This work was supported by NIH grants R35GM141906 and R21ES031364 and Damon Runyon Fellowship DRG-2371-19.

1. Preparation of seeded NGM plates in the field
<ol>
	<li>Before travel, weigh 23.005 g of Nematode Growth Medium (NGM) (see Table of Materials) powder and pre-pack in a sealable plastic bag. Make one bag for each liter of media desired. 
	NOTE: Pre-packaging before travel bypasses the need for a functional balance in the field.</li>
	<li>Prior to travel, prepare 1 mL of 1M MgSO4, 1 mL of 1M CaCl2, and 25 mL of 1M potassium phosphate buffer for each liter of media desired. To make 1 L of potassium phosphate buffer, dissolve 108.3 g of KH2PO4 and 35.6 g of K2HPO4 in water, as described in WormBook20.</li>
	<li>Before travel, make an overnight culture of OP50 (see Table of Materials) grown in LB at 37 &#176;C, as described in WormBook20. Aliquot the culture into 50 mL conical tubes and wrap the tops with paraffin film to prevent leakage.</li>
	<li>In the field, dissolve the contents of the NGM packet into 973 mL of double-distilled water (ddH20) or the purest, most sterile water available in a 1 L flask or bottle.</li>
	<li>Place the media flask or bottle, with a loose cap or aluminum foil cover, in a boiling hot water bath on a hot plate or stove. Stir occasionally until all the powder is dissolved and is clear (this takes ~30 min). 
	NOTE: If a magnetic hot plate is available, a stir bar is an excellent option to limit the amount of manual stirring.</li>
	<li>Remove the media from the water bath and cool to ~58 &#176;C with occasional shaking or with a stir bar. Once the media is cooled to 58&#176; C, use serological or standard pipettes to add 25 mL of 1M potassium phosphate buffer, 1 mL of 1M MgSO4, and 1 mL of 1M CaCl2, mixing well between each step.</li>
	<li>In the most sterile environment available, pipette or pour the media into plates of the desired size and allow to cool and solidify overnight. Pour one 60 mm plate (~10 mL) for each substrate sample. Pour one 35 mm plate (~3.5 mL) for each isofemale line; the number of small plates needed is difficult to predict in advance.</li>
	<li>Pipette 50 &#181;L of OP50 culture onto each plate and allow it to dry and grow overnight before use.</li>
</ol>
2. Collection of the nematode substrates
<ol>
	<li>Identify a bacteria-rich substrate in the field. Some examples include rotting fruit, flowers, fungi, and stems of herbaceous plants. Soil and leaf litter are also suitable, though they rarely contain Caenorhabditis.</li>
	<li>With a gloved hand, place a sample of this substrate (1-15 cm3) into a sealable plastic bag (see Table of Materials) labeled with a unique sample ID (Figure 1A).</li>
	<li>Record the sample ID, latitude, longitude, date, description of the substrate, and any other local environmental measurements relevant to the experiment, including ambient and substrate temperature, time of collection, condition of substrate, presence of substrate-associated macroinvertebrates, and so on. A smartphone app is available to streamline this process21.</li>
</ol>
3. Preparation of an array of Baermann funnels
<ol>
	<li>For each funnel, use scissors to cut a segment of rubber tubing (see Table of Materials) ~3 cm long.</li>
	<li>Fit the tubing segment over the end of a plastic funnel (see Table of Materials). This may take some effort as the fit is very tight.</li>
	<li>Slide a tubing clamp over the rubber tubing and clamp it shut.</li>
	<li>To make a funnel holder, use a scalpel to cut circular holes of 35 mm diameter in the bottom of a cardboard fly-vial tray (see Table of Materials) that has not been folded together from its flat shipping orientation. A standard tray can accommodate 12 of these holes in a 3 x 4 array.</li>
	<li>Invert the cardboard, fold down the sides once (not twice, as one would to make a fly-vial tray), and tape the sides together to elevate the inverted cardboard tray (Figure 2).</li>
	<li>Place funnels in the holes, first making sure that the tubing clamps are in the closed position.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63287/63287fig02.jpg" /> 
Figure 2: Repurposed cardboard fly vial trays, folded and cut to support 12 Baermann funnels each. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63287/63287fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Transfering of samples into the funnels
<ol>
	<li>Pour water (as sterile as available) into each funnel, filling it about 3 cm below the rim. If air bubbles are trapped in the tubing, tap the funnel to release them.</li>
