitoes).</li>
<li>Use one infected mouse for every 50-100 female mosquitoes. Euthanize the mice immediately by cervical dislocation after the feeding while the mice are still under anesthesia.</li>
</ol>
4. 	Dissecting Mosquito Midguts and Counting Oocysts
<ol>
<li>Nine days after the feeding, the mosquitoes will be examined for oocysts in the insect's midgut. The presence of oocysts in the mosquitoes that feed on single-clone infected mice indicates that gametocytes of the two parental strains are infectious.</li>
<li>To dissect a mosquito midgut, use an aspirator connected to a small container to collect 5 to 10 infected mosquitoes from the cage.</li>
<li>Using CO2 gas, anesthetize the mosquitoes and transfer them into a glass Petri dish on ice. Separate and discard male mosquitoes. The antennae of males are noticeably bushier in comparison to the females.</li>
<li>Fill two syringes with PBS and fit them with 26 gauge half-inch (G&frac12;) needles. Deposit about 100 &mu;L of PBS from a syringe onto a glass slide.</li>
<li>Transfer 5 to 10 anesthetized female mosquitoes onto the drop of PBS and place the slide on a dissecting microscope.</li>
<li>Using the same needles, remove each mosquito's wings and legs, then, holding the insect at the thorax and posterior abdomen, pull the body apart until the midgut, a white sac-like body, is released. Detach the midgut and discard the rest of the body.</li>
<li>Add more PBS on the midgut if the slides are dry. The midgut will degenerate if let dried. Place a cover slip over the midguts.</li>
<li>Examine the midguts under a light microscope at a 100x magnification. Count the numbers of oocysts in each midgut (Fig 3). Oocysts are recognizable as thick-wall circular structures that lie on the outer surface of the midguts. </li>
<li>When finished, dispose of syringes, needles, and slide in a sharps bin.</li>
<li>If oocysts are absent in the midguts, feed mosquitoes with infected mice with the higher number of gametocytes and give a longer feeding time. Temperature is critical for this step. Make sure the temperature is set at 24 to 26&deg; C.</li>
</ol>
5. 	Collecting Sporozoites from Mosquitoes by Centrifugation
<ol>
<li>Prepare sporozoite collection tubes as follows: cut the lid of a clean 0.5-mL centrifuge tube and punch a hole (0.1-0.2 mm diameter) at the bottom of the tube using a flamed 19 G needle.</li>
<li>Fill 1/3 of the tube with glass wool. Insert the filter tube into a 1.5-mL collection tube (one tube is sufficient to harvest sporozoites from ~100 mosquitoes)</li>
<li>On day 17 post feeding, collect mosquitoes (Step 4) from the cages into a small container using an aspirator.</li>
<li>Use CO2 gas to anesthetize mosquitoes.</li>
<li>Separate the females from the male mosquitoes. Transfer the anesthetized female mosquitoes from the container to a glass Petri dish on ice.</li>
<li>Prepare two syringes with 26 G&frac12; needles filled with PBS. Deposit about 100 &mu;L of PBS from a syringe onto a glass slide.</li>
<li>Transfer the mosquitoes onto a microscope slide. Remove the mosquitoes' head, wings, legs, and abdomen under a dissecting microscope. </li>
<li>Using fine-tipped forceps transfer the thoraxes into 0.5 mL PBS in a 1.5-mL homogenizer (mortar and pestle). Store the thoraxes on ice (sporozoite remains infective for 2-3 h). </li>
<li>Homogenize the thoraxes on ice to release sporozoites from salivary glands (see Figure 3), and transfer the lysates (homogenates) into the tube filled with glass wool (Step 5.1-5.2) using a glass pipette.</li>
<li>Centrifuge for 10 to 20 sec at 1000 rpm at room temperature.</li>
<li>Collect the flow-through (the purified sporozoite suspension) using a syringe with a 22-30 G needle. (Optional: drop 10-50 &mu;L of the flow-through on a microscopic slide, place a cover slip and visualize the live sporozoites under a standard light microscope, 400X magnification, see also Fig 3)</li>
<li>Inject i.p. 0.1-0.5 mL of the sporozoite suspension into mice (up to 200,000 parasites can be harvested from a single mosquito; see ref 12). In a successful case, the animal will become infected 72 hrs after an injection.</li>
</ol>
6. 	Cloning Progeny using Limited Dilution
<ol>
<li>Seventy-two hrs after the sporozoite injections, collect tail blood from the two donor mice infected with the different parasite strains for microscopic examination after Giemsa staining of thin blood smears and for measurement of red blood cell density (see Supplementary Materials for detailed procedures).</li>
<li>Pick a mouse displaying a 0.1 to 1% parasitemia and whose infected red blood cells contain mostly single parasites as a donor for the cloning procedure. Repeat step 6.1 next days if the mice are microscopically negative. If the animals are not infected within 14 days, repeat Step 2 to 5 and determine the presence of sporozoites in step 5.11).</li>
<li>Dilute the harvested tail blood in a cold 1:1 solution of calf serum and mammalian Ringer's solution (2.7 mM potassium chloride, 1.8 mM calcium chloride, 154 mM sodium chloride) so as to attain a concentration of 0.6 to 1 infected red blood cell per 100 &mu;L, and store the inoculum on ice. In this manner, most of individual mouse should receive either one or zero parasites.</li>
<li>Intravenously inject 100 &mu;L of diluted infected blood in each of 50 to 100 mice.</li>
<li>Five to seven days later, screen the mice for the presence of blood stage parasites by examining Giemsa-stained thin blood smears. In a successful experiment, 20-50% of the mice become infected and will show parasitemia of 0.1-5.0% at day 8 post infection.</li>
</ol>
7. 	Characterization of Progeny using Genetic Markers
<ol>
<li>Collect 20 to 40 &mu;L of mouse tail blood in a 1.5-mL microcentrifuge tube that contains 0.2 mL of chilled physiological citrate saline solution, vortex briefly, and centrifuge for 3 min at 3000 rpm at room temperature to pellet the blood cells. Extract DNA immediately or keep the pellet at -80&deg;C.</li>
<li>Prepare parasite DNA using any standard DNA extraction methods. For rapid isolation of DNA, use a High Pure PCR template preparation kit (Roche Applied Bioscience).</li>
<li>Recombinant progeny are identified after genotyping using microsatellite (MS) or single nucleotide polymorphism (SNP) on different chromosomes that can differentiate the two parental strains of the genetic cross. A simplified method using single fluorescent-labeled primers for MS genotyping of P. yoelii can be used (see ref 8,9).</li>
</ol>
8. 	 Cryopreservation of Blood-stage Malaria Parasites
<ol>
<li>Collect 0.5-1 mL of infected blood by cardiac puncture from an anesthetized mouse with parasitemia of 5-20% in 5-10 mL chilled physiological saline solution.</li>
<li>Centrifuge for 5 min at 2000 rpm at room temperature. Discard supernatant.</li>
<li>Add drop-wise 2x volume of Glycerolyte 57 solution (Fenwal Laboratories) onto blood pellets and mix well.</li>
<li>Transfer the suspension to cryotubes (0.2-0.5 mL per tube).</li>
<li>Store the cryopreserved blood at -80 &deg;C (up to 1 month) or in liquid nitrogen.</li>
</ol>
9. Representative Results:

