Authors thank Bloomberg Philanthropies for financial support to Johns Hopkins Malaria Research Institute (JHMRI). This work would not have been possible without the expertise provided by JHMRI insect and parasitology core facilities.

Blood collections described below have been approved by the Institutional Review Board of Johns Hopkins University. P. falciparum is cultured in fresh RBCs under sterile conditions in a biosafety level 2 (BSL2) facility and caution is used to handle biological materials. After each step involving blood or blood products, every plasticware or glassware is rinsed with 10% bleach within the hood prior to proper disposal.
1. Reagents and preparation
<ol>
	<li>Parasite isolate P. falciparum&#160;NF54 (see Table of Materials) was used, which can produce infectious gametocytes for up to two months while in continuous culture. 
	NOTE: Not all culture adapted parasite lines produce gametocytes, and low passage NF54 isolates are most consistent.</li>
	<li>O+ Erythrocytes: Dilute whole blood by adding equal volume of RPMI 1640 media and centrifuge at 500 x g for 5 min at room temperature in a swing out rotor with deacceleration set at 0. Carefully remove the supernatant and buffy coat (a white band between plasma and packed erythrocytes, which contains most of white blood cells and platelets) and add equal volume of RPMI. Repeat the washing step twice and after the final wash add one pellet volume of RPMI to get 50% hematocrit for storage at 4 &#176;C. 
	NOTE: Gametocytes mature over a period of two weeks which makes it critical to begin cultures with fresh erythrocytes, typically erythrocytes drawn within a week works well.</li>
	<li>O+ Human Serum: Pool at-least 6 units of serum together to minimize the effect of normal variation in serum from different individuals. Sterilize pooled human serum using 0.2 &#181;m filtration flasks. Aliquot in portions of 50 mL or less based on the need and store at -20 &#176;C. 
	NOTE: Erythrocytes and serum needs to be from a compatible blood type for P. falciparum cultures.</li>
	<li>500x Hypoxanthine (100 mL) solution: Dissolve 0.5 g of hypoxanthine in 100 mL of 1 M NaOH. Filter sterilize and make 5-10 mL aliquots for storage. Stocks can be stored up to a year at -20 &#176;C and up to 2 weeks at 4 &#176;C.</li>
	<li>Sodium bicarbonate (Optional): Dissolve 7.5 g sodium bicarbonate in 100 mL of deionized, tissue culture grade water and filter sterilize with 0.2 &#181;m filter. 
	NOTE: This protocol uses the candle jar method to provide microaerophilic conditions for P. falciparum in vitro culture and does not require sodium bicarbonate. However, if parasites are cultured using a malaria gas mix (5% O2, 5% CO2 and 90% N2), it is important to supplement culture media with 0.2% sodium bicarbonate.</li>
	<li>Complete media: To prepare 500 mL of complete media, add 1 mL of hypoxanthine solution and 50 mL of pooled human serum to 500 mL of RPMI 1640. Add 15 mL of 7.5% sodium bicarbonate if using a malaria gas mixture. Store complete media at 4 &#176;C and discard if the color of complete media changes from orange to pink. To avoid waste, make enough complete media to be used within three days.</li>
	<li>Exflagellation media (Optional): Make exflagellation or ookinete media by dissolving 200 mg NaHCO3, 5 mg Hypoxanthine and 100 &#181;L of xanthurenic acid (from 100 mM stock in water) to 100 mL of incomplete media (RPMI 1640 with glutamine and HEPES). 
	NOTE: Exflagellation is the process of male gamete formation inside the midgut of female Anopheles mosquito few minutes after it takes blood meal infected with gametocytes. Male gametocytes of Plasmodium give rise to 8 male gametes after an exflagellation event. In vitro, this process occurs spontaneously when culture temperature is lowered to room temperature (RT) and can be observed in cultured mature gametocytes.</li>
	<li>N-Acetylglucosamine (Optional): Make 500 mM stock solution of N-acetylglucosamine in water or incomplete media, aliquot and store at -20 &#176;C.</li>
	<li>NaCl solutions: Dissolve 0.9 g, 1.6 g and 12 g NaCl in 100 mL of tissue culture grade deionized water to make 0.9%, 1.6% and 12% NaCl solutions. Filter sterilize with 0.22 &#181;m filter and store at 4 &#176;C.</li>
</ol>
2. P. falciparum asexual stage culture
<ol>
	<li>To begin parasite culture, remove a low passage frozen vial of P. falciparum NF54, from the liquid nitrogen tank and quickly thaw in water bath set at 37 &#176;C.</li>
	<li>Transfer contents (~1 mL) to a 50 mL sterile centrifuge tube and dropwise add 0.2 volume of prewarmed 12% NaCl, while gently shaking the tube to ensure even mixing. Incubate for 5 min at RT with intermittent gentle shaking.</li>
	<li>Add 9 volumes of 1.6% NaCl dropwise, while continuing gentle mixing. Centrifuge the contents at 500 x g for 5 min at RT, carefully discard the supernatant. Add 9 volumes of 0.9% NaCl dropwise, while making sure to consistently mix the parasite pellet. Centrifuge again at 500 x g for 5 min at RT. Remove the supernatant and resuspend parasites into 5 mL of complete media.</li>
	<li>Transfer contents to one well of a 6 well tissue culture plate and add 100-200 &#181;L of packed RBCs.</li>
	<li>Incubate parasite culture at 37 &#176;C in a candle jar or in a modular incubator chamber purged with special gas mix of 5% O2, 5% CO2 and 90% N2.</li>
	<li>Replace media every day and monitor the growth by making a thin blood smear by drawing a few microliters from settled RBC layer after the culture supernatant has been aspirated during regular media change.</li>
	<li>Use sterile glass Pasteur pipette to draw and then tap in the center of the glass slide to transfer a drop of culture to the slide. Place another glass slide in front of the of the drop and draw it back to contact the RBCs, quickly pushing it forward in one motion to make thin smear of RBC monolayer.</li>
	<li>Place the slide horizontally on a drying rack and let it air dry. Fix the blood smear by dropping absolute methanol onto the smear. Allow the fixed smear to dry completely and then carefully pour 10% Geimsa-stain freshly diluted in water, until blood smear is completely covered. Allow cells to stain for approximately 15 min.</li>
