Funded in part by NIH grant RO1AI015503. Schistosome-infected mice were provided by the NIAID Schistosomiasis Resource Center at the Biomedical Research Institute (Rockville, MD) through NIH-NIAID Contract HHSN272201000005I for distribution through BEI Resources.

All animal care and experimental procedures were approved by the Institutional Animal Care and Use Committee of the University of Wisconsin-Madison under Protocol no. V005717. The following protocol involves working in a Biosafety Level 2 (BSL2) facility for human pathogens, although none of the schistosome stages depicted in this protocol are infective to humans or other mammals. Follow institutional policies for handling Risk Group 2 (RG2) human pathogens.
NOTE: We use female Swiss-Webster mice (6-week old) due to their higher susceptibility to infection10, thereby yielding larger egg burdens. Tucker et al. describe the mouse and snail infection protocols used in this model system11. The Table of Materials lists all of the specific equipment, materials, reagents and animal sources needed to carry out this experimental protocol.
1. Processing of infected livers
<ol>
	<li>Euthanize mice, 6-7 weeks post-infection, by CO2 asphyxiation (rate of 10-30% of the chamber&#39;s volume per minute). Monitor mice for cessation of respiration and heartbeat. 
	NOTE: Typically, up to 20 mice may be processed at a given time.</li>
	<li>After transferring euthanized mice to a biosafety cabinet, place mice on their backs and spray their ventral surfaces with 70% ethanol. Let soak for 1 min.</li>
	<li>Pinch and pull up skin of the lower abdomen and cut across the base of the pinched skin closest to the abdomen with sterile surgical scissors. Pull the cut skin anteriorly (i.e., toward the head) to reveal the liver. Note that the liver should be enlarged and speckled due to an egg-induced granulomatous response (Figure 2A).</li>
	<li>Remove the liver and place it in a beaker containing 200 mL of sterile 1.2% NaCl solution containing pen/strep.</li>
	<li>Repeat steps 1.3 and 1.4 with remaining mice. 
	NOTE: Twenty mice are usually processed in a single harvest.</li>
	<li>Place the livers in a sterile Petri dish and remove non-liver fat/connective tissues with sterile forceps.</li>
	<li>Transfer trimmed/cleaned livers to a sterile 250-mL centrifuge bottle containing ~200 mL of sterile 1.2% NaCl solution with pen/strep.</li>
	<li>Vigorously shake the bottle containing the livers and, using a sterile disposable pipette, remove the excess surface foam/floating debris. Slowly pour off the remaining saline solution into a container or sink containing 1% bleach (disinfectant).</li>
	<li>Add ~200 mL of fresh saline solution to the livers and repeat step 1.8 three more times.</li>
</ol>
2. Liver extraction and miracidial isolation
<ol>
	<li>Place the livers into a small sterile stainless-steel blender cup and add ~2 mL of 1.2% NaCl solution (Figure 2B).</li>
	<li>Cover the cup with foil and a petri dish to prevent spills, and blend for 1 min by alternating low and high speed (20 s low speed/10 s high speed; repeat) (Figure 2C).</li>
	<li>Evenly distribute the blended liver suspension into sterile 250-mL centrifuge bottles (for 20 livers use 4 bottles).</li>
	<li>Add sterile saline with antibiotics to ~200 mL total volume/bottle. Weigh/balance bottles prior to centrifugation.</li>
	<li>Centrifuge the blended livers at 290 x g for 15 min at 4 oC. Gently pour off supernatants into a container with bleach prior to discarding.</li>
	<li>Repeat steps 2.4-2.5 once. Be sure to thoroughly resuspend liver sediment prior to centrifugation.</li>
	<li>After discarding the last saline wash (Step 2.6), add 200 mL sterile pond water with pen/strep to the pelleted liver tissue, shake to resuspend sediment, and transfer the suspension to a sterilized 1-L volumetric flask that has been completely covered with aluminum foil, except for the top 3 cm of the neck (Figure 2D). 
	NOTE: Use two bottles (= 10 livers) per 1-L flask. Pond water simulates the natural freshwater environment needed to stimulate miracidial hatching from eggs.</li>
	<li>Using sterile pond water, fill up the flask to approximately 3 cm above the foil covering on neck. Then, without delay (as miracidia soon begin to hatch), remove residual foam and liver tissue floating to the surface by quickly pipetting up ~12 mL of pond water containing debris and discarding it into a separate discard tube. Replace with fresh sterile pond water.</li>
	<li>Repeat cleaning Step 2.8, three more times, checking for presence of miracidia in the discard tube. Cease repetition of cleaning step if miracidia appear in the wash solution.</li>
	<li>After the last wash, slowly add ~12 mL of warm sterile pond water (pre-warmed to 30 oC) to create a temperature gradient with clean, sterile warm water on top. 
	NOTE: This is best accomplished by slowly discharging water along the side of the flask neck to avoid mixing.</li>
	<li>Place a small Petri dish cover over the flask opening and shine a bright light across the top of the flask above the foil cover to illuminate the upper surface of water. 
	NOTE: Typically, within 5-10 min, swimming miracidia, attracted to the light, will concentrate in large numbers within the first 2-3 cm of pond water (Figure 3A).</li>
</ol>
3. Miracidial harvesting and cultivation
<ol>
	<li>When miracidia have amassed at the surface after 10 min, using a sterile transfer pipet, remove ~8 mL of pond water and transfer to a sterile 15-mL centrifuge tube. Place the tube containing miracidia on ice.</li>
	<li>Add back 8 mL of warm sterile pond water along the inside of the volumetric flask and wait another 10 min.</li>
	<li>Using new tubes each time, repeat steps 3.1 and 3.2 three more times. After the 4th collection, keep the final tube on ice for 10 min to allow the miracidia to settle. This set of tubes comprises the first harvest.</li>
	<li>Centrifuge tubes containing miracidia at 290 x g for 1 min at 4 oC in a pre-cooled refrigerated centrifuge.</li>
	<li>Immediately remove most of the supernatant (~7 mL) being careful not to disturb the parasite pellet (Figure 3B).</li>
	<li>Pool all the miracidia from this first harvest into one tube, then rinse all of the original tubes with ~1 mL sterile pond water and add to the &#34;pooled&#34; tube.</li>
	<li>Allow parasites to settle on ice for ~5 min, and again centrifuge the parasites at 290 x g for 1 min at 4 oC.</li>
	<li>Carefully remove the supernatant and add 6-8 mL sterile Chernin&#39;s balanced salt solution (CBSS+; see Table of Materials) containing pen/strep.</li>
	<li>Gently suspend the miracidia and aliquot them into a 24-well culture plate (1 mL per well), followed by rinsing the tube in 6-8 mL CBSS+ and adding 1 mL of rinse to each well. Incubate parasites at 26 oC under normoxic conditions. Concentrations of isolated miracidia will vary, but will usually average ~7000/mL.</li>
	<li>If parasites are still concentrating at the light source, repeat Steps 3.1 to 3.9 to continue harvesting late-hatching miracidia but increase the time interval between collections to 15-20 min. 
	NOTE: This new set of tubes represents the second harvest. However, because miracidial numbers typically are low, larvae from all tubes usually are pooled into a single culture well containing 2 mL CBSS+.</li>
	<li>At 24 h post-harvest and cultivation, gently resuspend sporocysts in their wells by pipetting.</li>
	<li>Wait a few min to allow parasites to settle to the bottom of the wells. Once they have settled, carefully remove the supernatant (~1.5 mL), which will contain most of the ciliated epidermal plates shed by miracidia during larval transformation. This &#34;transformation&#34; supernatant may either be discarded or incorporated into follow-up studies of larval secretory products.</li>
	<li>Add 1.5 mL of fresh CBSS+ to each well and gently pipette to resuspend sporocysts. Repeat step 3.12 if further washing or changing to a different medium is desired.</li>
</ol>