	<li>With gloved hands, place a lint-free wipe or specifically a Kimwipe (folded in half to make a square) over the funnel, and press down on the center so that it is submerged in the water.</li>
	<li>Manually break large solid pieces of natural substrates (fruit, flower, soil, leaf litter, etc.) into smaller fragments to minimize the distance worms must travel to fall out of the substrate. 
	NOTE: Leaf litter and awkwardly shaped samples can be preprocessed in a food processor or blender.</li>
	<li>Gently place a sample of the natural substrate (1-15 cm3) onto the tissue/lint-free wipe in a funnel without puncturing the tissue and without the sample protruding above the rim.</li>
	<li>Label the funnel, or the cardboard next to the funnel, with the sample ID corresponding to field collection notes.</li>
	<li>Fold the corners of the tissue/lint-free wipe over the sample (Figure 1B). Be careful to keep the sample contained within the tissue/lint-free wipe so that no soil or debris can pass to the bottom of the funnel. 
	NOTE: This step is to prevent the corners from draping over the edge of the funnel, where they would wick the water from the funnel over the sides.</li>
	<li>With hands, a spatula, or a pipette tip, pick out any active insects, millipedes, or other animals that may travel from funnel to funnel, cross-contaminating samples. Wrap the tissue/lint-free wipe entirely around the sample or lay a second tissue across the top of the sample to prevent cross-contamination.</li>
	<li>Add more water to funnels so that the entire sample is submerged (Figure 3).</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63287/63287fig03.jpg" /> 
Figure 3: Assembled Baermann funnels. Each sample is wrapped in tissue/lint-free wipe and submerged under water in the funnel, which is clamped shut. Over a period of ~12 h, the nematodes will migrate through the tissue and to the bottom of the funnel. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63287/63287fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Extraction of nematodes from the funnels
<ol>
	<li>Wait for ~12 h or overnight. During this time, active worms will wriggle out of the substrate, through the tissue/lint-free wipe and down to the bottom of the clamped funnel. 
	NOTE: Waiting much longer than 12 h risks worm mortality due to hypoxia or pathogenic infection, which may also be a risk at shorter durations for samples that are particularly crowded with worms and bacteria.</li>
	<li>Write the sample ID of a funnel on the bottom of a 60 mm NGM worm plate seeded with a spot of OP50 E. coli bacteria. Remove the lid from the plate.</li>
	<li>Remove the funnel containing that sample from the funnel stand. Using one hand to hold the funnel upright above the open worm plate, use the other hand to release pressure on the tubing clamp, allowing one drop of water to fall from the tubing onto the worm plate (Figure 1C). As soon as water drops from the funnel, quickly clamp it shut again to prevent flooding the NGM plate. 
	NOTE: To select worms attracted to OP50, including Caenorhabditis species, release the drop away from the bacterial lawn. When the water soaks into the plate or evaporates, Caenorhabditis nematodes will crawl into the bacterial lawn.</li>
	<li>Clean up: Throw out the contents of the funnels. Wash the funnels with hot water for subsequent reuse.</li>
</ol>
6. Establishing the cultures
<ol>
	<li>Observe the isolated nematodes under the stereomicroscope at a magnification of 5x-50x. The plates should include nematodes, and at much lower frequencies, small oligochaete annelids, tardigrades, rotifers, and small crustaceans&#160;(Figure 4). 
	&#8203;NOTE: If the funnel has been set up correctly, no mites, insects, or visible non-living material will have made it through the funnel.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63287/63287fig04.jpg" /> 
Figure 4: Contents of the first droplet released from a Baermann funnel onto an NGM plate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63287/63287fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>To establish isohermaphrodite or isofemale lines, use a worm pick to transfer each L4 hermaphrodite or mated adult female (recognizable by their larger body size and lack of the distinct male tail22) to a separate 35 mm NGM plate seeded with OP50 (Figure 1D). Use a lighter to sterilize the worm pick before and after transferring worms.</li>
	<li>Use paraffin film to wrap plates thoroughly for travel.</li>
</ol>