In a successful experiment, 5-10% of the cloned lines will be independent recombinant progeny.

 <img src="/files/ftp_upload/2365/2365fig1.jpg" alt="Figure 1" /> 
Figure 1. A schematic representation of the life cycle and genetic recombination events during a genetic cross. Genetic recombination events take place during the early phase of development in the mosquito. Following the fertilization of "haploid" male and female gametes of different genetic backgrounds in the midgut of the mosquito, "diploid" zygotes are formed. The zygote constitutes the only diploid stage of the parasite life cycle; meiosis follows within 12 hrs of fertilization, and all subsequent stages in the mosquito and mammals remain haploid. The resulting zygotes develop into ookinetes and oocysts. Growth and mitotic division of each oocyst produces thousands of sporozoites, which are released into mosquitoes' hemoceal. Those sporozoites that migrate to salivary glands are in position to be transmitted to a mammalian host, where development of the liver and red blood cell stages takes place. During the course of the red blood cell cycles, some parasites can differentiate into mature male and female gametocytes. The parasite life cycle is completed when these gametocytes are taken up by another mosquito, perpetuating transmission.
 <img src="/files/ftp_upload/2365/2365fig2.jpg" alt="Figure 2" /> 
Figure 2. Inheritance of nuclear genes in malaria parasites and production of recombinant progeny. Genetic recombination can be generated through chromosomal reassortment or by crossover events. Homozygous progeny produced by selfing between gametes of the same parental clone 1 or 2 (A and D), respectively. B) Heterozygous progeny produced by cross-fertilization between gametes of the two different parental clones, recombinant nuclei (blue ovals) being formed by chromosomal reassortment alone. C) Heterozygous progeny produced from cross-fertilization between gametes of the two different parent clones, recombinant nuclei (red ovals) being formed by crossovers between the non-sister chromatids of the homologous chromosomes.
 <img src="/files/ftp_upload/2365/2365fig3.jpg" alt="Figure 3" /> 
Figure 3. Morphology of the rodent malaria parasite Plasmodium yoelii. A) ring stage, B) trophozoite, C) schizont, D) merozoites, E) immature male gametocyte, F) mature male gametocyte, G) immature female gametocyte, H) mature female gametocyte, I) midgut with oocysts on day 10 post feeding (mature oocysts indicated by arrows; 200x magnification), J) mosquito salivary glands (100x magnification), K) sporozoites (400x magnification).