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	<li>Colley, D. G., Bustinduy, A. L., Secor, W. E., King, C. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+schistosomiasis.">Human schistosomiasis.</a> Lancet. 383, 2253-2264 (2014).</li><li>WHO. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Schistosomiasis:+number+of+people+treated+worldwide+in+2013.">Schistosomiasis: number of people treated worldwide in 2013.</a> Wkly. Epidemiol. Rec. 5 (90), 25-32 (2015).</li><li>Yoshino, T. P., Gourbal, B., Th&#233;ron, A., Jamieson, B. G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Schistosoma+sporocysts.">Schistosoma sporocysts.</a> Schistosoma: biology, pathology and control. , 118-148 (2017).</li><li>Bayne, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Successful+parasitism+of+vector+snail+Biomphalaria+glabrata+by+the+human+blood+fluke+(trematode)+Schistosoma+mansoni:+a+2009+assessment.">Successful parasitism of vector snail Biomphalaria glabrata by the human blood fluke (trematode) Schistosoma mansoni: a 2009 assessment.</a> Mol. Biochem. Parasitol. 165 (1), 8-18 (2009).</li><li>Yoshino, T. P., Coustau, C., Toledo, R., Fried, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunobiology+of+Biomphalaria-trematode+interactions.">Immunobiology of Biomphalaria-trematode interactions.</a> Biomphalaria snails and larval trematodes. , 159-189 (2011).</li><li>Coustau, C., 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=Advances+in+gastropod+immunity+from+the+study+of+the+interaction+between+the+snail+Biomphalaria+glabrata+and+its+parasites:+A+review+of+research+progress+over+the+last+decade.">Advances in gastropod immunity from the study of the interaction between the snail Biomphalaria glabrata and its parasites: A review of research progress over the last decade.</a> Fish Shellfish Immunol. 46 (1), 5-16 (2015).</li><li>Bayne, C. J., Hahn, U. K., Bender, R. 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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Schistosomiasis.">Schistosomiasis.</a> Curr Proto Immunol. 103, 19.1:19.1.1-19.1.58 (2013).</li><li>Basch, P. F., DiConza, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+miracidium-sporocyst+transition+in+Schistosoma+mansoni:+surface+changes+in+vitro+with+ultrastructural+correlation.">The miracidium-sporocyst transition in Schistosoma mansoni: surface changes in vitro with ultrastructural correlation.</a> J. Parasitol. 60 (6), 935-941 (1974).</li><li>Hansen, E. 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USA. 96, 4965-4970 (1999).</li><li>Yoshino, T. P., Bayne, C. J., Bickham, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molluscan+cells+in+culture:+primary+cell+cultures+and+cell+lines.">Molluscan cells in culture: primary cell cultures and cell lines.</a> Can. J. Zool. 91, 391-404 (2013).</li><li>Coustau, C., Yoshino, T. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flukes+without+snails:+Advances+in+the+in+vitro+cultivation+of+intramolluscan+stages+of+trematodes.">Flukes without snails: Advances in the in vitro cultivation of intramolluscan stages of trematodes.</a> Exp. Parasitol. 94, 62-66 (2000).</li><li>Milligan, J. N., Jolly, E. 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Authors have nothing to disclose.

