The authors acknowledge Dr. Qing Yu (Agriculture and Agri-Food Canada) for providing the Ditylenchus dipsaci culture and for advice on culture methods; Dr. Benjamin Mimee (Agriculture and Agri-Food Canada) and Nathalie Dauphinais (Agriculture and Agri-Food Canada) for advice on in vitro culture of plant-parasitic nematodes; Dr. Andrew Burns and Sean Harrington for helpful suggestions on the project and the manuscript. JK is an NSERC Alexander Graham Bell Canada Graduate Scholar. PJR is supported by a CIHR project grant (313296). PJR is a Canada Research Chair (Tier 1) in Chemical Genetics.

The D. dipsaci strain G-137 used for the present work was collected from Fish Lake 4 Variety garlic in Prince Edward County and was provided by Agriculture and Agri-food Canada. If starting a fresh culture, consult Poirier et al. for inoculation methodology15.
1. Culturing of D. dipsaci
<ol>
	<li>Prepare the media and plates following previously reported work15.
	<ol>
		<li>Prepare 500 mL of Nutrient agar (NA) media with 23 g/L of NA (see Table of Materials) and ultrapure water. Using sterile technique, pour 25 mL of autoclaved NA media into 20 disposable Petri dishes (100 mm diameter x 15 mm deep). Allow agar to solidify at room temperature (22 &#176;C) with the lid on for ~2 h and set aside for later use.</li>
		<li>Prepare 500 mL of Gamborg B-5 (GA) media containing 3.2 g/L of GA basal medium with minimal organics, 20 g/L of sucrose, 15 g/L of agar, and distilled water (see Table of Materials). Using sterile technique, pour 50 mL of autoclaved GA media in 10 disposable Petri dishes (100 mm diameter x 25 mm deep). Allow agar to solidify at room temperature (22 &#176;C) with the lid on for ~5 h and store upright at room temperature in a sterile bag until the sprout transfer. 
		NOTE: Excess media plates are made in case contamination occurs.</li>
	</ol>
	</li>
	<li>Perform seed sterilization following a method modified from a previous work15.
	<ol>
		<li>Autoclave one 2 L beaker with a stir bar, 2 forceps, a glass petri dish, and 1 L of distilled water. Prepare 200 mL of 95% EtOH solution and 200 mL of a 15% commercial bleach solution.</li>
		<li>Use pea seeds (see Table of Materials). Pour 150 seeds into a sterile 2 L beaker with a stir bar near a Bunsen burner flame on a lab bench.</li>
		<li>Add 200 mL of 95% EtOH to the seeds within the beaker, stir vigorously on the stir plate for 5 min, and then pour off EtOH in a waste container.</li>
		<li>Pour bleach solution into the beaker to completely immerse the seeds. Stir vigorously on a stir plate for 20 min then pour off bleach in a waste container.</li>
		<li>Pour distilled water into the beaker to immerse seeds and stir vigorously on a stir plate for 20 min. Repeat water washes three times, pouring off distilled water after each wash. After the final water wash, pour sterilized seeds into the glass Petri dish.</li>
		<li>To check for contamination, transfer 6 seeds to each 10 cm NA plate (prepared in step 1.1.1) in the laminar flood hood using sterilized forceps. Arrange the seeds around the plate&#39;s circumference (plumper seeds work best). Wrap the plates individually in laboratory wrapping film and incubate in the dark for 3 days at 26 &#176;C. 
		NOTE: Plating more seeds than needed will allow for selective use of non-contaminated seeds.</li>
	</ol>
	</li>
	<li>Perform sprout transfer following the step below.
	<ol>
		<li>In a laminar flow hood, use sterilized forceps to plate 2 non-contaminated seeds on each GA plate (prepared in step 1.1.2). Wrap the plates individually in laboratory wrapping film and incubate at room temperature for 7-10 days to allow seeds to sprout.</li>
	</ol>
	</li>
	<li>Perform in vitro rearing following a method modified from a previous work15.
	<ol>
		<li>Prepare 50 mL of 20 g/L sucrose solution. Filter sterilize sucrose solution and set aside.</li>
		<li>In a laminar flow hood, cut a piece of agar containing root tissue (~2 cm3) from an existing culture plate. Pipette 500 &#181;L of sucrose solution on new GA plate with pea seedlings and place agar cube on top of sucrose. Wrap the plates individually in laboratory wrapping film and maintain the culture in a box lined with aluminum foil at room temperature (~22 &#176;C).</li>
		<li>Subculture nematodes on fresh GA plates every 8-9 weeks to maintain the culture. Nematodes are ready to be extracted after ~8 weeks.</li>
	</ol>
	</li>
</ol>
2. Extraction and collection of D. dipsaci
<ol>
	<li>Perform the extraction of D. dipsaci following the steps below.