	<li>Wash off the excess stain by rinsing the slide under clean tap water and allow the slides to dry in vertical position.</li>
	<li>Determine parasitemia by viewing thin blood smear on a microscope using oil immersion 100x objective. Count the number of infected erythrocytes among a total of at-least 500 RBCs to determine percent parasitemia.</li>
</ol>
3. P. falciparum gametocyte culture
NOTE: Gametocyte cultures take two weeks to produce mature gametocytes infectious to mosquitoes. The steps of the gametocyte culture are outlined in Figure 1. P. falciparum isolates usually lose the ability to produce gametocyte after long term in vitro culture13. To ensure quality of gametocytes, culture should be initiated from low passage feeder culture, not more than 2 months old since thawing. Pre-warm media to 37 &#176;C and perform all the procedures on a slide warmer set at 38 &#176;C. Make sure gametocyte cultures are not out of incubator for extended period to minimize temperature fluctuations.
<ol>
	<li>Seed the gametocyte culture (day 0) using mixed asexual stage feeder culture at 0.3-1% parasitemia at 4% hematocrit.</li>
	<li>To set up a six well gametocyte culture, centrifuge 5 mL of feeder culture at 500 x g for 5 min at RT. Discard the supernatant and resuspend the pellet in 30 mL of complete media.</li>
	<li>Add 1.2 mL of packed RBCs, mix, and dispense 5 mL to each well of the 6 well plate and incubate in a candle jar at 37 &#176;C. 
	NOTE: Gametocyte culture set up described above is based on a mixed asexual stage feeder culture at 5% parasitemia.</li>
	<li>Change media daily for 15-18 days, without the addition of fresh blood, by carefully aspirating about 70-80% culture supernatant to avoid removing blood cells.</li>
	<li>Add 5 mL of fresh complete media to each well. While changing media, slowly add media using a serological pipette against the wall of the well to avoid disturbing the settled RBC layer. 
	NOTE: Because 1-2 mL of the culture medium will be left during media change, total volume of culture will be between 6-7 mL, after day 1 of gametocyte culture. While adapting this protocol, it is advisable to make blood smears every alternate day to make sure parasites are healthy. Making a blood smear can often deplete the number of cells in culture. To avoid this, draw a very small volume.</li>
	<li>To quantify mature gametocytemia, make blood smear between days 15-18 and count the number of mature gametocytes among total number of cells. 
	NOTE: Mature gametocytes can easily be identified with their classic crescent shape with smooth rounded ends (Figure 5B).</li>
	<li>Count a minimum of 1,000 RBCs and calculate the percentage of RBCs infected with mature gametocytes.</li>
	<li>To quantify exflagellation events, take 200 &#181;L of gametocyte culture and centrifuge at 500 x g for 5 min at RT in a pre-warmed tube. Resuspend the pellet in 20 &#181;L of exflagellation media and transfer to a glass slide with cover slip.</li>
	<li>After 15 min incubation at room temperature, begin counting exflagellation centers in phase contrast mode using 10x objective. Count exflagellation centers in at-least four fields to calculate exflagellation events. 
	NOTE: A detailed methodology of exflagellation assay has been described earlier by Delves et al10. More than 20 exflagellation events per field is considered suitable for membrane feeding assay.</li>
	<li>Gametocyte cultures typically have low levels of residual asexual stages. If the experiment requires pure gametocytes, treat the culture with 50 mM N-acetylglucosamine.</li>
	<li>Add 0.5 mL of N-acetylglucosamine from 10x stock solution per well (5 mL) for a minimum of 3 days while changing media to clear residual asexual stages. 
	NOTE: N-acetylglucosamine can block invasion of sexually committed merozoites too, treatment should be initiated only after day 7 of gametocyte culture.</li>
</ol>
4. Mosquito infection using standard membrane feeding assay (SMFA)
NOTE: Gametocytes grown in vitro can be fed to mosquitos using glass membrane feeders. Setting up of the blood feeding apparatus is shown in Figure 2. As described above, always maintain gametocytes at 37 &#176;C to avoid activation before they are ingested by mosquitoes. Prewarm plasticware, reagents and equipment&#39;s used with gametocyte culture to 37 &#176;C.
<ol>
	<li>Starve 3-7-day old female Anopheles mosquitoes by removing their sugar water for 6.5 h or overnight before membrane feeding.</li>
	<li>Transfer them using motorized or mouth operated aspirators to paper cups covered with double layer of fine mesh fabric. Alternatively, knock down mosquitoes in a cold room or refrigerator to transfer them to the paper cups. 
	NOTE: One-pint size cup can be used to feed up to 100 mosquitoes.</li>
	<li>Centrifuge washed blood at 500 x g for 5 min at RT and discard supernatant. Add an equal volume of freshly thawed human serum to reconstitute the whole blood. Transfer the blood and serum mix to the water bath set at 37 &#176;C for 30 min.</li>
	<li>Transfer gametocyte culture to pre-warmed 15 mL plastic tube and centrifuge at 600 x g for 5 min at 37 &#176;C. Carefully aspirate the culture supernatant and make thin blood smear to determine mature gametocytemia as described above.</li>
	<li>To make the final blood feed, dilute gametocyte pellet to a desired concentration using reconstituted whole blood. Keep the blood feed at 37 &#176;C until mosquitoes and glass feeders are ready. 
	NOTE: Mature gametocyte concentrations between 0.02 to 0.3% will provide consistent mosquito infectivity. However, it is advisable to optimize gametocytemia by feeding various concentrations to different cups of mosquitoes.</li>
	<li>Ensure that the glass membrane feeders have two openings, a narrow top opening to pipette the infected blood and a bottom opening for membrane attachment. Cut a paraffin film into squares, stretch to an even thickness and stick on to the opening of feeder to create sealed compartment for the blood feed.</li>