Miracidia of S. mansoni, isolated and manipulated as described herein, can infect only the snail intermediate host, and therefore do not represent a human biohazard during this phase of larval development. However, to avoid accidental exposure/infection of snails, care should be taken to perform miracidial isolations in a different location from areas where susceptible Biomphalaria snail species may be present or maintained. A separate room, registered as BSL2 space, is highly recommended. In addition, ...

The human blood flukes Schistosoma spp., are the causative agents of schistosomiasis, a neglected tropical disease afflicting an estimated 230 million people worldwide1. The most widespread species, Schistosoma mansoni, is geographically distributed in South America, the Caribbean archipelago, the Middle East and sub-Saharan Africa2. The life cycle of S. mansoni, and other schistosomes, is complex, involving a mammalian definitive host and freshwater snails of the genus Biomphalaria that serve as obligate intermediate hosts.
S. mansoni male and female adult worm pairs living in the posterior mesenteric veins of the mammalian host reproduce, resulting in the release of embryonated eggs (ova) that become lodged in the small venules of the intestinal mucosa. Eggs then breaks through the vessel walls, migrate through mucosal tissue, and eventually enter the intestinal lumen where they are voided with the feces. When ova enter freshwater and free-swimming larvae (miracidia) hatch, they must, in short order, find a suitable Biomphalaria snail to infect in order to continue its life cycle. This infection process involves miracidial attachment to the snail&#39;s outer body surface followed by active penetration and entry of the larva into the host. Soon after entry, the miracidium begins to shed its outer ciliated epidermal plates as it forms a syncytium that will become the new outer surface (tegument) of the next larval stage, the primary or mother sporocyst. Through asexual reproduction, each primary sporocyst produces and releases a second generation of sporocysts, termed secondary or daughter sporocysts, which in turn, generate and release large numbers of the final intramolluscan stage, the cercaria3. Upon escape from the snail host, free-swimming cercariae are capable of attaching to and directly penetrating the skin of a human or other suitable mammalian host. Upon entry into their new host, cercariae transform to the parasitic schistosomula stage and invade the vascular system. They, in turn, migrate to the lung and then to the hepatic vein where they mature to adult worms, form male and female pairs and travel to the mesenteric veins to complete their life cycle.
Successful intramolluscan or &#34;snail phase&#34; development of larval schistosomes is essential to continuation of the cycle of human host-to-host transmission. Of critical importance is the period following entry of the free-swimming miracidium into the snail host and its early transformation to the primary sporocyst stage. Depending on the physiological and immunological compatibility between host and parasite, it is during this phase of larval development that initial success or failure to establish infections is determined4,5,6. Subsequent development of primary sporocysts capable of asexually producing secondary sporocysts, which then generate human-infective cercariae, further require a permissive physiological host environment that provides for all of the parasite&#39;s growth and reproductive needs.
To date, little is known about the underlying physiological, biochemical and molecular processes controlling schistosome-snail interactions, especially the molecules and pathways involved in regulating larval development and reproduction, or modulating host immune responses. Ready access to large numbers of these larval stages under in vitro culture conditions that permit experimental manipulation would greatly facilitate the discovery of developmental pathways and underlying mechanisms of cell growth, differentiation and reproduction. Critical parasite pathways or mechanisms, identified through in vitro experimentation, could then be used to disrupt larval growth/reproduction in the snail host, or to identify snail host immune mechanisms involved in recognition and elimination of miracidial or sporocyst stages.
In this article and video presentation, we provide a detailed description of a method for isolating large numbers of free-swimming S. mansoni miracidia for introduction into in vitro culture that may then be subjected to follow-up experimental manipulation (see Figure 1 for a schematic overview of the protocol workflow). Although similar procedures have been described previously7,8,9, we felt that a detailed protocol would be useful to researchers who wish to employ this model to address still unanswered questions related to intramolluscan larval development, cellular proliferation, and schistosome-snail immune interactions. In addition, this approach could easily be adapted for isolation of eggs from other medically important trematodes such as bladder-dwelling S. haematobium, liver flukes or lung flukes.