<ol>
	<li>Frezal, L., Felix, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=C.+elegans+outside+the+Petri+dish.">C. elegans outside the Petri dish.</a> Elife. 4, 05849 (2015).</li><li>Greene, J. S., 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=Balancing+selection+shapes+density-dependent+foraging+behaviour.">Balancing selection shapes density-dependent foraging behaviour.</a> Nature. 539 (7628), 254-258 (2016).</li><li>Reddy, K. 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=Antagonistic+paralogs+control+a+switch+between+growth+and+pathogen+resistance+in+C.+elegans.">Antagonistic paralogs control a switch between growth and pathogen resistance in C. elegans.</a> PLOS Pathogens. 15 (1), 1007528 (2019).</li><li>Schulenburg, H., Felix, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+natural+biotic+environment+of+Caenorhabditis+elegans.">The natural biotic environment of Caenorhabditis elegans.</a> Genetics. 206 (1), 55-86 (2017).</li><li>Zhang, F., 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=Caenorhabditis+elegans+as+a+model+for+microbiome+research.">Caenorhabditis elegans as a model for microbiome research.</a> Frontiers in Microbiology. 8, 485 (2017).</li><li>Andersen, E. 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=Chromosome-scale+selective+sweeps+shape+Caenorhabditis+elegans+genomic+diversity.">Chromosome-scale selective sweeps shape Caenorhabditis elegans genomic diversity.</a> Nature Genetics. 44 (3), 285-290 (2012).</li><li>Cook, D. E., Zdraljevic, S., Roberts, J. P., Andersen, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CeNDR,+the+Caenorhabditis+elegans+natural+diversity+resource.">CeNDR, the Caenorhabditis elegans natural diversity resource.</a> Nucleic Acids Research. 45, 650-657 (2017).</li><li>Crombie, T. 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=Deep+sampling+of+Hawaiian+Caenorhabditis+elegans+reveals+high+genetic+diversity+and+admixture+with+global+populations.">Deep sampling of Hawaiian Caenorhabditis elegans reveals high genetic diversity and admixture with global populations.</a> Elife. 8, 50465 (2019).</li><li>Lee, D., 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=Balancing+selection+maintains+hyper-divergent+haplotypes+in+Caenorhabditis+elegans.">Balancing selection maintains hyper-divergent haplotypes in Caenorhabditis elegans.</a> Nature Ecology &amp; Evolution. 5 (6), 794-807 (2021).</li><li>Rockman, M. V., Kruglyak, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recombinational+landscape+and+population+genomics+of+Caenorhabditis+elegans.">Recombinational landscape and population genomics of Caenorhabditis elegans.</a> PLOS Genetics. 5 (3), 1000419 (2009).</li><li>Evans, K. S., van Wijk, M. H., McGrath, P. T., Andersen, E. C., Sterken, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+QTL+to+gene:+C.+elegans+facilitates+discoveries+of+the+genetic+mechanisms+underlying+natural+variation.">From QTL to gene: C. elegans facilitates discoveries of the genetic mechanisms underlying natural variation.</a> Trends in Genetics. 37, 933-947 (2021).</li><li>Gaertner, B. E., Phillips, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Caenorhabditis+elegans+as+a+platform+for+molecular+quantitative+genetics+and+the+systems+biology+of+natural+variation.">Caenorhabditis elegans as a platform for molecular quantitative genetics and the systems biology of natural variation.</a> Genetics Research (Cambridge Core). 92 (5-6), 331-348 (2010).</li><li>Noble, L. M., Rockman, M. V., Teotonio, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene-level+quantitative+trait+mapping+in+Caenorhabditis+elegans.">Gene-level quantitative trait mapping in Caenorhabditis elegans.</a> G3 Genes|Genomes|Genetics. 11 (2), 061 (2021).</li><li>Van Bezooijen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods and techniques for nematology. , (2006).</li><li>Barri&#232;rre, A., F&#233;lix, M. -. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+C.+elegans.+and+related+nematodes.">Isolation of C. elegans. and related nematodes.</a> Wormbook. , (2014).</li><li>Baermann, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eine+einfache+Methode+zur+Auffindung+von+Anklostomum+(Nematoden)+Larven+in+Erdproben.">Eine einfache Methode zur Auffindung von Anklostomum (Nematoden) Larven in Erdproben.</a> Geneeskundig tijdschrift voor Nederlandsch-Indi&#235;. 57, 131-137 (1917).</li><li>Gray, N. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ecology+of+nematophagous+fungi+Panagrellus+redivivus+as+the+target+organism.">Ecology of nematophagous fungi Panagrellus redivivus as the target organism.</a> Plant and Soil. 297, 293-297 (1983).</li><li>Mallez, S., Castagnone, C., Espada, M., Viera, P., Eisenback, J., Mota, M., Guillemaud, T., Castagnone-Sereno, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=First+insights+into+the+genetic+diversity+of+the+pinewood+nematode+in+its+native+area+using+new+polymorphic+microsatellite+loci.">First insights into the genetic diversity of the pinewood nematode in its native area using new polymorphic microsatellite loci.</a> PLOS ONE. 8 (3), 4-11 (2013).</li><li>Kerfahi, D., Tripathi, M. B., Porazinska, L. D., Park, J., Go, R., Adams, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Do+tropical+rain+forest+soils+have+greater+nematode+diversity+than+High+Arctic+tundra?+A+metagenetic+comparison+of+Malaysia+and+Svalbard.">Do tropical rain forest soils have greater nematode diversity than High Arctic tundra? A metagenetic comparison of Malaysia and Svalbard.</a> Global Ecology and Biogeography. 25, 716-728 (2016).</li><li>Stiernagle, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maintenance+of+C.+elegans.">Maintenance of C. elegans.</a> WormBook. , 1-11 (2006).</li><li>Di Bernardo, M., Crombie, T. A., Cook, D. E., Andersen, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=easyFulcrum:+An+R+package+to+process+and+analyze+ecological+sampling+data+generated+using+the+Fulcrum+mobile+application.">easyFulcrum: An R package to process and analyze ecological sampling data generated using the Fulcrum mobile application.</a> PLOS ONE. 16, 0254293 (2021).</li><li>Lints, R., Hall, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Handbook+of+C.+elegans+male+anatomy.">Handbook of C. elegans male anatomy.</a> WormAtlas. , (2005).</li><li>Kiontke, K., F&#233;lix, M. -. A., Ailion, M., Rockman, M. V., Braendle, C., P&#233;nigault, J. -. B., Fitch, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+phylogeny+and+molecular+barcodes+for+Caenorhabditis,+with+numerous+new+species+from+rotting+fruits.">A phylogeny and molecular barcodes for Caenorhabditis, with numerous new species from rotting fruits.</a> BMC Evolutionary Biology. 11, 339 (2011).</li></ol>