	<li>Attach membrane feeders to a circulatory water-bath by connecting tubing on each side to allow for the passage of warm water through the jacket around membrane feeders. 
	NOTE: Several glass feeders can be connected in series to accommodate for multiple feeding conditions.</li>
	<li>After all the feeders are connected to the water bath, turn it on and inspect for any leaks.</li>
	<li>Place feeders in the center of netting on mosquito cups with membrane-side down and secure feeders using clamps or tape. Make sure membrane feeders are not tilted to any side to allow even distribution of the blood feed and close contact of the entire feeder with the netting on cups, to allow mosquitoes access from inside the cup.</li>
	<li>Pipette about 200&#8211;1000 &#956;L of blood feed into membrane feeders. 
	NOTE: Volume of the blood feed will vary based on the number of mosquitoes and size of the membrane feeder. For 14 mm glass feeder and 50 mosquitoes use 200 &#956;L of blood feed.</li>
	<li>Make sure loading was done properly and blood feed was overlaid on the paraffin film.</li>
	<li>Allow mosquitoes to feed for about 30 min with intermittent monitoring. 
	NOTE: Gently blow mosquitoes with mouth to provide CO2 which will induce improved blood feeding.</li>
	<li>Remove unfed mosquitoes by knocking them down in a cold room. Visibly inspect mosquitoes for bulge and redness in the abdomen as a sign of fresh blood meal. Alternatively, use a mouth operated aspirator to selectively remove unfed mosquitoes.</li>
	<li>Discard unfed mosquitoes, after soaking them in 70% ethanol and put fed mosquitoes back into the mosquito cup.</li>
	<li>Double cage mosquito cups and transfer them to high containment incubator specified for mosquitoes infected with human malaria parasite P. falciparum.</li>
	<li>Place cotton pads soaked in 10% sucrose on the mosquito cups to provide them with sugar meal. Replace cotton pads every other day, until mosquitoes are dissected.</li>
</ol>
5. Mosquito mid-gut dissection and oocyst load quantification
NOTE: A schematic of midgut dissection is shown in Figure 3.
<ol>
	<li>7-8 days post blood feeding, transfer mosquito cups to 4 &#176;C for 10 min to knock down mosquitoes.</li>
	<li>Transfer mosquitoes to be dissected to a Petri dish on ice using fine-tipped forceps.</li>
	<li>Soak mosquitoes in 70% ethanol for 1-2 min to euthanize them. Add PBS to the Petri dish to wash off the ethanol after all the mosquitoes are euthanized. 
	NOTE: Before adding PBS make sure all the mosquitoes are dead.</li>
	<li>Mount a glass slide on dissecting microscope and pipette 100 &#956;L of PBS in the center. Carefully transfer one mosquito using forceps into the PBS and leave the rest of the mosquitoes in the Petri dish on ice.</li>
	<li>Using fine-tipped forceps, hold the third segment of the abdomen from the posterior end of the mosquito and with second forceps hold the junction between the thorax and abdomen. Gently pull the abdomen until the midgut is fully exposed.</li>
	<li>Discard the rest of the mosquito tissue and transfer the midgut to a clean slide containing a few drops of PBS.</li>
	<li>When the desired number of mosquitoes have been dissected, carefully remove the PBS using a pipette and stain the midgut with 0.2% mercurochrome for 2-5 min.</li>
	<li>Remove excess mercurochrome and line-up midguts on the slide so that they can be easily visualized under the light microscope.</li>
	<li>Place a cover slip and count oocysts on each midgut under 10x objective. Oocysts stain pink and are circular in shape (Figure 6C).</li>
</ol>
6. Mosquito salivary gland dissection and sporozoite load quantification
NOTE: A schematic of salivary gland dissection is shown in Figure 4.
<ol>
	<li>14-18 days post blood feeding, place mosquito cups at 4 &#176;C for 10 min. 
	NOTE: It is important to wait till mosquitoes stop moving.</li>
	<li>While waiting for mosquitoes to stop moving, prepare tools for dissection.</li>
	<li>Place a glass plate on a dissection microscope stage. Prepare 2 sets of 25 G needles on 1 mL syringes and a 9&#8221; Pasteur pipette and rubber bulb. Place 70% EtOH and dissection medium (HBSS, L-15 or PBS) in a 6 well plate.</li>
	<li>In the cold room, transfer anesthetized mosquitoes in a 6 well plate containing 70% EtOH. Gently move mosquitoes with forceps to make sure all mosquitoes are soaked, then transfer mosquitoes to the dissection medium by using forceps to wash off 70% EtOH.</li>
	<li>Move to the dissection room and place mosquitoes on a dissection microscope stage.</li>
	<li>Remove excess medium from mosquitoes, add new medium as necessary during dissection. Avoid mosquitoes from drying out yet too much liquid makes dissection more difficult.</li>
	<li>Using 2 syringes with 25 G needles, hold mosquito thorax and head, and gently pull the head upward to pull salivary glands from thorax.</li>
	<li>Disconnect salivary glands from head and thorax with a needle.</li>
	<li>Temporary place salivary glands in a separate droplet of medium on the glass plate.</li>
	<li>After dissecting 15-20 salivary glands, collect them in a low retention tube using the Pasteur pipette. 
	NOTE: Once salivary glands enter the wide part of the Pasteur pipette, they will stick to the glass and it is difficult to get them out.</li>
	<li>Pellet salivary glands by short pulse spin in a table-top centrifuge. Remove the dissection medium without disrupting the pellet and Add 100 &#181;L of new dissection medium.</li>
	<li>Grind salivary glands with a small homogenizer for 1 min to obtain sporozoites.</li>
	<li>Place 10 &#181;L of dissection medium containing sporozoites on a hemocytometer. 
	NOTE: Depending on the number of salivary glands, one may need to dilute the sporozoite solution to 1:10 to 1:50 with medium before counting.</li>
	<li>Count the number of sporozoites in two of the four quadrants and calculate number of sporozoites/mosquito:</li>
</ol>
<img alt="Equation 1" src="/files/ftp_upload/61426/61426equ01v3.jpg" />
<img alt="Equation 2" src="/files/ftp_upload/61426/61426equ02v3.jpg" />
<img alt="Equation 3" src="/files/ftp_upload/61426/61426equ03.jpg" />