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We demonstrate the techniques for the production of a genetic cross in the rodent malaria Plasmodium yoelii, which is also applicable to production of genetic crosses in other rodent malarias. Infections of mice with single parental clones are usually performed to determine successful transmission of the parental parasites to ensure that the parents are competent in producing functional gametes before performing a cross.
Successful transmission through a mosquito is influenced by mul...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Glyerolyte 57 solution</td>
<td>&nbsp;</td>
<td>Cenmed</td>
<td>4A7833</td>
<td>&nbsp;</td>
<tr>
<td>Mouse Mus musculus</td>
<td>&nbsp;</td>
<td>Charles River Laboratory</td>
<td>&nbsp;</td>
<td>Female, inbred, strain Balb/C</td>
</tr>
<tr>
<td>Heat-inactivated calf serum</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>26010-066</td>
<td>&nbsp;</td>
<tr>
<td>Phosphate buffered saline (PBS) solution</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>10010-072</td>
<td>pH 7.4; Cell Culture grade </td>
</tr>
<tr>
<td>Malaria parasite Plasmodium yoelii yoelii 17XNL(1.1)</td>
<td>&nbsp;</td>
<td>MR4</td>
<td>MRA-593</td>
<td>deposited by DJ Carucci</td>
</tr>
<tr>
<td>Malaria parasite Plasmodium yoelii nigeriensis N67</td>
<td>&nbsp;</td>
<td>MR4</td>
<td>MRA-427</td>
<td>deposited by W Peters, BL Robinson, R Killick Kendrick</td>
</tr>
<tr>
<td>Mosquito Anopheles stephensi</td>
<td>&nbsp;</td>
<td>MR4</td>
<td>MRA-128</td>
<td>deposited by MQ Benedict</td>
</tr>
<tr>
<td>Cellometer automatic cell counter</td>
<td>&nbsp;</td>
<td>Nexcelom Biosciences</td>
<td>Cellometer Auto T4</td>
<td>&nbsp;</td>
<tr>
<td>Cellometer CP2 disposable hemacytometer</td>
<td>&nbsp;</td>
<td>Nexcelom Biosciences</td>
<td>Cellometer CP2 </td>
<td>&nbsp;</td>
<tr>
<td>High Pure PCR template preparation kit</td>
<td>&nbsp;</td>
<td>Roche Applied Science</td>
<td>11 796 828 001</td>
<td>&nbsp;</td>
<tr>
<td>Calcium chloride</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>C5670</td>
<td>Cell culture tested; insect cell culture tested</td>
</tr>
<tr>
<td>Giemsa stain, modified</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>GS500 </td>
<td>&nbsp;</td>
<tr>
<td>Ketamine hydrochloride</td>
<td>&nbsp;</td>
<td>Fort Dodge Animal Health</td>
<td> NDC 0856-2013-01</td>
<td> Pharmaceutical grade; concentration to 100 mg/mL</td>
</tr>
<tr>
<td>Potassium chloride</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>P5405</td>
<td>Cell culture tested; insect cell culture tested</td>
</tr>
<tr>
<td>Sodium chloride</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>S5886</td>
<td>Cell culture tested; insect cell culture tested</td>
</tr>
<tr>
<td>Trisodium citrate dihydrate</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>S4641</td>
<td>&nbsp;</td>
<tr>
<td>Xylazine</td>
<td>&nbsp;</td>
<td>Akorn Inc.</td>
<td> 4811-20ml</td>
<td>Pharmaceutical grade; concentration to 20 mg/mL</td>
</tr>
<tr>
<td>Glass wool</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>32848-003</td>
<td>&nbsp;</td>
<tr>
<td>Glass capillary (1 &mu;L)</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>53440-001</td>
<td>&nbsp;</td>
<tr>
<td>Hemocytometer</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>15170-168</td>
<td>Complete chamber set</td>
</tr>
<tr>
<td>Homogenizer</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>KT749520-0090</td>
<td>Pestle with matching tube, 1.5 mL </td>
</tr>		</tbody>
		</table>
		</div>
			<div>
			SUPPLEMENTARY MATERIALS:
<ul>
<li>Maintenance of laboratory mice</li>
<li>Maintenance of laboratory mosquitoes</li>
<li>Microscopic examination of thin blood smears stained with Giemsa stain</li>
<li>Measurement of red blood cell density</li>
</ul>