<table><tbody><tr><td>&lt;strong&gt;Chernin&#039;s balanced salt solution (CBSS+)&lt;/strong&gt;</td><td></td><td></td><td>&lt;strong&gt;For 1L of solution&lt;/strong&gt;</td></tr><tr><td>2.8 g sodium chloride</td><td>Fisher Scientific</td><td>S271-3</td><td>Dissolve salts, except calcium chloride, and&amp;nbsp;</td></tr><tr><td>0.15 g of potassium chloride&amp;nbsp;</td><td>Sigma-Aldrich</td><td>P5405</td><td>sugars in 800 mL ddH2O</td></tr><tr><td>0.07 g sodium phosphate, dibasic anhydrous</td><td>Fisher Scientific</td><td>S374-500</td><td>Dissolve calcium chloride separately in 200&amp;nbsp;</td></tr><tr><td>0.45 g magnesium sulfate heptahydrate&amp;nbsp;</td><td>Sigma-Aldrich</td><td>M1880</td><td>mL ddH2O</td></tr><tr><td>0.53 g calcium chloride dihydrate&amp;nbsp;</td><td>Mallinckrodt</td><td>4160</td><td>Slowly add calcium soln to the salt/sugar soln with&amp;nbsp;</td></tr><tr><td>0.05 g sodium bicarbonate&amp;nbsp;&amp;nbsp;</td><td>Fisher Scientific</td><td>S233-3</td><td>with constant mixing</td></tr><tr><td>1 g glucose</td><td>MP Biomedicals</td><td>152527</td><td>Adjust to pH 7.2 and filter sterilize using a&amp;nbsp;</td></tr><tr><td>1 g trehalose</td><td>Sigma-Aldrich</td><td>T0167</td><td>0.22 &amp;micro;m disposable bottle-top filter</td></tr><tr><td>10 mL 100X penicillin/streptomycin</td><td>Hyclone</td><td>SV30010</td><td>Add filtered penicillin and streptomycin soln&amp;nbsp;&lt;br/&gt; prior use</td></tr><tr><td>&lt;strong&gt;Incomplete Bge&amp;nbsp; medium (Ibge)&lt;/strong&gt;</td><td></td><td></td><td>&lt;strong&gt;For 900 mL solution&lt;/strong&gt;</td></tr><tr><td>220 mL Schneider&amp;rsquo;s Drosophila medium modified</td><td>Lonza</td><td>04-351Q</td><td>Mix Schneider&#039;s medium with 680 mL ddH2O</td></tr><tr><td>4.5 g lactalbumin enzymatic hydrolysate</td><td>Sigma-Aldrich</td><td>L9010</td><td>Add lactalbumin hydrolysate and galactose</td></tr><tr><td>1.3 g galactose</td><td>Sigma-Aldrich</td><td>G0625</td><td>Adjust to pH 7.2 and filter sterilize using a 0.22 &amp;micro;m&lt;br/&gt; pre-sterilized disposable bottle-top filter</td></tr><tr><td>&lt;strong&gt;Complete Bge medium (cBge)&lt;/strong&gt;</td><td></td><td></td><td>&lt;strong&gt;For 100 mL of solution&lt;/strong&gt;</td></tr><tr><td>90 mL Incomplete Bge medium</td><td></td><td></td><td>To heat-inactivate FBS: Incubate thawed FBS in&amp;nbsp;</td></tr><tr><td>9 mL heat-inact. fetal bovine serum (FBS) (Optima)</td><td>Atlanta Biologicals</td><td>S12450</td><td>waterbath at 60&amp;deg;C for 1 hr while gently&amp;nbsp;</td></tr><tr><td>1 mL 100X penicillin/streptomycin</td><td>Hyclone</td><td>SV30010</td><td>swirling the bottle every 10 min&lt;br/&gt; Aliquot heat-inactivated FBS into 15-mL tubes&amp;nbsp;&lt;br/&gt; and store at -20&amp;deg;C.&amp;nbsp; Mix medium + FBS and&lt;br/&gt; filter sterilize using a 0.22 &amp;micro;m pre-sterilized&amp;nbsp;&lt;br/&gt; disposable bottle-top filter&amp;nbsp;&lt;br/&gt; Add penicillin and streptomycin prior to use</td></tr><tr><td>&lt;strong&gt;Pond water (stock solution)&lt;/strong&gt;</td><td></td><td></td><td>&lt;strong&gt;1L of stock solution&lt;/strong&gt;</td></tr><tr><td>12.5 g calcium carbonate</td><td>Fisher Scientific</td><td>C64-500</td><td>Mix all salts in 1L of ddH2O</td></tr><tr><td>1.25 g magnesium carbonate</td><td>Fisher Scientific</td><td>M27-500</td><td>Note that the salts will not have completely&amp;nbsp;</td></tr><tr><td>1.25 g&amp;nbsp; sodium chloride</td><td>Fisher Scientific</td><td>S271-3</td><td>dissolved.&amp;nbsp; Shake vigorously to suspend&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;</td></tr><tr><td>0.25 g&amp;nbsp; potassium chloride</td><td>Sigma-Aldrich</td><td>P5405</td><td>salts prior to making the working soln&amp;nbsp;&amp;nbsp;</td></tr><tr><td>&lt;strong&gt;Pond water (working solution)&lt;/strong&gt;</td><td></td><td></td><td>&lt;strong&gt;1.5L of solution&lt;/strong&gt;</td></tr><tr><td>0.8 mL stock solution pond water (shake prior use) in&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;</td><td></td><td></td><td>Mix stock to ddH2O</td></tr><tr><td>1500 mL of ddH2O</td><td></td><td></td><td>Sterilize pond water by autoclaving (slow&amp;nbsp;cycle)</td></tr><tr><td>1.5 mL of 100X penicillin/streptomycin&amp;nbsp;</td><td>Hyclone</td><td>SV30010</td><td>Add penicillin and streptomycin prior use</td></tr><tr><td>&lt;strong&gt;Saline solution (1.2% NaCl)&lt;/strong&gt;</td><td></td><td></td><td>&lt;strong&gt;1.5L of solution&lt;/strong&gt;</td></tr><tr><td>18 g sodium chloride in 1500 mL of ddH2O</td><td>Fisher Scientific</td><td>S271-3</td><td>Autoclave saline solution to sterilize</td></tr><tr><td>1.5 mL of 100X penicillin/ streptomycin</td><td>Hyclone</td><td>SV30010</td><td>Add penicillin and streptomycin prior use</td></tr><tr><td>&lt;strong&gt;Additional equipment and material:&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>7-L mouse euthanizing chamber</td><td></td><td></td><td>Following approved IACUC protocol no. V001551</td></tr><tr><td>Mice</td><td>Taconic Biosciences</td><td></td><td>Swiss-Webster, female, 6-wk old, murine&amp;nbsp;&amp;nbsp;</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; tank and regulator</td><td></td><td></td><td>pathogen-free</td></tr><tr><td>24-well tissue culture plate</td><td>TPP</td><td>92424</td><td></td></tr><tr><td>1-L volumetric flasks</td><td></td><td></td><td></td></tr><tr><td>Light source (150W)</td><td>Chiu Tech Corp</td><td>Model F0-150</td><td></td></tr><tr><td>Centrifuge, refrigerated, swinging bucket</td><td>Eppendorf</td><td>Model 5810R</td><td></td></tr><tr><td>Centrifuge bottles (250 mL)</td><td>Nalgene</td><td></td><td></td></tr><tr><td>15-mL centrifuge tubes, sterile</td><td>Corning</td><td>430053</td><td></td></tr><tr><td>Sterile disposable transfer pipets&amp;nbsp;</td><td>Fisher Scientific</td><td>1371120</td><td></td></tr><tr><td>0.22 &amp;micro;m pre-sterilized disposable bottle-top filter&amp;nbsp;</td><td>EMD Millipore</td><td>SCGPS05RE</td><td></td></tr><tr><td>Stainless steel blender</td><td>Waring Commercial</td><td>Model 51BL31</td><td></td></tr><tr><td>Blender cup, 100 mL capacity</td><td>Waring Commercial</td><td></td><td></td></tr><tr><td>Inverted compound microscope</td><td>Nikon Instruments&amp;nbsp;</td><td>Eclipse TE300</td><td></td></tr></tbody></table>