The authors declare no conflicts of interest.

The central principle of this method is that nematodes will pass through the tissue submerged in water, while their substrate and larger invertebrate contaminants will not. The critical steps of the protocol are (1) collecting an appropriate substrate, (2) submerging the substrate, wrapped in a filtering material, in water, (3) collecting worms that have passed through the filter and sunk to the bottom of the water, and (4) isolating individual worms to create isofemale or isohermaphrodite lines. All other parts of the method are amenable to modification as necessary according to the resources available, the nature of the substrates, or the fieldwork goals. Some modifications worth considering are as follows.
Baiting the worms by planting fruit 
If there is not an ample supply of bacteria-rich rotting material to be found in the field site, one may wish to bring a sample, such as a piece of apple or tomato, to leave to rot. Pin the bait down well with a few stakes so that larger animals do not remove them and so it can easily be found later for collection. Avoid direct sunlight, where the bait might dry before it rots.
Constructing funnel apparatus out of whatever materials are available 
Any structure with holes large enough to accommodate the tubing clamp and stable enough to support a top-heavy funnel will work. For a single funnel, a drinking glass is a suitable holder. Any kind of tissue&#160;&#8211;&#160;facial tissue, toilet paper, or paper towels &#8211;&#160;can be used.
Putting less material in funnels, or adjusting the wait time, to prevent hypoxia or infection 
If the sample has a very high concentration of worms or bacteria, nematodes may begin dying from hypoxia or infection before the 12 h incubation is complete. If this is a concern, the researcher may check funnels earlier or prepare an additional funnel with a very small subsample of the highly populated substrate.
Adjusting the NGM plate preparation to experimental needs and constraints 
NGM plates can be prepared with whatever media and food source is appropriate to the experiment. The field protocol described above is designed to minimize baggage weight. Depending on baggage limitations and the fieldwork timing, bringing already-poured NGM plates &#8211;&#160;either prepared in the lab or purchased commercially &#8211;&#160;may be preferable rather than pouring plates in the field.
Performing the funnel isolations in the laboratory 
The protocol describes carrying out the complete procedure under field conditions to capture the nematode population as it occurs in nature. For some research goals, it may be sufficient to isolate nematodes later, after traveling back to the lab with samples in sealed bags. Even then, the Baermann funnel method provides a cleaner and complete sample of the surviving nematodes than other isolation methods. However, samples in sealed bags may experience selection during travel, as they are exposed to potential extremes of temperature and hypoxia. This can be minimized by performing isolations as soon as possible after sample collection.
A common alternative method to the Baermann funnel involves placing the substrate directly onto an NGM plate and waiting for worms to crawl out, which is either highly labor-intensive or results in the incomplete collection of the population. It also yields plates contaminated with mites and insect larvae. The Baermann funnel method is a low-cost, low-tech, low-labor strategy for quickly separating the entire population of active worms from their substrate.
The Baermann funnel method is not universally applicable for collecting all types of wild nematodes. Some plant-dwelling nematodes take much longer than 12 h to emerge from their substrate and will be absent from a droplet released too early, while some insect parasites will crawl to the top of the funnel rather than the bottom, also evading collection14. Alternatives to the Baermann funnel either require more specialized equipment or more labor to recover the worms. However, they may still be preferred if the caveats above are a problem for the experiment. Alternative options, reviewed by van Bezooijen14, include the funnel spray method, which provides a constant mist of water to funnels, adding oxygen and allowing the overflow of bacteria in suspension. This allows for a more extended extraction period of nematodes from plants. The blender centrifugal flotation method recovers slow-moving, inactive, or upward-crawling nematodes by separating them by their specific gravity, the Oostenbrink elutriator applies an undercurrent to separate settling sediment from suspended nematodes, and Cobb&#39;s Method uses a series of sieves to isolate nematodes by their size, shape, and sedimentation rate14. To collect rhabditids, though, the Baermann funnel effectively produces clean samples quickly and with minimal effort.

Valuable biological insights are emerging as researchers who study&#160;C. elegans in the lab expand their focus to C. elegans and related rhabditid nematodes in the wild. Studies of wild nematodes place genes and genomes in their natural context, revealing functions potentially obscured by laboratory conditions1,2,3,4,5. These studies generate insights into the prerequisite for evolution itself, genetic variation6,7,8,9,10. The natural genetic variation captured by wild samples also provides inroads into the genetic basis of many complex traits11,12,13.
When designing studies that require the isolation of natural populations of nematodes, particularly&#160;when performing remote fieldwork, practical considerations come to the fore. This protocol aims to cleanly isolate entire populations of active nematodes that can be cultured on OP50 from bait or wild substrates. The method is well suited for extracting free-living rhabditid and diplogasterid nematodes, including Caenorhabditis, Oscheius, and Pristionchus.
There are many techniques for isolating nematodes from their substrates14,15. The most basic approach is to place the substrate directly onto a nematode-medium plate, picking animals as they crawl out8,15. This method requires large amounts of time and labor if the goal is to isolate all nematodes from a sample. More sophisticated techniques take advantage of the animals&#39; weight, size, mobility, or some combination of these14. Each method has its advantages and disadvantages in terms of setup and throughput. They also differ in their sampling biases and may select for certain nematodes if the animals in the sample vary along the axis of the method&#39;s separation principle.
The Baermann funnel method was first described in 1917 by Dutch Physician G. K. T. F. Baermann, who invented the device on Java while studying soil-dwelling nematodes, including the parasitic hookworm16. The Baermann funnel functions based on the principle of mobility. The substrate is placed in a funnel lined with a cloth or paper filter (a &#34;Kimwipe&#34; is used for this study, referred to as &#34;lint-free wipe&#34; in the current protocol) and sealed shut at the bottom. The funnel is then filled with water, submerging the sample while the filter separates it from the sealed outlet. Active nematodes in the sample release themselves into the water and swim through the filter, eventually settling at the bottom of the funnel. The funnel outlet is opened, and a drop of nematodes is expelled onto a plate (Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63287/63287fig01.jpg" /> 
Figure 1: Summary of the Baermann funnel technique. (A) Collecting&#160;a bacteria-rich sample from a site of interest. (B) Submerging the sample in a Baermann funnel and waiting for worms to wriggle out and sink. (C) Releasing a single drop from the funnel. (D) Moving single hermaphrodites or mated females to separate plates. Illustration created by Ramin Rahni. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63287/63287fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The Baermann funnel will not work for every kind of nematode (see Discussion section for specific alternatives) and is best suited to those that are active forms in the size range of Caenorhabditis or smaller14. However, if a study can use the Baermann funnel, there are many advantages. The method is practical in the field, requiring limited setup, hands-on time, and cost. The researcher is left with a clean sample without the obstruction of the substrate on the plate, which makes picking easy. Using a filter also prevents contamination of the plate by insect larvae or mites, which chew up plates or prey on nematodes in the sample. Most importantly, the Baermann funnel efficiently extracts nearly the whole population from the substrate14, which may be required depending on the study&#39;s design. For example, researchers interested in counting wild populations&#39; stage or sex distribution, finding slow-growing or rare genotypes, or sampling nematodes not attracted to OP50 might benefit from this method. This is appropriate for researchers studying ecological17, population genetic18, or metagenetic19 questions as the sampling scheme takes a snapshot of the population at the sampling time.
The present manuscript describes a complete protocol for isolating populations of nematodes using the Baermann funnel and establishing isofemale and isohermaphrodite lines in the field, using equipment chosen for easy transport. For researchers conducting fieldwork near their labs, many of these steps can be omitted or simplified.