Maintenance of laboratory mice
Females of inbred laboratory mouse strain BALB/c, aged 5 to 8 weeks old, are used in the study. Mice are housed in a standard solid-bottom polycarbonate cage with wire-bar lid, equipped with feeder and a water bottle. Mice are maintained at a constant temperature (25 &plusmn; 1&deg;C) on 12:12 hour light:dark cycle. Mice are allowed to feed on 2018S Harlan Teklad Global 19% protein extruded rodent diet (sterilizable; from Harlan-Teklad) and supplied with acidified drinking water ad libitum. Experiments on animals are performed in accordance with the guidelines and regulations set forth by the Animal Care and Use Committee at the National Institute of Allergy and Infectious Disease under protocol LMVR11E (National Institutes of Health, Bethesda, Maryland).
Maintenance of laboratory mosquitoes
Mosquitoes are from a laboratory-bred colony of Anopheles stephensi. The adults are maintained in nylon cages kept in a temperature- and humidity-controlled room (23 to 25&deg;C for Plasmodium yoelii and Plasmodium chabaudi, and 19 to 21&deg;C for Plasmodium berghei; 80 to 95% humidity; on 12:12 hours light:dark cycle). Adult mosquitoes are fed with 10% glucose and 2.00% para-aminobenzoic acid (PABA) supplemented water solution. To obtain high-quality adults, 500 larvae are grown in a low-density condition in 1 L of distilled water in a 1,000-cm3 open dish supplied with approximately 1 mg of sodium bicarbonate. After hatching, the larvae are given tetramin powder (PETCO) until they develop into the pupa stage and are transferred to the adult mosquito cages for emerging.
Microscopic examination of thin blood smears stained with Giemsa stain
Using clean scissors snip off the tip (1.0 mm) of the infected mouse's tail. Place one drop (0.5-1.0 &mu;L) of tail blood onto a clean specimen slide. Mouse will stop bleeding in 1-2 min. Place a clean spreader slide on top of the blood drop, maintaining it at a 45&deg; angle relative to the specimen slide, and allow the blood to adsorb to the entire width of the spreader. Hold the specimen slide and push forward the spreader slide rapidly and smoothly to produce a thin smear. Let the blood film dry, and then immerse the slides in absolute methanol. Allow the slide to air dry once more before covering it with Giemsa stain (10% Giemsa dye in distilled water). After incubating the thin blood films for 10-15 min at room temperature, carefully rinse the slides with tap water and let it air dry. Examine the number of infected red blood cells (iRBC; see Figure 3 for morphology of infected RBC) under a light microscope with immersion oil at 1000x magnification (with 100x objective lens) and calculate parasitemia (the number of iRBC per 100 RBC counted). Different strains of malaria parasites vary in growth rate and pathogenicity. Monitoring of blood stage parasitaemias can be performed 24hrs after injections, depending on the dose of the blood stage malaria parasites. For example, mice will be microscopically positive 24 hrs when injected with 107 infected RBC intraperitoneally or 106 infected RBC intravenously.
Measurement of red blood cell density
Like the levels of parasitaemias, red blood cell (RBC) density in infected mice varies throughout the course of infection. RBC density should be measured within 1-2 hrs before the start of the single- and mixed-clone infection and the cloning experiments. There are two methods for measurement of RBC density: a manual counting using Neubauer hemocytometer and an automatic counting using a Cellometer (Nexcelom Bioscience). In both methods, withdraw 1 &mu;L of mouse tail blood using a glass capillary (VWR) and dilute in 10 mL of PBS and mix well. To use a Neubauer hemocytometer, load 20 &mu;L of the suspension onto the hemacytometer. Place the hemacytometer on a light microscope with 10x objective lens. The hemacytometer contains a grid divided into 9 large squares, and 4 large squares at the corner are further divided into 16 small squares. Count the total number of cells in each of the 16 small squares in the four corner squares. To avoid counting bias or counting cells that overlap a grid line, count a cell as "in" if it overlaps the top or right lines and "out" if it overlaps the bottom or left lines. Estimate the number of cells per one small square and divide by 0.00625 (the volume of one small square is 6.25 nL). This yields the number of cells per microliter (&mu;L). From this data, calculate the final red blood cell density by multiplying with 10,000 (a dilution factor). Rinse the cover slip and counting chamber with distilled water and 70% ethanol; air dry. Alternatively, load 20 &mu;L of the suspension onto a Cellometer counting chamber slide. Insert the slide into a Cellometer slide chamber (the reader). Start the Cellometer software, select the "red blood cell" option, and enter a dilution factor of 10,000. Record the RBC density.		</div>

protocol for production of a genetic cross of the rodent malaria parasites

Variation in response to antimalarial drugs and in pathogenicity of malaria parasites is of biologic and medical importance. Linkage mapping has led to successful identification of genes or loci underlying various traits in malaria parasites of rodents1-3 and humans4-6. The malaria parasite Plasmodium yoelii is one of many malaria species isolated from wild African rodents and has been adapted to grow in laboratories. This species reproduces many of the biologic characteristics of the human malaria parasites; genetic markers such as microsatellite and amplified fragment length polymorphism (AFLP) markers have also been developed for the parasite7-9. Thus, genetic studies in rodent malaria parasites can be performed to complement research on Plasmodium falciparum. Here, we demonstrate the techniques for producing a genetic cross in P. yoelii that were first pioneered by Drs. David Walliker, Richard Carter, and colleagues at the University of Edinburgh10.

Genetic crosses in P. yoelii and other rodent malaria parasites are conducted by infecting mice Mus musculus with an inoculum containing gametocytes of two genetically distinct clones that differ in phenotypes of interest and by allowing mosquitoes to feed on the infected mice 4 days after infection. The presence of male and female gametocytes in the mouse blood is microscopically confirmed before feeding. Within 48 hrs after feeding, in the midgut of the mosquito, the haploid gametocytes differentiate into male and female gametes, fertilize, and form a diploid zygote (Fig. 1). During development of a zygote into an ookinete, meiosis appears to occur11. If the zygote is derived through cross-fertilization between gametes of the two genetically distinct parasites, genetic exchanges (chromosomal reassortment and cross-overs between the non-sister chromatids of a pair of homologous chromosomes; Fig. 2) may occur, resulting in recombination of genetic material at homologous loci. Each zygote undergoes two successive nuclear divisions, leading to four haploid nuclei. An ookinete further develops into an oocyst. Once the oocyst matures, thousands of sporozoites (the progeny of the cross) are formed and released into mosquito hemoceal. Sporozoites are harvested from the salivary glands and injected into a new murine host, where pre-erythrocytic and erythrocytic stage development takes place. Erythrocytic forms are cloned and classified with regard to the characters distinguishing the parental lines prior to genetic linkage mapping. Control infections of individual parental clones are performed in the same way as the production of a genetic cross.

Watch this Scientific Journal Video about Protocol for Production of a Genetic Cross of the Rodent Malaria Parasites at JoVE.com

Protocol for Production of a Genetic Cross of the Rodent Malaria Parasites

Genetic crosses of rodent malaria parasites are performed by feeding two genetically distinct parasites to mosquitoes. Recombinant progeny are cloned from mouse blood after allowing mosquitoes to bite infected mice. This video shows how to produce genetic crosses of Plasmodium yoelii and is applicable to other rodent malaria parasites.