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Proteolytic+Cleavage+of+the+Plasmodium+falciparum+Circumsporozoite+Protein+Is+a+Target+of+Protective+Antibodies.">Proteolytic Cleavage of the Plasmodium falciparum Circumsporozoite Protein Is a Target of Protective Antibodies.</a> Journal of Infectious Diseases. 212 (7), 1111-1119 (2015).</li><li>Swearingen, K. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interrogating+the+Plasmodium+Sporozoite+Surface:+Identification+of+Surface-Exposed+Proteins+and+Demonstration+of+Glycosylation+on+CSP+and+TRAP+by+Mass+Spectrometry-Based+Proteomics.">Interrogating the Plasmodium Sporozoite Surface: Identification of Surface-Exposed Proteins and Demonstration of Glycosylation on CSP and TRAP by Mass Spectrometry-Based Proteomics.</a> PLoS Pathogens. 12 (4), 1005606 (2016).</li><li>Ifediba, T., Vanderberg, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complete+in+vitro+maturation+of+Plasmodium+falciparum+gametocytes.">Complete in vitro maturation of Plasmodium falciparum gametocytes.</a> Nature. 294 (5839), 364-366 (1981).</li><li>Miura, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+inter-laboratory+comparison+of+standard+membrane-feeding+assays+for+evaluation+of+malaria+transmission-blocking+vaccines.">An inter-laboratory comparison of standard membrane-feeding assays for evaluation of malaria transmission-blocking vaccines.</a> Malaria Journal. 15, 463 (2016).</li><li>Miura, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Qualification+of+Standard+Membrane-Feeding+Assay+with+Plasmodium+falciparum+Malaria+and+Potential+Improvements+for+Future+Assays.">Qualification of Standard Membrane-Feeding Assay with Plasmodium falciparum Malaria and Potential Improvements for Future Assays.</a> PLoS One. 8 (3), 57909 (2013).</li></ol>