mass isolation and in vitro cultivation of intramolluscan stages of the human blood fluke schistosoma mansoni

Human blood flukes, Schistosoma spp., have a complex life cycle that involves asexual and sexual developmental phases within a snail intermediate and mammalian final host, respectively. The ability to isolate and sustain the different life cycle stages under in vitro culture conditions has greatly facilitated investigations of the cellular, biochemical and molecular mechanisms regulating parasite growth, development and host interactions. Transmission of schistosomiasis requires asexual reproduction and development of multiple larval stages within the snail host; from the infective miracidium, through primary and secondary sporocysts, to the final cercarial stage that is infective to humans. In this paper we present a step-by-step protocol for mass hatching and isolation of Schistosoma mansoni miracidia from eggs obtained from livers of infected mice, and their subsequent introduction into in vitro culture. It is anticipated that the detailed protocol will encourage new researchers to engage in and broaden this important field of schistosome research.

The vast majority of miracidia typically will have been collected in the first two &#34;harvests&#34;. Incubation of isolated miracidia in CBSS+ triggers the miracidium-to-sporocyst transformation process (Figure 4A-C). Within the first hour following transfer to culture wells containing CBSS+, the majority of miracidia cease swimming (Figure 4A). At 6 h in culture, miracidia are in the process of actively sheddi...

Watch this Scientific Journal Video about Mass Isolation and In Vitro Cultivation of Intramolluscan Stages of the Human Blood Fluke Schistosoma Mansoni at JoVE.com

Mass Isolation and In Vitro Cultivation of Intramolluscan Stages of the Human Blood Fluke Schistosoma Mansoni

This article describes a protocol for the large-scale axenic isolation and collection of free-swimming miracidia of the human blood fluke Schistosoma mansoni and their subsequent processing for introduction into in vitro culture.

Mass Isolation and In Vitro Cultivation of Intramolluscan Stages of the Human Blood Fluke Schistosoma Mansoni

Human blood flukes, Schistosoma spp., have a complex life cycle that involves asexual and sexual developmental phases within a snail intermediate and ...

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Research

JoVE Journal

Developmental Biology

9.1K Views.  University of Wisconsin, School of Veterinary Medicine. This article describes a protocol for the large-scale axenic isolation and collection of free-swimming miracidia of the human blood fluke Schistosoma mansoni and their subsequent processing for introduction into in vitro culture. 

Video: Mass Isolation and In Vitro Cultivation of Intramolluscan Stages of the Human Blood Fluke Schistosoma Mansoni

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.

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 ...