Variation in response to antimalarial drugs and in pathogenicity of malaria parasites is of biologic and medical importance. Linkage mapping has led to ...

protocol-for-production-genetic-cross-rodent-malaria

Research

JoVE Journal

Immunology and Infection

15.1K Views.  National Institutes of Health. Genetic crosses of rodent malaria parasites are performed by feeding two genetically distinct parasites to mosquitoes. Recombinant progeny are cloned from mouse blood after allowing mosquitoes to bite infected mice. This video shows how to produce genetic crosses of Plasmodium yoelii and is applicable to other rodent malaria parasites.

Video: Protocol for Production of a Genetic Cross of the Rodent Malaria Parasites

In vivo Imaging of Transgenic Leishmania Parasites in a Live Host

Distinct species of Leishmania, a protozoan parasite of the family Trypanosomatidae, typically cause different human disease manifestations. The most common forms of disease are visceral leishmaniasis (VL) and cutaneous leishmaniasis (CL). Mouse models of leishmaniasis are widely used, but quantification of parasite burdens during murine disease requires mice to be euthanized at various times after infection. Parasite loads are then measured either by microscopy, limiting dilution assay, or qPCR amplification of parasite DNA. The in vivo imaging system (IVIS) has an integrated software package that allows the detection of a bioluminescent signal associated with cells in living organisms. Both to minimize animal usage and to follow infection longitudinally in individuals, in vivo models for imaging Leishmania spp. causing VL or CL were established. Parasites were engineered to express luciferase, and these were introduced into mice either intradermally or intravenously. Quantitative measurements of the luciferase driving bioluminescence of the transgenic Leishmania parasites within the mouse were made using IVIS. Individual mice can be imaged multiple times during longitudinal studies, allowing us to assess the inter-animal variation in the initial experimental parasite inocula, and to assess the multiplication of parasites in mouse tissues. Parasites are detected with high sensitivity in cutaneous locations. Although it is very likely that the signal (photons/second/parasite) is lower in deeper visceral organs than the skin, but quantitative comparisons of signals in superficial versus deep sites have not been done. It is possible that parasite numbers between body sites cannot be directly compared, although parasite loads in the same tissues can be compared between mice. Examples of one visceralizing species (L. infantum chagasi) and one species causing cutaneous leishmaniasis (L. mexicana) are shown. The IVIS procedure can be used for monitoring and analyzing small animal models of a wide variety of Leishmania species causing the different forms of human leishmaniasis.

In vivo Imaging of Transgenic Leishmania Parasites in a Live Host

An in vivo imaging system is used to generate quantitative measurements of murine infection with the Trypanosomatid protozoan Leishmania. This is a non-invasive and non-lethal method for detecting parasites expressing luciferase within many tissues throughout the course of chronic Leishmania spp. infection.

Distinct species of Leishmania, a protozoan parasite of the family Trypanosomatidae, typically cause different human disease manifestations. The most ...

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

Mosquitoes are vectors for a number of disease-causing pathogens such as the yellow fever virus, malaria parasites and filarial worms. Laboratories are investigating anti-pathogen components of the innate immune system in disease vector species in the hopes of generating transgenic mosquitoes that are refractory to such pathogens1, 2. The innate immune system of mosquitoes consists of several lines of defense 3. Pathogens that manage to escape the barrier imposed by the epithelium-lined mosquito midgut 4 enter the hemolymph and encounter circulating hemocytes, important cellular components that encapsulate and engulf pathogens 5, 6. Researchers have not found evidence for hematopoietic tissues in mosquitoes and current evidence suggests that the number of hemocytes is fixed at adult emergence and numbers may actually decline as the mosquito ages 7. The ability to properly collect and identify hemocytes from medically important insects is an essential step for studies in cellular immunity. However, the small size of mosquitoes and the limited volume of hemolymph pose a challenge to collecting immune cells. 
Two established methods for collecting mosquito hemocytes include expulsion of hemolymph from a cut proboscis 8, and volume displacement (perfusion), in which saline is injected into the membranous necklike region between the head and thorax (i.e., cervix) and the perfused hemolymph is collected from a torn opening in a distal region of the abdomen 9, 10. These techniques, however, are limited by low recovery of hemocytes and possible contamination by fat body cells, respectively 11. More recently a method referred to as high injection/recovery improved recovery of immunocytes by use of anticoagulant buffers while reducing levels of contaminating scales and internal tissues 11. While that method allows for an improved method of collecting and maintaining hemocytes for primary culture, it entails a number of injection and collecting steps that are not necessary if the downstream goal is to collect, fix and stain hemocytes for diagnostics. Here, we demonstrate our method of collecting mosquito hemolymph that combines the simplicity of perfusion, using anticoagulant buffers in place of saline solution, with the accuracy of high injection techniques to isolate clean preparations of hemocytes in Aedes mosquitoes.

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

A simplified yet accurate method to collect and stain mosquito hemocytes is described. Our method combines the simplicity of perfusion with the accuracy of high injection techniques to isolate clean preparations of hemocytes in Aedes mosquitoes. This method facilitates studies requiring knowledge of the types of hemocytes and their abundance.