The Authors have nothing to disclose.

Methods described here have been successfully used at the Johns Hopkins Malaria Research Institute for more than 10 years15,16,17,18,19,20,21,22. Gametocytes produced using this protocol have been used for high throughput gametocytocidal assays<sup class="x...

Malaria is caused by Plasmodium parasites and is transmitted to their vertebrate hosts via infectious bite of female Anopheles mosquitoes. According to the 2019 World Health Organization (WHO) report, there were an estimated 405,000 deaths, from a total of 228 million cases of malaria1. Most of the malaria related deaths were concentrated in the African region, especially among children below five years of age. While the overall incidence rate of malaria has declined globally from 2010, in recent years decline has plateaued and additional control strategies are urgently needed to eliminate the disease.
Cyclic asexual blood stages of malaria parasites cause disease pathogenesis and a small subset of these differentiate into female and male gametocytes. Plasmodium falciparum&#160;gametocytes are unique in nature as they take 7-10 days to develop through five morphologically distinct stages. Immature gametocytes from stage I to IV are sequestered in bone marrow parenchyma and largely remain absent from peripheral circulation2,3,4,5. Erythrocytes infected with mature stage V gametocytes are released in the bloodstream and freely circulate to be taken up by mosquitoes. Once inside the mosquito midgut, gametocytes are activated, through a change in temperature and exposure to the midgut environment, transform into female and male gametes and begin development of the mosquito stages, which culminates with the infective stages of sporozoites in the mosquito salivary glands6,7.
Since Trager and Jenson8 described a standardized method to culture P. falciparum, studies on the asexual blood stages have greatly advanced. However, the lack of a reliable culture system for sexual stages has made it difficult to study P. falciparum gametocytes, transmission biology and mosquito stages. In recent years, several methods have been published which have aided laboratories in establishing gametocyte cultures9,10,11,12. This manuscript describes standardized and reliable protocol to culture P. falciparum gametocytes that can represent a valuable resource for the malaria research community. This method enables the robust production of mature and infectious gametocytes which along with a standardized mosquito feeding protocol, results in highly reliable mosquito infectivity. These methods were established to maintain uninterrupted supply of gametocytes, and mosquito stage parasites. In this manuscript, we describe a thorough gametocyte culture protocol (Figure 1), preparation of glass membrane feeders and infection of mosquitoes using these membrane feeders (Figure 2), dissection of midgut (Figure 3) and salivary gland of mosquitoes (Figure 4), and quantification of infection in mosquito after midgut and salivary gland dissection.