<table><tbody><tr><td>1 L glass bottle</td><td>NA</td><td>NA</td><td>Step 1 - vessel for making plate media</td></tr><tr><td>1 mL pipette tips</td><td>any</td><td>NA</td><td>Step 1 - dispense worm media additives and seed plates</td></tr><tr><td>1 mL pipetter</td><td>any</td><td>NA</td><td>Step 1 - dispense worm media additives and seed plates</td></tr><tr><td>35 mm worm plates</td><td>Tritech</td><td>T3500</td><td>Step 1 - worm plates</td></tr><tr><td>4mm ID rubber tubing</td><td>Fisher</td><td>14178A</td><td>Step 3 - part of the funnel</td></tr><tr><td>60 mm worm plates</td><td>Greiner</td><td>628161</td><td>Step 1 - worm plates</td></tr><tr><td>65 mm Funnel</td><td>Fisher</td><td>22170156</td><td>Step 3 - part of the funnel</td></tr><tr><td>Calcium chloride</td><td>Fisher</td><td>C79-500</td><td>Step 1 - worm plate medium</td></tr><tr><td>Cooking pot</td><td>NA</td><td>NA</td><td>Step 1 - a water bath for melting the agar to pour plates</td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; strain OP50</td><td>Caenorhabditis Genetics Center</td><td>OP50</td><td>Step 1 - worm food</td></tr><tr><td>Field microscope</td><td>OMAX</td><td>G223CS</td><td>Step 6 - for viewing worms in the field</td></tr><tr><td>Fly vial storage tray</td><td>Azer Scientific</td><td>ES-260T</td><td>Step 3 - to build a platform to hold 12 funnels</td></tr><tr><td>Hot plate or stove</td><td>NA</td><td>NA</td><td>Step 1 - to heat the water bath</td></tr><tr><td>Kimwipes</td><td>KimberlyClark</td><td>34120</td><td>Step 4 - filter for the funnels</td></tr><tr><td>LB media</td><td>Gibco</td><td>10855021</td><td>Step 1 - for growing OP50</td></tr><tr><td>Lighter</td><td>NA</td><td>NA</td><td>Step 6 - sterilize the worm pick</td></tr><tr><td>Magnesium sulfate</td><td>Fisher</td><td>M63-500</td><td>Step 1 - worm plate medium</td></tr><tr><td>NGM-lite agar</td><td>USBiological</td><td>N1000</td><td>Step 1 - worm plate medium</td></tr><tr><td>Parafilm M wrapping film</td><td>Fisher</td><td>13-374-10</td><td>Step 6 - pack worm plates for travel</td></tr><tr><td>Potassium phosphate dibasic</td><td>Fisher</td><td>BP363-500</td><td>Step 1 - worm plate medium</td></tr><tr><td>Potassium phosphate monobasic</td><td>Fisher</td><td>P285-3</td><td>Step 1 - worm plate medium</td></tr><tr><td>scalpel</td><td>NA</td><td>NA</td><td>Step 3 - cut holes for the funnels</td></tr><tr><td>scissors</td><td>NA</td><td>NA</td><td>Step 3 - cut the rubber tubing</td></tr><tr><td>Tape</td><td>NA</td><td>NA</td><td>Step 3 - tape the funnel holder together</td></tr><tr><td>Tubing clamps</td><td>Fisher</td><td>5869</td><td>Step 3 - part of the funnel</td></tr><tr><td>Worm pick</td><td>NA</td><td>NA</td><td>Step 6 - for isolating individual worms</td></tr><tr><td>Ziplock-style storage bags</td><td>NA</td><td>NA</td><td>Step 2 - to bag a substrate sample</td></tr></tbody></table>

rapid isolation of wild nematodes by baermann funnel

Beyond being robust experimental model organisms, Caenorhabditis elegans and its relatives are also real animals that live in nature. Studies of wild nematodes in their natural environments are valuable for understanding many aspects of biology, including the selective regimes in which distinctive genomic and phenotypic characters evolve, the genetic basis for complex trait variation, and the natural genetic diversity fundamental to all animal populations. This manuscript describes a simple and efficient method for extracting nematodes from their natural substrates, including rotting fruits, flowers, fungi, leaf litter, and soil. The Baermann funnel method, a classical nematology technique, selectively isolates active nematodes from their substrates. Because it recovers nearly all active worms from the sample, the Baermann funnel technique allows for the recovery of rare and slow-growing genotypes that co-occur with abundant and fast-growing genotypes, which might be missed in extraction methods that involve multiple generations of reproduction. The technique is also well suited to addressing metagenetic, population-genetic, and ecological questions. It captures the entire population in a sample simultaneously, allowing an unbiased view of the natural distribution of ages, sexes, and genotypes. The protocol allows for deployment at scale in the field, rapidly converting substrates into worm plates, and the authors have validated it through fieldwork on multiple continents.