Immunology and Infection

Cercarial Transformation and in vitro Cultivation of Schistosoma mansoni Schistosomules

Schistosome parasites are the causative agents of schistosomiasis, a chronically debilitating disease that affects over 200 million people globally and ranks second to malaria among parasitic diseases in terms of public health and socio-economic impact (1-4). Schistosome parasites are trematode worms with a complex life cycle interchanging between a parasitic life in molluscan and mammalian hosts with intervening free-swimming stages. Briefly, free-swimming cercariae infect a mammalian host by penetrating the skin with the aid of secreted proteases, during which time the cercariae lose their tails, transforming into schistosomules. The schistosomules must now evade the host immune system, develop a gut for digestion of red blood cells, and migrate though the lungs and portal circulation en route to their final destination in the hepatic portal system and eventually the mesenteric veins (for S. mansoni) where male and female worms pair and mate, producing hundreds of eggs daily. Some of the eggs are excreted from the body into fresh water, where the eggs hatch into free-swimming miracidia (5-10). The miracidia infect specific snail species and transform into mother and daughter sporocysts, which in turn, produce infective cercariae, completing the life cycle. Unfortunately, the entire schistosome life cycle cannot be cultured in vitro, but infective cercariae can be transformed into schistosomules, and the schistosomules can be cultured for weeks for the analysis of schistosome development in vitro or microarray analysis. In this protocol, we provide a visual description of cercarial transformation and in vitro culturing of schistosomules. We shed infectious cercariae from the snail host Biomphalaria glabrata and manually transform them into schistosomules by detaching their tails using an emulsifying double-ended needle. The in vitro cercarial transformation and schistosomules culture techniques described avoid the use of a mammalian host, which simplifies visualization of schistosomes and facilitates the collection of the parasite for experimental analysis. in vitro transformation and culturing techniques of schistosomes have been done for years (11, 12), but no visual protocols have been developed that are available to the entire community.

Cercarial Transformation and in vitro Cultivation of Schistosoma mansoni Schistosomules

We describe an in vitro method for culturing schistosomula of the flatworm parasite Schistosoma mansoni, via the harvesting and transformation of infective cercariae from the fresh water snail intermediate host Biomphalaria glabrata.

Schistosome parasites are the causative agents of schistosomiasis, a chronically debilitating disease that affects over 200 million people globally and ...

Using Eggs from Schistosoma mansoni as an In vivo Model of Helminth-induced Lung Inflammation

Schistosoma parasites are blood flukes that infect an estimated 200 million people worldwide 1. In chronic infection with Schistosoma, the severe pathology, including liver fibrosis and splenomegaly, is caused by the immune response to the parasite eggs rather than the parasite itself 2. Parasite eggs induce a Th2 response characterized by the production of IL-4, IL-5 and IL-13, the alternative activation of macrophages and the recruitment of eosinophils. Here, we describe injection of Schistosoma mansoni eggs as a model to examine parasite-specific Th2 cytokine responses in the lung and draining lymph nodes, the formation of pulmonary granulomas surrounding the egg, and airway inflammation. 
Following intraperitoneal sensitization and intravenous challenge, S. mansoni eggs are transported to the lung via the pulmonary arteries where they are trapped within the lung parenchyma by granulomas composed of lymphocytes, eosinophils and alternatively activated macrophages 3-6. Associated with granuloma formation, inflammation in the broncho-alveolar spaces, expansion of the draining lymph nodes and CD4 T cell activation can be observed. Here we detail the protocol for isolating Schistosoma mansoni eggs from infected livers (modified from 7), sensitizing and challenging mice, and recovering the organs (broncho-alveolar lavage (BAL), lung and draining lymph nodes) for analysis. We also include representative histologic and immunologic data and suggestions for additional immunologic analysis. 
Overall, this method provides an in vivo model to investigate helminth-induced immunologic responses in the lung, which is broadly applicable to the study of Th2 inflammatory diseases including helminth infection, fibrotic diseases, allergic inflammation and asthma. Advantages of this model for the study of type 2 inflammation in the lung include the reproducibility of a potent Th2 inflammatory response in the lung and draining lymph nodes, the ease of assessment of inflammation by histologic examination of the granulomas surrounding the egg, and the potential for long-term storage of the parasite eggs.

Using Eggs from Schistosoma mansoni as an In vivo Model of Helminth-induced Lung Inflammation

Schistosoma mansoni eggs are potent stimulators of the T helper type 2 (Th2) immune response, characteristic of parasite infection, asthma and allergic inflammation. This protocol utilizes S. mansoni egg injection to generate a CD4 Th2 cytokine-induced inflammatory response in the lung, characterized by lung granuloma formation around the egg, eosinophilia and macrophage alternative activation.

Schistosoma parasites are blood flukes that infect an estimated 200 million people worldwide 1. In chronic infection with Schistosoma, the severe ...

Transient Transduction of the Strobilated Forms of Echinococcus granulosus

Cystic echinococcosis or hydatid disease is one of the most important zoonotic parasitic diseases caused by Echinococcus granulosus, a small tapeworm harbored in the intestine of canines. There is an urgent need for applied genetic research to understand the mechanisms of pathogenesis and disease control and prevention. However, the lack of an effective gene evaluation system impedes direct interpretation of the functional genetics of cestode parasites, including the Echinococcus species. The present study demonstrates the potential of lentiviral gene transient transduction in the metacestode and strobilated forms of E. granulosus. Protoscoleces (PSCs) were isolated from hydatid cysts and transferred to specific biphasic culture media to develop into strobilated worms. The worms were transfected with harvested third-generation lentivirus, along with HEK293T cells as a transduction process control. A pronounced fluorescence was detected in the strobilated worms over 24 h and 48 h, indicating transient lentiviral transduction in E. granulosus. This work presents the first attempt at lentivirus-based transient transduction in tapeworms and demonstrates the promising outcomes with potential implications in experimental studies on flatworm biology.

Transient Transduction of the Strobilated Forms of Echinococcus granulosus

We describe a rapid transient transduction technique in different developmental stages of Echinococcus granulosus using third-generation lentiviral vectors.