<table>
	<tbody>
		<tr>
			<td>10% Sugar solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10ml serological pipet</td>
			<td>Falcon</td>
			<td>357551</td>
			<td></td>
		</tr>
		<tr>
			<td>15 ml conical tube</td>
			<td>Falcon</td>
			<td>352096</td>
			<td></td>
		</tr>
		<tr>
			<td>1ml serological pipet</td>
			<td>Falcon</td>
			<td>357521</td>
			<td></td>
		</tr>
		<tr>
			<td>25 ml serological pipet</td>
			<td>Falcon</td>
			<td>357535</td>
			<td></td>
		</tr>
		<tr>
			<td>37&#176;C Incubator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml conical tube</td>
			<td>Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>5ml serological pipet</td>
			<td>Falcon</td>
			<td>357543</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well tissue culture plates</td>
			<td>Falcon</td>
			<td>353046</td>
			<td></td>
		</tr>
		<tr>
			<td>70% Ethanol</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>9&#34; glass pipet</td>
			<td>Fisherbrand</td>
			<td>13-678-6B</td>
			<td></td>
		</tr>
		<tr>
			<td>Anopheles Mosquitoes</td>
			<td>JHMRI, Insectary core</td>
			<td></td>
			<td>We use A. stephensi or A. gambiae (keele)</td>
		</tr>
		<tr>
			<td>cell counter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Circulating water bath</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>fine tip forceps</td>
			<td>Fisherbrand</td>
			<td>12-000-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Geimsa stain</td>
			<td>Sigma</td>
			<td>GS1L</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass desiccator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass membrane feeder</td>
			<td>Chemglass Life Sciences</td>
			<td>CG183570</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>Fisherbrand</td>
			<td>12-552-3</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Sigma</td>
			<td>H6648</td>
			<td></td>
		</tr>
		<tr>
			<td>Human Blood O+</td>
			<td>JHU</td>
			<td></td>
			<td>Wash RBCs three times with RPMI and refrigerate at 50% heamatocrit</td>
		</tr>
		<tr>
			<td>Human Serum O+</td>
			<td>Interstate blood bank</td>
			<td></td>
			<td>Pool at-least 6 units of serum from different donors and freeze down aliquots at -20&#176;C.</td>
		</tr>
		<tr>
			<td>Hypoxanthine</td>
			<td>Sigma</td>
			<td>H9337</td>
			<td>Make 500x stock in 1M NaOH</td>
		</tr>
		<tr>
			<td>Mercurochrome</td>
			<td>Sigma</td>
			<td>M7011</td>
			<td>Prepare 1% stock solution in PBS that can be diluted to 0.1% when needed</td>
		</tr>
		<tr>
			<td>Micro Pipette</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td></td>
			<td>Any microscope with 10x, 40x and 100x objective will work.</td>
		</tr>
		<tr>
			<td>Mosquito cups</td>
			<td>Neptune cups</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>N-acetylglucosamine</td>
			<td>Sigma</td>
			<td>A3286</td>
			<td>Optional and needed only when pure gametocytes are required.</td>
		</tr>
		<tr>
			<td>Netting</td>
			<td></td>
			<td></td>
			<td>Make sure it can contain mosquitoes and allow blood feeding</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Thermo Scientific</td>
			<td>249964</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipet tips</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetman</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plasmodium falciparum NF54</td>
			<td>BEI Resources</td>
			<td>MRA-1000</td>
			<td>Freeze down large numbers of early passage culture to make sure you have a constant supply</td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Corning</td>
			<td>CV-041-CV</td>
			<td>Media contains glutamine and HEPES</td>
		</tr>
		<tr>
			<td>Slide warmer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma</td>
			<td>S6297</td>
			<td>Optional for media, add only when using malaria gas mix during culture incubation</td>
		</tr>
		<tr>
			<td>water bath</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Xanthurenic Acid</td>
			<td>Sigma</td>
			<td>D120804</td>
			<td>For flagellation media</td>
		</tr>
	</tbody>
</table>

plasmodium falciparum gametocyte culture and mosquito infection through artificial membrane feeding