This protocol was used to isolate nematodes from fruit, flowers, fungi, soil, and stems on Barro Colorado Island, Panam&#225;, at the Smithsonian Tropical Research Institute field station in August of 2018. Of 131 substrates processed by a single investigator over four days, 130 substrates (99.2%) yielded nematodes. Forty-four of the substrates (33.6%) yielded Caenorhabditis nematodes (Figure 5). Subsequent analysis of cultures established from these forty-four substrates, by PCR and mating tests23, revealed the presence of six different Caenorhabditis species &#8211;&#160;C. becei, C. tropicalis, C. briggsae, C. sp. 24, C. sp. 57, and C. panamensis.
This protocol was used again to isolate nematodes from various substrates in the Chernobyl Exclusion Zone, Ukraine, over four days in August of 2019. Live worms were recovered from 62 out of 63 soil samples, 1 out of 17 invertebrate samples, 31 out of 75 fruit samples, 1 out of 12 bait samples (see Discussion section), and no worms were recovered from mushroom, river reed, or wolf feces samples (one sample collected of each). Subsequent sequencing of 18S ribosomal DNA15 identified these nematodes as Oscheius, Panagrolaimus, Acrobeloides, Mesorhabditis, Panagrellus, Pristionchus, and Pelodera, but no Caenorhabditis were identified (Figure 5).
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63287/63287fig05.jpg" /> 
Figure 5: Success rates of two collection trips. Panama in 2018 (left) and Ukraine in 2019 (right). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63287/63287fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Rapid Isolation of Wild Nematodes by Baermann Funnel at JoVE.com

Rapid Isolation of Wild Nematodes by Baermann Funnel

This protocol outlines a method for efficiently extracting live nematodes from natural substrates in the field.

Beyond being robust experimental model organisms, Caenorhabditis elegans and its relatives are also real animals that live in nature. Studies of wild ...

rapid-isolation-of-wild-nematodes-by-baermann-funnel

Nanyang Technological University

Research

JoVE Journal

Genetics

7.9K Views.  New York University. This protocol outlines a method for efficiently extracting live nematodes from natural substrates in the field.

Video: Rapid Isolation of Wild Nematodes by Baermann Funnel

Helminth Collection and Identification from Wildlife

Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are &#34;roundworms&#34; (nematodes), &#34;thorny-headed worms&#34; (acanthocephalans), &#34;flukes&#34; (trematodes), and &#34;tapeworms&#34; (cestodes). The optimum method for collecting helminths is to examine a host that has been dead less than 4-6 hr since most helminths will still be alive. A thorough necropsy should be conducted and all major organs examined. Organs are washed over a 106 &#956;m sieve under running water and contents examined under a stereo microscope. All helminths are counted and a representative number are fixed (either in 70% ethanol, 10% buffered formalin, or alcohol-formalin-acetic acid). For species identification, helminths are either cleared in lactophenol (nematodes and small acanthocephalans) or stained (trematodes, cestodes, and large acanthocephalans) using Harris&#39; hematoxylin or Semichon&#39;s carmine. Helminths are keyed to species by examining different structures (e.g. male spicules in nematodes or the rostellum in cestodes). The protocols outlined here can be applied to any vertebrate animal. They require some expertise on recognizing the different organs and being able to differentiate helminths from other tissue debris or gut contents. Collection, preservation, and staining are straightforward techniques that require minimal equipment and reagents. Taxonomic identification, especially to species, can be very time consuming and might require the submission of specimens to an expert or DNA analysis.

Wild animals are commonly parasitized by a wide range of helminths.&#160; The four major types of helminths are &#8220;roundworms&#8221; (nematodes), &#8220;thorny-headed worms&#8221; (acanthocephalans), &#8220;flukes&#8221; (trematodes), and &#8220;tapeworms&#8221; (cestodes).&#160; Here we describe how helminths are collected from a vertebrate animal and how they are preserved and taxonomically identified.&#160;

Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are &#34;roundworms&#34; (nematodes), ...

Environment

Soil Sampling and Isolation of Entomopathogenic Nematodes (Steinernematidae, Heterorhabditidae)

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to two families: Steinernematidae and Heterorhabditidae. Until now, more than 70 species have been described in the Steinernematidae and there are about 20 species in the Heterorhabditidae. The nematodes have a mutualistic partnership with Enterobacteriaceae bacteria and together they act as a potent insecticidal complex that kills a wide range of insect species.
Herein, we focus on the most common techniques considered for collecting EPN from soil. The second part of this presentation focuses on the insect-baiting technique, a widely used approach for the isolation of EPN from soil samples, and the modified White trap technique which is used for the recovery of these nematodes from infected insects. These methods and techniques are key steps for the successful establishment of EPN cultures in the laboratory and also form the basis for other bioassays that consider these nematodes as model organisms for research in other biological disciplines. The techniques shown in this presentation correspond to those performed and/or designed by members of S. P. Stock laboratory as well as those described by various authors.

Soil Sampling and Isolation of Entomopathogenic Nematodes Steinernematidae, Heterorhabditidae

Entomopathogenic nematodes are soil-inhabiting roundworms that parasitize a wide range of insects. We demonstrate sampling methods used for the isolation of these nematodes from the soil using two techniques: the insect baiting and the modified White trap, for the recovery of nematodes from soil samples and infected insect cadavers, respectively.

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to ...

In vivo and In vitro Rearing of Entomopathogenic Nematodes (Steinernematidae and Heterorhabditidae)

Entomopathogenic nematodes (EPN) (Steinernematidae and Heterorhabditidae) have a mutualistic partnership with Gram-negative Gamma-Proteobacteria in the family Enterobacteriaceae. Xenorhabdus bacteria are associated with steinernematids nematodes while Photorhabdus are symbionts of heterorhabditids. Together nematodes and bacteria form a potent insecticidal complex that kills a wide range of insect species in an intimate and specific partnership. Herein, we demonstrate in vivo and in vitro techniques commonly used in the rearing of these nematodes under laboratory conditions. Furthermore, these techniques represent key steps for the successful establishment of EPN cultures and also form the basis for other bioassays that utilize these organisms for research. The production of aposymbiotic (symbiont&#8211;free) nematodes is often critical for an in-depth and multifaceted approach to the study of symbiosis. This protocol does not require the addition of antibiotics and can be accomplished in a short amount of time with standard laboratory equipment. Nematodes produced in this manner are relatively robust, although their survivorship in storage may vary depending on the species used. The techniques detailed in this presentation correspond to those described by various authors and refined by P. Stock&#8217;s Laboratory, University of Arizona (Tucson, AZ, USA). These techniques are distinct from the body of techniques that are used in the mass production of these organisms for pest management purposes.