 Mosquitoes are vectors for a number  of disease-causing pathogens such as the yellow fever virus, malaria parasites  and filarial worms. Laboratories are ...

Quantitative Analyses of all Influenza Type A Viral Hemagglutinins and Neuraminidases using Universal Antibodies in Simple Slot Blot Assays

Hemagglutinin (HA) and neuraminidase (NA) are two surface proteins of influenza viruses which are known to play important roles in the viral life cycle and the induction of protective immune responses1,2. As the main target for neutralizing antibodies, HA is currently used as the influenza vaccine potency marker and is measured by single radial immunodiffusion (SRID)3. However, the dependence of SRID on the availability of the corresponding subtype-specific antisera causes a minimum of 2-3 months delay for the release of every new vaccine. Moreover, despite evidence that NA also induces protective immunity4, the amount of NA in influenza vaccines is not yet standardized due to a lack of appropriate reagents or analytical method5. Thus, simple alternative methods capable of quantifying HA and NA antigens are desirable for rapid release and better quality control of influenza vaccines.
Universally conserved regions in all available influenza A HA and NA sequences were identified by bioinformatics analyses6-7. One sequence (designated as Uni-1) was identified in the only universally conserved epitope of HA, the fusion peptide6, while two conserved sequences were identified in neuraminidases, one close to the enzymatic active site (designated as HCA-2) and the other close to the N-terminus (designated as HCA-3)7. Peptides with these amino acid sequences were synthesized and used to immunize rabbits for the production of antibodies. The antibody against the Uni-1 epitope of HA was able to bind to 13 subtypes of influenza A HA (H1-H13) while the antibodies against the HCA-2 and HCA-3 regions of NA were capable of binding all 9 NA subtypes. All antibodies showed remarkable specificity against the viral sequences as evidenced by the observation that no cross-reactivity to allantoic proteins was detected. These universal antibodies were then used to develop slot blot assays to quantify HA and NA in influenza A vaccines without the need for specific antisera7,8. Vaccine samples were applied onto a PVDF membrane using a slot blot apparatus along with reference standards diluted to various concentrations. For the detection of HA, samples and standard were first diluted in Tris-buffered saline (TBS) containing 4M urea while for the measurement of NA they were diluted in TBS containing 0.01% Zwittergent as these conditions significantly improved the detection sensitivity. Following the detection of the HA and NA antigens by immunoblotting with their respective universal antibodies, signal intensities were quantified by densitometry. Amounts of HA and NA in the vaccines were then calculated using a standard curve established with the signal intensities of the various concentrations of the references used. 
Given that these antibodies bind to universal epitopes in HA or NA, interested investigators could use them as research tools in immunoassays other than the slot blot only.

A simple slot blot method was developed for the quantification of influenza viral hemagglutinin and neuraminidase using universal antibodies targeting their most conserved sequences identified through bioinformatics analyses. This innovative approach may provide a useful alternative to quantitative determination of all viral hemagglutinin and neuraminidase.

Hemagglutinin (HA) and neuraminidase (NA) are two surface proteins of influenza viruses which are known to play important roles in the viral life cycle ...

A Simple Chelex Protocol for DNA Extraction from Anopheles spp.

Endemic countries are increasingly adopting molecular tools for efficient typing, identification and surveillance against malaria parasites and vector mosquitoes, as an integral part of their control programs1,2,3,4,5. For sustainable establishment of these accurate approaches in operations research to strengthen malaria control and elimination efforts, simple and affordable methods, with parsimonious reagent and equipment requirements are essential6,7,8. Here we present a simple Chelex-based technique for extracting malaria parasite and vector DNA from field collected mosquito specimens.
We morphologically identified 72 Anopheles gambiae sl. from 156 mosquitoes captured by pyrethrum spray catches in sleeping rooms of households within a 2,000 km2 vicinity of the Malaria Institute at Macha. After dissection to separate the head and thorax from the abdomen for all 72 Anopheles gambiae sl. mosquitoes, the two sections were individually placed in 1.5 ml microcentrifuge tubes and submerged in 20 &mu;l of deionized water. Using a sterile pipette tip, each mosquito section was separately homogenized to a uniform suspension in the deionized water. Of the ensuing homogenate from each mosquito section, 10 &mu;l was retained while the other 10 &mu;l was transferred to a separate autoclaved 1.5 ml tube. The separate aliquots were subjected to DNA extraction by either the simplified Chelex or the standard salting out extraction protocol9,10. The salting out protocol is so-called and widely used because it employs high salt concentrations in lieu of hazardous organic solvents (such as phenol and chloroform) for the protein precipitation step during DNA extraction9.
Extracts were used as templates for PCR amplification using primers targeting arthropod mitochondrial nicotinamide adenine dinucleotide dehydrogenase (NADH) subunit 4 gene (ND4) to check DNA quality11, a PCR for identification of Anopheles gambiae sibling species10 and a nested PCR for typing of Plasmodium falciparum infection12. Comparison using DNA quality (ND4) PCR showed 93% sensitivity and 82% specificity for the Chelex approach relative to the established salting out protocol. Corresponding values of sensitivity and specificity were 100% and 78%, respectively, using sibling species identification PCR and 92% and 80%, respectively for P. falciparum detection PCR. There were no significant differences in proportion of samples giving amplicon signal with the Chelex or the regular salting out protocol across all three PCR applications. The Chelex approach required three simple reagents and 37 min to complete, while the salting out protocol entailed 10 different reagents and 2 hr and 47 min' processing time, including an overnight step. Our results show that the Chelex method is comparable to the existing salting out extraction and can be substituted as a simple and sustainable approach in resource-limited settings where a constant reagent supply chain is often difficult to maintain.