Malaria remains one of the most important public health problems, causing significant morbidity and mortality. Malaria is a mosquito borne disease transmitted through an infectious bite from the female Anopheles mosquito. Malaria control will eventually rely on a multitude of approaches, which includes ways to block transmission to, through and from mosquitoes. To study mosquito stages of malaria parasites in the laboratory, we have optimized a protocol to culture highly infectious Plasmodium falciparum gametocytes, a parasite stage required for transmission from the human host to the mosquito vector. P. falciparum gametocytes mature through five morphologically distinct steps, which takes approximately 1-2 weeks. Gametocyte culture described in this protocol is completed in 15 days and are infectious to mosquitoes from days 15-18. These protocols were developed to maintain a continuous cycle of infection competent gametocytes and to maintain uninterrupted supply of mosquito stages of the parasite. Here, we describe the methodology of gametocyte culture and how to infect mosquitoes with these parasites using glass membrane feeders.

Here we present results from a series of membrane feeds using P. falciparum NF54 gametocyte cultures generated using the protocol above (see (Figure 5). Gametocyte culture was initiated with approximately 0.5% mixed stage asexual culture on Day 0, which grew to a peak parasitemia of approximately 15% by Day 4 and Day 5. As shown in Figure 5A at this high parasitemia, the parasites are stressed and the asexual stage culture crashes. However, this stress ...

Watch this Scientific Journal Video about Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding at JoVE.com

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Detailed investigations on mosquito stages of malaria parasites are critical to design effective transmission blocking strategies. This protocol demonstrates how to effectively culture infectious gametocytes and then feed these gametocytes to mosquitoes to generate mosquito stages of P. falciparum.

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Malaria remains one of the most important public health problems, causing significant morbidity and mortality. Malaria is a mosquito borne disease ...

plasmodium-falciparum-gametocyte-culture-mosquito-infection-through

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JoVE Journal

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7.3K Views.  Johns Hopkins University. Detailed investigations on mosquito stages of malaria parasites are critical to design effective transmission blocking strategies. This protocol demonstrates how to effectively culture infectious gametocytes and then feed these gametocytes to mosquitoes to generate mosquito stages of P. falciparum.

Video: Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Harvesting Sperm and Artificial Insemination of Mice

Rodents of the genus Peromyscus (deer mice) are the most prevalent native North American mammals. Peromyscus species are used in a wide range of research including toxicology, epidemiology, ecology, behavioral, and genetic studies. Here they provide a useful model for demonstrations of artificial insemination. 

Methods similar to those displayed here have previously been used in several deer mouse studies, yet no detailed protocol has been published. Here we demonstrate the basic method of artificial insemination. This method entails extracting the testes from the rodent, then isolating the sperm from the epididymis and vas deferens. The mature sperm, now in a milk mixture, are placed in the female&#x2019;s reproductive tract at the time of ovulation. Fertilization is counted as day 0 for timing of embryo development. Embryos can then be retrieved at the desired time-point and manipulated.

Artificial insemination can be used in a variety of rodent species where exact embryo timing is crucial or hard to obtain. This technique is vital for species or strains (including most Peromyscus) which may not mate immediately and/or where mating is hard to assess. In addition, artificial insemination provides exact timing for embryo development either in mapping developmental progress and/or transgenic work. Reduced numbers of animals can be used since fertilization is guaranteed. This method has been vital to furthering the Peromyscus system, and will hopefully benefit others as well.

This protocol demonstrates methods for extracting sperm from the testes of males and then inseminating female mice. This procedure is useful when precise time is needed in developmental studies as well as transgenic work.

Rodents of the genus Peromyscus (deer mice) are the most prevalent native North American mammals.  Peromyscus species are used in a wide range of research ...

Protocol for Plasmodium falciparum Infections in Mosquitoes and Infection Phenotype Determination

Once a gene is identified as potentially refractory for malaria, it must be evaluated for its role in preventing Plasmodium infections within the mosquito.    This protocol illustrates how the extent of plasmodium infections of mosquitoes can be assayed.  The techniques for preparing the gametocyte culture, membrane feeding mosquitoes human blood, and assaying viral titers in the mosquito midgut are demonstrated.