Cystic echinococcosis or hydatid disease is one of the most important zoonotic parasitic diseases caused by Echinococcus granulosus, a small tapeworm ...

Genetics

Establishment and Optimization of a High Throughput Setup to Study Staphylococcus epidermidis and Mycobacterium marinum Infection as a Model for Drug Discovery

Zebrafish are becoming a valuable tool in the preclinical phase of drug discovery screenings as a whole animal model with high throughput screening possibilities. They can be used to bridge the gap between cell based assays at earlier stages and in vivo validation in mammalian models, reducing, in this way, the number of compounds passing through to testing on the much more expensive rodent models. In this light, in the present manuscript is described a new high throughput pipeline using zebrafish as in vivo model system for the study of Staphylococcus epidermidis and Mycobacterium marinum infection. This setup allows the generation and analysis of large number of synchronous embryos homogenously infected. Moreover the flexibility of the pipeline allows the user to easily implement other platforms to improve the resolution of the analysis when needed. The combination of the zebrafish together with innovative high throughput technologies opens the field of drug testing and discovery to new possibilities not only because of the strength of using a whole animal model but also because of the large number of transgenic lines available that can be used to decipher the mode of action of new compounds.

Establishment and Optimization of a High Throughput Setup to Study Staphylococcus epidermidis and Mycobacterium marinum Infection as a Model for Drug Discovery

This video article describes the high throughput pipeline that has been successfully established to infect and analyze large numbers of zebrafish embryos providing a new powerful tool for compound testing and drug discovery using a whole animal vertebrate organism.

Zebrafish are becoming a valuable tool in the preclinical phase of drug discovery screenings as a whole animal model with high throughput screening ...

Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites

Malaria infection begins when the sporozoite stage of Plasmodium is inoculated into the skin of a mammalian host through a mosquito bite. The highly motile parasite not only reaches the liver to invade hepatocytes and transform into erythrocyte-infective form. It also migrates into the skin and to the proximal lymph node draining the injection site, where it can be recognized and degraded by resident and/or recruited myeloid cells. Intravital imaging reported the early recruitment of brightly fluorescent Lys-GFP positive leukocytes in the skin and the interactions between sporozoites and CD11c+ cells in the draining lymph node. We present here an efficient procedure to recover, identify and enumerate the myeloid cell subsets that are recruited to the mouse skin and draining lymph node following intradermal injection of immunizing doses of sporozoites in a murine model. Phenotypic characterization using multi-parametric flow cytometry provides a reliable assay to assess early dynamic cellular changes during inflammatory response to Plasmodium infection.

Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites

We describe here a protocol for isolating myeloid cells from mouse skin and draining lymph node following intradermal injection of Plasmodium sporozoites. Flow cytometry of collected cells provides a reliable assay to characterize the skin and draining lymph node inflammatory response to the parasite.

Malaria infection begins when the sporozoite stage of Plasmodium is inoculated into the skin of a mammalian host through a mosquito bite. The highly ...

A Rapid Strategy for the Isolation of New Faustoviruses from Environmental Samples Using Vermamoeba vermiformis

The isolation of giant viruses is of great interest in this new era of virology, especially since these giant viruses are related to protists. Giant viruses may be potentially pathogenic for many species of protists. They belong to the recently described order of Megavirales. The new lineage Faustovirus that has been isolated from sewage samples is distantly related to the mammalian pathogen African swine fever virus. This virus is also specific to its amoebal host, Vermamoeba vermiformis, a protist common in health care water systems. It is crucial to continue isolating new Faustovirus genotypes in order to enlarge its genotype collection and study its pan-genome. We developed new strategies for the isolation of additional strains by improving the use of antibiotic and antifungal combinations in order to avoid bacterial and fungal contaminations of the amoeba co-culture and favoring the virus multiplication. We also implemented a new starvation medium to maintain V. vermiformis in optimal conditions for viruses co-culture. Finally, we used flow cytometry rather than microscopic observation, which is time-consuming, to detect the cytopathogenic effect. We obtained two isolates from sewage samples, proving the efficiency of this method and thus widening the collection of Faustoviruses, to better understand their environment, host specificity and genetic content.

A Rapid Strategy for the Isolation of New Faustoviruses from Environmental Samples Using Vermamoeba vermiformis

We describe here the latest advances in viral isolation for the characterization of new genotypes of Faustovirus, a new asfarvirus-related lineage of giant viruses. This protocol can be applied to the high throughput isolation of viruses, especially giant viruses infecting amoeba.

The isolation of giant viruses is of great interest in this new era of virology, especially since these giant viruses are related to protists. Giant ...

Isolation and Characterization of Extracellular Vesicles from Adult Schistosoma japonicum

Extracellular vesicles (EVs) are membranous vesicles released by a variety of cells into the extracellular microenvironment. EVs represent a population of heterogeneous vesicles, whose size range between 40 and 1,000 nm. Accumulated evidence indicated that EVs play important regulatory roles in pathogen-host interactions. A deep understanding of schistosome EVs should provide insights into the mechanisms underlying schistosome-host interactions, enabling development of novel strategies against schistosomiasis. Here, we aim to further study EVs functions in schistosomes by presenting a protocol for the isolation and characterization of EVs from adult Schistosoma japonicum (S. japonicum). EVs were isolated from in vitro culture medium using centrifugation combined with a commercial exosome isolation kit. The isolated S. japonicum EVs (SjEVs) typically possess a diameter of 100 - 400 nm, and are characterized by transmission electronic microscopy and western blotting. The usage of PKH67 dye-labeled SjEVs has demonstrated that SjEVs are internalized by the recipient cells. Overall, our protocol provides an alternative method for isolating EVs from adult schistosomes; the isolated SjEVs may be suitable for functional analysis.