A Simple Chelex Protocol for DNA Extraction from Anopheles spp.

A rapid and affordable way to extract quality malaria parasite and vector DNA from mosquito specimens is described. Capitalizing on chelating properties of Chelex resin, the simple method enables genotyping of malaria parasites in mosquito mid-gut and salivary gland phases, as well as molecular identification of the Anopheles sibling species by PCR.

Endemic countries are increasingly adopting molecular tools for  efficient typing, identification and surveillance against malaria parasites and  vector ...

Genetic Manipulation in &Delta;ku80 Strains for Functional Genomic Analysis of Toxoplasma gondii

Targeted genetic manipulation using homologous recombination is the method of choice for functional genomic analysis to obtain a detailed view of gene function and phenotype(s). The development of mutant strains with targeted gene deletions, targeted mutations, complemented gene function, and/or tagged genes provides powerful strategies to address gene function, particularly if these genetic manipulations can be efficiently targeted to the gene locus of interest using integration mediated by double cross over homologous recombination.
Due to very high rates of nonhomologous recombination, functional genomic analysis of Toxoplasma gondii has been previously limited by the absence of efficient methods for targeting gene deletions and gene replacements to specific genetic loci. Recently, we abolished the major pathway of nonhomologous recombination in type I and type II strains of T. gondii by deleting the gene encoding the KU80 protein1,2. The &Delta;ku80 strains behave normally during tachyzoite (acute) and bradyzoite (chronic) stages in vitro and in vivo and exhibit essentially a 100% frequency of homologous recombination. The &Delta;ku80 strains make functional genomic studies feasible on the single gene as well as on the genome scale1-4.
Here, we report methods for using type I and type II &Delta;ku80&Delta;hxgprt strains to advance gene targeting approaches in T. gondii. We outline efficient methods for generating gene deletions, gene replacements, and tagged genes by targeted insertion or deletion of the hypoxanthine-xanthine-guanine phosphoribosyltransferase (HXGPRT) selectable marker. The described gene targeting protocol can be used in a variety of ways in &Delta;ku80 strains to advance functional analysis of the parasite genome and to develop single strains that carry multiple targeted genetic manipulations. The application of this genetic method and subsequent phenotypic assays will reveal fundamental and unique aspects of the biology of T. gondii and related significant human pathogens that cause malaria (Plasmodium sp.) and cryptosporidiosis (Cryptosporidium).

Genetic Manipulation in &lt;em&gt;&amp;Delta;ku80&lt;/em&gt; Strains for Functional Genomic Analysis of &lt;em&gt;Toxoplasma gondii&lt;/em&gt;

Here we report a method for using type I and type II &Delta;ku80 strains of Toxoplasma gondii to efficiently generate targeted gene deletions and gene replacements for functional genomic analysis.

Targeted genetic manipulation using homologous recombination is the method of choice for functional genomic analysis to obtain a detailed view of gene ...

Retroviral Transduction of Helper T Cells as a Genetic Approach to Study Mechanisms Controlling their Differentiation and Function

Helper T cell development and function must be tightly regulated to induce an appropriate immune response that eliminates specific pathogens yet prevents autoimmunity. Many approaches involving different model organisms have been utilized to understand the mechanisms controlling helper T cell development and function. However, studies using mouse models have proven to be highly informative due to the availability of genetic, cellular, and biochemical systems. One genetic approach in mice used by many labs involves retroviral transduction of primary helper T cells. This is a powerful approach due to its relative ease, making it accessible to almost any laboratory with basic skills in molecular biology and immunology. Therefore, multiple genes in wild type or mutant forms can readily be tested for function in helper T cells to understand their importance and mechanisms of action. We have optimized this approach and describe here the protocols for production of high titer retroviruses, isolation of primary murine helper T cells, and their transduction by retroviruses and differentiation toward the different helper subsets. Finally, the use of this approach is described in uncovering mechanisms utilized by microRNAs (miRNAs) to regulate pathways controlling helper T cell development and function.

Many experimental systems have been utilized to understand the mechanisms regulating T cell development and function in an immune response. Here a genetic approach using retroviral transduction is described, which is economic, time efficient, and most importantly, highly informative in identifying regulatory pathways.

Helper T cell development and function must be tightly regulated to induce an appropriate immune response that eliminates specific pathogens yet prevents ...

A Simple Red Blood Cell Lysis Method for the Establishment of B Lymphoblastoid Cell Lines

A number of methods exist for the transformation of B lymphocytes by the Epstein Barr virus in vitro into immortalized cell lines. We have developed a new method with a powerful and simple strategy for the establishment of B-LCLs, the red blood cell lysis method. This method simplified the PBMC separation procedure with red blood cell removal, and used as little as 0.5 mL of whole blood for establishing EBV-immortalized cell lines, which can proliferate to large cell numbers in a relatively short amount time with a 100% success rate. The method is simple, reliable, time saving, and applicable to treating a large number of the clinical samples.

