恶性疟原虫 通过人工膜喂养进行配子体培养和蚊子感染

Gametocytes are responsible for malaria parasite transmission. Generating infectious gametocytes in the lab is key to studying transmission biology in mosquito stages and to generating sporozoites for later-stage results. These methods are simple for any laboratory to adopt, and visual demonstration will aid in the sharing of these procedures with the world-wide malaria community.

Demonstrating the procedure with me will be my colleagues, Godfree Mlambo, our senior research associate, and Sachie Kanatani, our postdoctoral fellow, both from the Johns Hopkins Malaria Research Institute. To set up a gametocyte culture, spin down five milliliters of feeder culture in a centrifuge, and re-suspend the pellet in 30 milliliters of complete medium. Mix 1.2 milliliters of packed red blood cells with the tube contents, and add five milliliters of the resulting suspension to each well of a six-well culture plate.

Then place the plate in a candle jar at 37 degrees Celsius. Every day for the next 15 to 18 days, carefully aspirate 70 to 80%of the culture supernatant from each well and use a serological pipette to add five milliliters of fresh medium without blood cells down the wall of each well to feed the cultures without disturbing the settled red blood cell layer. To quantify mature gametocytemia, on day 15 to 18, make a blood smear, and count the number of matured gametocytes in a minimum of 1000 red blood cells to allow calculation of the percentage of mature, gametocyte-infected red blood cells.

To quantify the exflagellation events, centrifuge 200 microliters of the gametocyte culture in a pre-warmed tube, and re-suspend the pellet in 20 microliters of exflagellation medium. Transfer the gametocyte suspension to a glass slide with a cover slip, and incubate the slide for 15 minutes at room temperature. At the end of the incubation, use a 10X objective to count the exflagellation centers in phase contrast mode in at least four fields to allow calculation of the exflagellation events.

To infect mosquitoes using the standard membrane feeding assay, remove the sugar water from a three to seven-day-old female anopheles mosquito cage for 6.5 to 18 hours, and use an aspirator to transfer up to 100 mosquitoes into individual paper cups covered with a double-layer of fine mesh fabric. Collect the washed blood by centrifugation, and reconstitute the blood with an equal volume of freshly-thawed human serum. Pre-warm reconstituted blood in a 37-degrees Celsius water bath for 30 minutes, transfer the gametocyte culture into pre-warmed, 15-milliliter plastic tubes for centrifugation, and make a thin blood smear of the pellet to determine the mature gametocytemia as demonstrated.

To make the final blood feed, use reconstituted whole blood to dilute the gametocyte pellet to the appropriate experimental concentration, and place the blood feed at 37 degrees Celsius until the mosquitoes and glass feeders are ready. Stretch a square piece of paraffin film to an even thickness and place the film over the top opening of the feeder to create a sealed compartment for the blood feed. Use a connecting tube on each side to attach the membrane feeder to a circulatory water bath to allow the passage of warm water through the jacket around the feeder.

When all of the feeders for the experiment have been connected, turn on the bath and check all of the equipment for leaks. Place the feeders in the center of the netting on the mosquito cups, membrane side down, and secure the feeders to the netting in an upright position. When all of the feeders have been optimally positioned, add 200 to 1000 microliters of blood feed to each membrane feeder, and check that the feed has been properly overlaid onto the paraffin film.

After about 30 minutes of feeding with intermittent monitoring, knock down the mosquitoes in a cold room and visibly inspect the insects for bulge and redness in the abdomen. Place the fed mosquitoes back into the mosquito cups, and place cotton pads soaked in 10%sucrose on the mosquito cups to provide them with a sugar meal and then double-cage the cups to secure infected mosquitoes, then transfer the mosquitoes to a high-containment incubator specifically for mosquitoes infected with the human malaria parasite, P.falciparum. To harvest the midguts from the infected mosquitoes, pipette 100 microliters of PBS into the center of a glass slide mounted onto a dissecting microscope stage, and use forceps to carefully transfer one mosquito into the PBS.

Use fine-tipped forceps to grasp the segment of the abdomen from the posterior end of the mosquito, and use a second pair of forceps to hold the junction between the thorax and the abdomen, then gently pull the abdomen until the midgut is fully exposed. When all of the mosquitoes have been dissected in the same manner, use a pipette to carefully remove the PBS, and stain in the midguts with 0.2%Mercurochrome for two to five minutes. At the end of the incubation, remove the excess dye, and line the midguts on the slide for easy visualization under a light microscope, then place a cover slip over the midguts and count the number of oocysts in each midgut under the 10X objective.

For salivary gland harvest, place the mosquitoes in a glass plate under a dissecting microscope in just enough medium to keep the mosquito tissue hydrated and use two syringes equipped with 25-gauge needles to grasp mosquito thorax and head. Gently pull the head upward to pull the salivary glands from the thorax, and use a needle to disconnect the salivary glands from the head and thorax. When 15 to 20 salivary glands have been collected, use a Pasteur pipette to transfer the glands to a low-retention tube, and pellet the glands by a short pull spin in a tabletop centrifuge.

Re-suspend the salivary gland in 100 microliters of fresh dissection medium, and use a small homogenizer to grind the salivary glands for one minute to release the sporozoites, then count the number of sporozoites in 10 microliters of dissection medium in two quadrants of a hemocytometer. Here, a time course of P.falciparum NF54 gametocyte culture growth, as demonstrated, can be observed. The gametocyte culture was initiated with an approximately 0.5%mixed-stage asexual culture on day zero that grew to a peak parasitemia of approximately 15%by days four and five.

At this high parasitemia, the parasites are stressed, resulting in crashing of the asexual stage culture. This stress, however, results in the induction of gametocytogenesis, with early gametocytes appearing around day six to seven, and asexual parasitemia slowly declining while remaining at a low level. The majority of the gametocytes matured to stage five by day 15, at which point they become infectious to mosquitoes and are ready to be fed.

These representative images of Giemsa's stain blood smears show gametocyte culture progression at different time points after gametocyte culture initiation. Here, results from series-of-membrane feeds using P.falciparum NF54 gametocytes generated using this protocol can be observed. Between 8 to 10 days after blood feeding, the number of oocysts varies both within and between experiments, and requires 25 to 50 mosquitoes per cohort to determine the effects of various experimental conditions.

As shown in the table, overall, the average number of sporozoites in this analysis was consistent. However, there was one experiment in which zero sporozoites were obtained, illustrating the occasional failure of the assay. The success of this protocol depends on how well the gametocytes infect the mosquitoes.

Keeping the gametocytes as still as possible when changing the medium during the two-week culture helps. Using these methods, laboratories can perform studies on gametocyte biology, transmission blocking, vector and parasite interactions, and the sporozoite stages.