In vivo and In vitro Rearing of Entomopathogenic Nematodes Steinernematidae and Heterorhabditidae

The goal of this presentation is to demonstrate in vivo and in vitro techniques for the rearing of entomopathogenic nematodes. In vivo methods consider the rearing of these nematodes with an insect host, whereas the in vitro methods utilize rich agar media.

Entomopathogenic nematodes (EPN) (Steinernematidae and Heterorhabditidae) have a mutualistic partnership with Gram-negative Gamma-Proteobacteria in the ...

Bioengineering

Cultivation of Heligmosomoides Polygyrus: An Immunomodulatory Nematode Parasite and its Secreted Products

Heligmosomoides polygyrus (formerly known as Nematospiroides dubius, and also referred to by some as H. bakeri) is a gastrointestinal helminth that employs multiple immunomodulatory mechanisms to establish chronic infection in mice and closely resembles prevalent human helminth infections. H. polygyrus has been studied extensively in the field of helminth-derived immune regulation and has been found to potently suppress experimental models of allergy and autoimmunity (both with active infection and isolated secreted products). The protocol described in this paper outlines management of the H. polygyrus life cycle for consistent production of L3 larvae, recovery of adult parasites, and collection of their excretory-secretory products (HES).

Cultivation of Heligmosomoides Polygyrus: An Immunomodulatory Nematode Parasite and its Secreted Products

Heligmosomoides polygyrus is a murine nematode with powerful immunomodulatory capabilities that closely resemble those of highly-prevalent human helminth infection. Here we describe a protocol for the long-term maintenance of the H. polygyrus lifecycle.

Heligmosomoides polygyrus (formerly known as Nematospiroides dubius, and also referred to by some as H. bakeri) is a gastrointestinal helminth that ...

Immunology and Infection

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Biology

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Caenorhabditis elegans (C. elegans)&#160;has proven to be an excellent model for studying host-microbe interactions and the microbiome, especially in the context of the intestines. Recently, ecological sampling of wild Caenorhabditis nematodes has discovered a diverse array of associated microbes, including bacteria, viruses, fungi, and microsporidia. Many of these microbes have interesting colonization or infection phenotypes that warrant further study, but they are often unculturable. This protocol presents a method to enrich the desired intestinal microbes in C. elegans and related nematodes and reduce the presence of the many contaminating microbes adhering to the cuticle. This protocol involves forcing animals into the dauer stage of development and using a series of antibiotic and detergent washes to remove external contamination. As dauer animals have physiological changes that protect nematodes from harsh environmental conditions, any intestinal microbes will be protected from these conditions. But, for enrichment to work, the microbe of interest must be maintained when animals develop into dauers. When the animals leave the dauer stage, they are singly propagated into individual lines. F1 populations are then selected for desired microbes or infection phenotypes and against visible contamination. These methods will allow researchers to enrich unculturable microbes in the intestinal lumen, which make up part of the natural microbiome of C. elegans and intracellular intestinal pathogens. These microbes can then be studied for colonization or infection phenotypes and their effects on the host fitness.

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Wild Caenorhabditis nematodes are associated with many microbes, often in the gut lumen or infecting the intestine. This protocol details a method to enrich unculturable microbes colonizing the intestine, taking advantage of the resistance of the dauer cuticle.

Caenorhabditis elegans (C. elegans)&#160;has proven to be an excellent model for studying host-microbe interactions and the microbiome, especially in the ...

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

The genus Strongyloides consists of multiple species of skin-penetrating nematodes with different host ranges, including Strongyloides stercoralis and Strongyloides ratti. S.&#160;stercoralis is a human-parasitic, skin-penetrating nematode that infects approximately 610 million people, while the rat parasite S.&#160;ratti is closely related to S.&#160;stercoralis and is often used as a laboratory model for S.&#160;stercoralis. Both S.&#160;stercoralis and S.&#160;ratti are easily amenable to the generation of transgenics and knockouts through the exogenous nucleic acid delivery technique of intragonadal microinjection, and as such, have emerged as model systems for other parasitic helminths that are not yet amenable to this technique.
Parasitic Strongyloides adults inhabit the small intestine of their host and release progeny into the environment via the feces. Once in the environment, the larvae develop into free-living adults, which live in feces and produce progeny that must find and invade a new host. This environmental generation is unique to the Strongyloides species and similar enough in morphology to the model free-living nematode Caenorhabditis elegans that techniques developed for C. elegans can be adapted for use with these parasitic nematodes, including intragonadal microinjection. Using intragonadal microinjection, a wide variety of transgenes can be introduced into Strongyloides. CRISPR/Cas9 components can also be microinjected to create mutant Strongyloides larvae. Here, the technique of intragonadal microinjection into Strongyloides, including the preparation of free-living adults, the injection procedure, and the selection of transgenic progeny, is described. Images of transgenic Strongyloides larvae created using CRISPR/Cas9 mutagenesis are included. The aim of this paper is to enable other researchers to use microinjection to create transgenic and mutant Strongyloides.

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

The functional genomic toolkit for the parasitic nematodes Strongyloides stercoralis and Strongyloides ratti includes transgenesis, CRISPR/Cas9-mediated mutagenesis, and RNAi. This protocol will demonstrate how to use intragonadal microinjection to introduce transgenes and CRISPR components into S.&#160;stercoralis and S.&#160;ratti.

The genus Strongyloides consists of multiple species of skin-penetrating nematodes with different host ranges, including Strongyloides stercoralis and ...