A simple red blood cell lysis method for establishing B-LCLs was developed with high immortalization efficiency, requiring a small amount of blood and saving time from initiation to cryopreservation.

A number of methods exist for the transformation of B lymphocytes by the Epstein Barr virus in vitro into immortalized cell lines. We have developed a new ...

Phenotypic Analysis of Rodent Malaria Parasite Asexual and Sexual Blood Stages and Mosquito Stages

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. However, effective malaria parasite targets for vaccine and chemotherapy development are still limited. This is largely due to the unavailability of relevant and practical in vivo infection models for human Plasmodium species, most notably for P.&#160;falciparum and P. vivax. Therefore, rodent malaria species have been extensively used as practical alternative in vivo models for malaria vaccine, drug targeting, immune response, and functional characterization studies of conserved Plasmodiumspp. genes. Indeed, rodent malaria models have proven to be invaluable, especially for exploring mosquito transmission and liver stage biology, and were indispensable for immunological studies. However, there are discrepancies in the methods used to evaluate the phenotypes of transgenic and wild-type asexual and sexual blood-stage parasites.&#160;Examples of these discrepancies are the choice of an intravenous vs. intraperitoneal infection of rodents with blood-stage parasites and the evaluation of male gamete exflagellation. Herein, we detail standardized experimental methods to evaluate the phenotypes of asexual and sexual blood stages in transgenic parasites expressing reporter-gene or wild-type rodent malaria parasite species. We also detail the methods to evaluate the phenotypes of malaria parasite mosquito stages (gametes, ookinetes, oocysts, and sporozoites) inside Anopheles mosquito vectors. These methods are detailed and simplified here for the lethal and non-lethal strains of P. berghei and P. yoelii but can also be applied with some adjustments to P. chabaudi and P. vinckei rodent malaria species.

Due to the striking similarities of the life cycle and biology of rodent malaria parasites to human malaria parasites, rodent malaria models have become indispensable for malaria research. Herein, we standardized some of the most important techniques used in the phenotypic analysis of wild-type and transgenic rodent malaria species.

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. ...

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro

Natural killer (NK) cells belong to the innate immune system and are a first-line anti-cancer immune defense; however, they are suppressed in the tumor microenvironment and the underlying mechanism is still largely unknown. The lack of a consistent and reliable source of NK cells limits the research progress of NK cell immunity. Here, we report an in vitro system that can provide high quality and quantity of bone marrow-derived murine NK cells under a feeder-free condition. More importantly, we also demonstrate that siRNA-mediated gene silencing successfully inhibits the E4bp4-dependent NK cell maturation by using this system. Thus, this novel in vitro NK cell differentiating system is a biomaterial solution for immunity research.

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro

Here, we present a protocol to mass-produce gene-silencing murine NK cells by using a feeder-free differentiation system for mechanistic study in vitro and in vivo.

Natural killer (NK) cells belong to the innate immune system and are a first-line anti-cancer immune defense; however, they are suppressed in the tumor ...

Quantification of Intracellular Growth Inside Macrophages is a Fast and Reliable Method for Assessing the Virulence of Leishmania Parasites

The lifecycle of Leishmania, the causative agent of leishmaniasis, alternates between promastigote and amastigote stages inside the insect and vertebrate hosts, respectively. While pathogenic symptoms of leishmaniasis can vary widely, from benign cutaneous lesions to highly fatal visceral disease forms depending on the infective species, all Leishmania species reside inside host macrophages during the vertebrate stage of their lifecycle. Leishmania infectivity is therefore directly related to its ability to invade, survive and replicate within parasitophorous vacuoles (PVs) inside macrophages. Thus, assessing the parasite&#39;s ability to replicate intracellularly serves as a dependable method for determining virulence. Studying leishmaniasis development using animal models is time-consuming, tedious and often difficult, particularly with the pathogenically important visceral forms. We describe here a methodology to follow the intracellular development of Leishmania in bone marrow-derived macrophages (BMMs). Intracellular parasite numbers are determined at 24 h intervals for 72 - 96 h following infection. This method allows for a reliable determination of the effects of various genetic factors on Leishmania virulence. As an example, we show how a single allele deletion of the Leishmania Mitochondrial Iron Transporter gene (LMIT1) impairs the ability of the Leishmania amazonensis mutant strain LMIT1/&#916;Lmit1 to grow inside BMMs, reflecting a drastic reduction in virulence compared to wild-type. This assay also allows precise control of experimental conditions, which can be individually manipulated to analyze the influence of various factors (nutrients, reactive oxygen species, etc.) on the host-pathogen interaction. Therefore, the appropriate execution and quantification of BMM infection studies provide a non-invasive, rapid, economical, safe and reliable alternative to conventional animal model studies.

Quantification of Intracellular Growth Inside Macrophages is a Fast and Reliable Method for Assessing the Virulence of Leishmania Parasites

All pathogenic Leishmania species reside and replicate inside macrophages of their vertebrate hosts. Here, we present a protocol to infect murine bone marrow-derived macrophages in culture with Leishmania, followed by precise quantification of intracellular growth kinetics. This method is useful for studying individual factors influencing host-pathogen interaction and Leishmania virulence.

The lifecycle of Leishmania, the causative agent of leishmaniasis, alternates between promastigote and amastigote stages inside the insect and vertebrate ...