Once a gene is identified as potentially refractory for malaria, it must be evaluated for its role in preventing Plasmodium infections within the mosquito. This protocol illustrates how the extent of plasmodium infections of mosquitoes can be assayed. The techniques for preparing the gametocyte culture, membrane feeding mosquitoes human blood, and assaying viral titers in the mosquito midgut are demonstrated.

Once a gene is identified as potentially refractory for malaria, it must be evaluated for its role in preventing Plasmodium infections within the ...

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

High-fat Feeding Paradigm for Larval Zebrafish: Feeding, Live Imaging, and Quantification of Food Intake

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich meal, which consists of an emulsion of chicken egg yolk liposomes created by sonicating egg yolk in embryo media. Detailed instructions are provided to screen larvae for egg yolk consumption so that larvae that fail to feed will not confound experimental results. The chicken egg yolk liposomes can be spiked with fluorescent lipid analogs, including fatty acids and cholesterol, enabling both systemic and subcellular visualization of dietary lipid processing. Several methods are described to mount larvae that are conducive to short- and long-term live imaging with both upright and inverted objectives at high and low magnification. Additionally presented is an assay to quantify larval food intake by extracting the lipids of larvae fed fluorescent lipid analogs, spotting the lipids on a thin layer chromatography plate, and quantifying the fluorescence. Finally, critical aspects of the procedures, important controls, options for modifying the protocols to address specific experimental questions, and potential limitations are discussed. These techniques can be applied not only to focused, hypothesis driven inquiries, but also to a variety of screens and live imaging techniques to study dietary lipid metabolism and the control of food intake.

Zebrafish are emerging as a valuable model of dietary lipid processing and metabolic disease. Described are protocols of lipid-rich larval feeds, live imaging of dietary fluorescent lipid analogs, and quantification of food intake. These techniques can be applied to a variety of screening, imaging, and hypothesis driven inquiry techniques.

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

Isolation and Culture of Human Mature Adipocytes Using Membrane Mature Adipocyte Aggregate Cultures (MAAC)

White adipose tissue (WAT) dysregulation plays a central role in development of insulin resistance and type 2 diabetes (T2D). To develop new treatments for T2D, more physiologically relevant in vitro adipocyte models are required. This study describes a new technique to isolate and culture mature human adipocytes. This method is entitled MAAC (membrane mature adipocyte aggregate culture), and compared to other adipocyte in vitro models, MAAC possesses an adipogenic gene signature that is the closest to freshly isolated mature adipocytes. Using MAAC, adipocytes can be cultured from lean and obese patients, different adipose depots, co-cultured with different cell types, and importantly, can be kept in culture for 2 weeks. Functional experiments can also be performed on MAAC including glucose uptake, lipogenesis, and lipolysis. Moreover, MAAC responds robustly to diverse pharmacological agonism and can be used to study adipocyte phenotypic changes, including the transdifferentiation of white adipocytes into brown-like fat cells.

Isolation and Culture of Human Mature Adipocytes Using Membrane Mature Adipocyte Aggregate Cultures MAAC

Membrane mature adipocyte aggregate culture (MAAC) is a new method to culture mature human adipocytes. Here we detail how to isolate adipocytes from human adipose and how to set up MAAC.

White adipose tissue (WAT) dysregulation plays a central role in development of insulin resistance and type 2 diabetes (T2D). To develop new treatments ...

Large-Scale Preparation of Synovial Fluid Mesenchymal Stem Cell-Derived Exosomes by 3D Bioreactor Culture

Exosomes secreted by mesenchymal stem cells (MSCs) have been suggested as promising candidates for cartilage injuries and osteoarthritis treatment. Exosomes for clinical application require large-scale production. To this aim, human synovial fluid MSCs (hSF-MSCs) were grown on microcarrier beads, and then cultured in a dynamic three-dimension (3D) culture system. Through utilizing 3D dynamic culture, this protocol successfully obtained large-scale exosomes from SF-MSC culture supernatants. Exosomes were harvested by ultracentrifugation and verified by a transmission electron microscope, nanoparticle transmission assay, and western blotting. Also, the microbiological safety of exosomes was detected. Results of exosome detection suggest that this approach can produce a large number of good manufacturing practices (GMP)-grade exosomes. These exosomes could be utilized in exosome biology research and clinical osteoarthritis treatment.

Here, we present a protocol to produce a large number of GMP-grade exosomes from synovial fluid mesenchymal stem cells using a 3D bioreactor.

Exosomes secreted by mesenchymal stem cells (MSCs) have been suggested as promising candidates for cartilage injuries and osteoarthritis treatment. ...

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

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