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. To improve the mating efficiency of B. Xylophilus and to increase the chances of mating observation, virgin adults were cultured and used for the investigation. Eggs were obtained by keeping the nematodes in water and allowing the females to lay eggs for 10 min. The second-stage juveniles (J2) were synchronized by incubating the eggs for 24 h at 25 &#176;C in the dark, and the early J4 were obtained by culturing the J2 with grey mold, Botrytis cinerea, for another 52 h. At this time point, most J4 nematodes could be clearly distinguished as being male or female using their genital morphology. The male and female J4 were collected and cultured separately in two different Petri dishes for 24 h to get virgin adult nematodes. A virgin male and a virgin female were paired in a drop of water in the well of a concave slide. The mating behavior was filmed with a video recorder under a stereomicroscope. The whole period of the mating process was 82.8 &#177;3.91 min (mean &#177;SE) and could be divided into 4 different phases: searching, contacting, copulating, and lingering. The mean minutes of duration were 21.8 &#177; 2.0, 28.0 &#177; 1.9, 25.8 &#177; 0.7 and 7.2 &#177; 0.5, respectively. Eleven sub-behaviors were described: cruising, approaching, encountering, touching, hooping, locating, attaching, ejaculating, separating, quiescence, and roaming. Interestingly, obvious intra-sexual competition was observed when one female was grouped with 3 males or one male with 3 females. This protocol is useful and valuable, not only in investigating the mating behavior of B. xylophilus, but also in acting as a reference for ethological studies of other nematodes.

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A protocol for investigating the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, is presented. Behavioral features of B. xylophilus are described in the mating process.

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. ...

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

Caenorhabditis elegans is one of the major model organisms in biology, but only recently have researchers focused on its natural ecology. The relative sparsity of information about C. elegans in its natural context comes from the challenges involved in the identification of the small nematode in nature. Despite these challenges, an increasing focus on the ecology of C. elegans has caused a wealth of new information regarding its life outside of the laboratory. The intensified search for C. elegans in nature has contributed to the discovery of many new Caenorhabditis species and revealed that congeneric nematodes frequently cohabitate in the wild, where they feed on microbial blooms associated with rotting plant material. The identification of new species has also revealed that the androdioecious mating system of males and self-fertilizing hermaphrodites has evolved three times independently within Caenorhabditis. The other two selfing species, C. briggsae and C. tropicalis, share the experimental advantages of C. elegans and have enabled comparative studies into the mechanistic basis of important traits, including self-fertilization. Despite these advances, much remains to be learned about the ecology and natural diversity of these important species. For example, we still lack functional information for many of their genes, which might only be attained through an understanding of their natural ecology. To facilitate ecological research of selfing Caenorhabditis nematodes, we developed a highly scalable method to collect nematodes from the wild. Our method makes use of mobile data collection platforms, cloud-based databases, and the R software environment to enhance researchers&#39; ability to collect nematodes from the wild, record associated ecological data, and identify wild nematodes using molecular barcodes.

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

This protocol can be used to perform large-scale ecological surveys of selfing Caenorhabditis nematodes. The primary advantage of this method is the efficient organization and analysis of ecological and molecular data associated with the nematodes collected from nature.

Caenorhabditis elegans is one of the major model organisms in biology, but only recently have researchers focused on its natural ecology. The relative ...

Isolation and Characterization of the Natural Microbiota of the Model Nematode Caenorhabditis elegans

The nematode Caenorhabditis elegans interacts with a large diversity of microorganisms in nature. In general, C. elegans is commonly found in rotten plant matter, especially rotten fruits like apples or on compost heaps. It is also associated with certain invertebrate hosts such as slugs and woodlice. These habitats are rich in microbes, which serve as food for C. elegans and which can also persistently colonize the nematode gut. To date, the exact diversity and consistency of the native C. elegans microbiota across habitats and geographic locations is not fully understood. Here, we describe a suitable approach for isolating C. elegans from nature and characterizing the microbiota of worms. Nematodes can be easily isolated from compost material, rotting apples, slugs, or attracted by placing apples on compost heaps. The prime time for finding C. elegans in the Northern Hemisphere is from September until November. Worms can be washed out of collected substrate material by immersing the substrate in buffer solution, followed by the collection of nematodes and their transfer onto nematode growth medium or PCR buffer for subsequent analysis. We further illustrate how the samples can be used to isolate and purify the worm-associated microorganisms and to process worms for 16S ribosomal RNA analysis of microbiota community composition. Overall, the described methods may stimulate new research on the characterization of the C. elegans microbiota across habitats and geographic locations, thereby helping to obtain a comprehensive understanding of the diversity and stability of the nematode's microbiota as a basis for future functional research.

Isolation and Characterization of the Natural Microbiota of the Model Nematode Caenorhabditis elegans

Caenorhabditis elegans is one of the main model species in biology, yet almost all research is performed in the absence of its naturally associated microbes. The methods described here will help to improve our understanding of the diversity of associated microbes as a basis for future functional C. elegans research.

The nematode Caenorhabditis elegans interacts with a large diversity of microorganisms in nature. In general, C. elegans is commonly found in rotten plant ...

Rapid Isolation of Wild Nematodes by Baermann Funnel

Summary

Explore More Videos

Chapters in this video

Helminth Collection and Identification from Wildlife

Soil Sampling and Isolation of Entomopathogenic Nematodes (Steinernematidae, Heterorhabditidae)

In vivo and In vitro Rearing of Entomopathogenic Nematodes (Steinernematidae and Heterorhabditidae)

Cultivation of Heligmosomoides Polygyrus: An Immunomodulatory Nematode Parasite and its Secreted Products

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

Isolation and Characterization of the Natural Microbiota of the Model Nematode Caenorhabditis elegans