Isolation and Characterization of Extracellular Vesicles from Adult Schistosoma japonicum

Here, we present a protocol to describe extracellular vesicles (EVs) isolation in in vitro cultured medium from adult Schistosoma japonicum.

Extracellular vesicles (EVs) are membranous vesicles released by a variety of cells into the extracellular microenvironment. EVs represent a population of ...

Isolation And Dendritic Cell-Uptake of Small Extracellular Vesicles from Echinococcus granulosus

The secretion of extracellular vesicles by cestodes is crucial for enabling cellular communication not only among parasites but also with host tissues. In particular, small extracellular vesicles (sEVs) act as nano-carriers transferring natural antigens, which are critical in host immunomodulation and parasite survival. This article presents a step-by-step protocol to isolate sEVs from larval stage cultures of Echinococcus granulosus and analyzes their uptake by dendritic cells obtained from murine bone marrow, which acquire adhesion and antigen presentation capacity during their maturation after one week of in vitro culture. This article provides comprehensive information for generating, purifying, and quantifying sEVs using ultracentrifugation alongside parallel analyses of dynamic light scattering and transmission electron microscopy. Additionally, a detailed experimental protocol is outlined for isolating and cultivating mouse bone marrow cells and driving their differentiation into dendritic cells using Flt3L. These dendritic cells can present antigens to na&#239;ve T cells, thereby modulating the type of immune response in vivo. Thus, alternative protocols, including confocal microscopy and flow cytometry analysis, are proposed to check the acquired maturational phenotype of dendritic cells previously exposed to parasitic sEVs. Finally, it is worth noting that the described protocol can be applied as a whole or in individual parts to carry out parasite in vitro culture, isolate extracellular vesicles, generate bone marrow-derived dendritic cell cultures, and perform uptake assays with these cells.

Here, we describe in vitro culture conditions, isolation, and increased generation of extracellular vesicles (EVs) from Echinococcus granulosus. The small EVs were characterized by dynamic light scattering and transmission electron microscopy. The uptake by bone marrow-derived dendritic cells and their phenotypic modulation were studied using confocal microscopy and flow cytometry.

The secretion of extracellular vesicles by cestodes is crucial for enabling cellular communication not only among parasites but also with host tissues. In ...

A Whole Mount In Situ Hybridization Method for the Gastropod Mollusc Lymnaea stagnalis

Whole mount in situ hybridization (WMISH) is a technique that allows for the spatial resolution of nucleic acid molecules (often mRNAs) within a &#39;whole mount&#39; tissue preparation, or developmental stage (such as an embryo or larva) of interest. WMISH is extremely powerful because it can significantly contribute to the functional characterization of complex metazoan genomes, a challenge that is becoming more of a bottleneck with the deluge of next generation sequence data. Despite the conceptual simplicity of the technique much time is often needed to optimize the various parameters inherent to WMISH experiments for novel model systems; subtle differences in the cellular and biochemical properties between tissue types and developmental stages mean that a single WMISH method may not be appropriate for all situations. We have developed a set of WMISH methods for the re-emerging gastropod model Lymnaea stagnalis that generate consistent and clear WMISH signals for a range of genes, and across all developmental stages. These methods include the assignment of larvae of unknown chronological age to an ontogenetic window, the efficient removal of embryos and larvae from their egg capsules, the application of an appropriate Proteinase-K treatment for each ontogenetic window, and hybridization, post-hybridization and immunodetection steps. These methods provide a foundation from which the resulting signal for a given RNA transcript can be further refined with probe specific adjustments (primarily probe concentration and hybridization temperature).

A Whole Mount In Situ Hybridization Method for the Gastropod Mollusc Lymnaea stagnalis

The goal of this protocol is to provide users with a set of methods for the high-throughput decapsulation of Lymnaea stagnalis embryos and larvae in preparation for whole mount in situ hybridization, and for subsequent pre- and post-hybridization treatments.

Mass Isolation and In Vitro Cultivation of Intramolluscan Stages of the Human Blood Fluke Schistosoma Mansoni

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

Cercarial Transformation and in vitro Cultivation of Schistosoma mansoni Schistosomules

Using Eggs from Schistosoma mansoni as an In vivo Model of Helminth-induced Lung Inflammation

Transient Transduction of the Strobilated Forms of Echinococcus granulosus

Establishment and Optimization of a High Throughput Setup to Study Staphylococcus epidermidis and Mycobacterium marinum Infection as a Model for Drug Discovery

Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites

A Rapid Strategy for the Isolation of New Faustoviruses from Environmental Samples Using Vermamoeba vermiformis

Isolation and Characterization of Extracellular Vesicles from Adult Schistosoma japonicum

Isolation And Dendritic Cell-Uptake of Small Extracellular Vesicles from Echinococcus granulosus

A Whole Mount In Situ Hybridization Method for the Gastropod Mollusc Lymnaea stagnalis