	<ol>
		<li>Autoclave a 50 mL beaker, an 80 mm funnel, 150 mL distilled water, and coffee filters. Place the funnel into the beaker and line the funnel with sterile coffee filter.</li>
		<li>In a laminar flow hood, cut the agar and the root tissue into 1 cm3 using a sterile scalpel. Transfer the agar cubes into the coffee filter-lined funnel and slowly pour distilled water on the agar to moisten the coffee filter.</li>
		<li>Remove the coffee filter-lined funnel from the beaker and fill the beaker with distilled water until the water level is just touching the bottom of the filter once the coffee filter-lined funnel is replaced.</li>
		<li>Cover the coffee filter-lined funnel and beaker with aluminum foil. Leave overnight (16 h) on the benchtop to allow worms to move through the coffee filter into the collection beaker. 
		NOTE: A nematode-culture plate is ready for extraction when worms have crawled into the agar when examined with a dissection microscope. Typically, a plate is ready for extraction 6-8 weeks after its initial inoculation.</li>
	</ol>
	</li>
	<li>Perform the collection of D. dipsaci. 
	NOTE: The next day, the D. dipsaci worms will have settled to the bottom of the beaker.
	<ol>
		<li>Remove the coffee filter-lined funnel and aspirate the top 40 mL of water from the collection beaker; ensure not to disrupt the settled worms. Using a 10 mL plastic serological pipette, collect the remaining liquid into a 15 mL conical centrifuge tube. 
		&#8203;NOTE: This collection can be used directly in assays. Place at 4 &#176;C if they will not be used within 24 hours. The worms can be left for 3 days at 4 &#176;C with no visible impact on mobility. Removing the funnel may disturb the worms in the collection beaker. Let them settle to the bottom before collecting.</li>
	</ol>
	</li>
</ol>
3. In vitro small molecule screen
<ol>
	<li>Prepare the assay plates following the steps below.
	<ol>
		<li>Pour autoclaved distilled water into a sterile trough and, using a multichannel pipette, dispense 40 &#181;L of distilled water from the trough into each well of a flat-bottom 96-well plate.</li>
	</ol>
	</li>
	<li>Prepare pinning tool and add the chemicals
	<ol>
		<li>Set up pinner trays (see Table of Materials) near a Bunsen burner flame on a lab bench. Add the following to successive pinner trays: 25 mL of pin cleaning solution, 35 mL of 50% DMSO (in water), 45 mL of distilled water, 55 mL of 70% EtOH, and 65 mL of 95% EtOH. Place one piece of blotting paper in front of each tray</li>
		<li>Clean the pinning tool by emersing the pins in the cleaning solution and moving the pins up and down three times (3x) in the solution. In this protocol, &#39;3x&#39; and &#39;10x&#39; are defined as moving the pinner up and down in a solution three or ten times, respectively. Blot the pins on the blotting paper. Repeat this procedure once more.
		<ol>
			<li>Next, rinse the pins 3x in distilled water, followed by blotting the pins. Repeat the procedure once more.</li>
			<li>Lastly, rinse the pins 3x in 95% EtOH, followed by blotting the pins. Repeat once more. Flame the pinner and allow the ethanol to evaporate.</li>
		</ol>
		</li>
		<li>Add chemicals from the 96-well chemical stock plates to assay plates by pinning 3x into the chemical plate, then transferring the pins 10X into the assay plate. Blot onto paper in front of the cleaning solution.</li>
		<li>Clean pinning tool between plates by washing in the following order, blotting in between on blotting paper: 3x in 50% DMSO (once), 3x in distilled water (once), 3x in 75% EtOH (once), 3x in 95% EtOH (twice). Flame the pinner and allow ethanol to evaporate.</li>
		<li>Repeat step 3.2.2 when all pinning is completed. 
		NOTE: The screening was performed at a final concentration of 60 &#181;M.</li>
	</ol>
	</li>
	<li>Addition of worms
	<ol>
		<li>Count the number of nematodes from the collection by first resuspending and then pipetting 5 &#181;L using low retention tips onto a slide for observation. Count the number of nematodes in 5 &#181;L using a dissection microscope.
		<ol>
			<li>Adjust the concentration to 2 worms/&#181;L using sterile distilled water. Using a multichannel pipette and a trough, add 10 &#181;L (~20 worms) to each well of the 96-well plates. 
			&#8203;NOTE: Approximately 15,000 D. dipsaci nematodes will be collected per culture plate using the described culture and collection method. Twenty worms are used per well because the small number facilitates the clear visualization and accounting of mobile and immobile ones.</li>
		</ol>
		</li>
		<li>Seal the plates in laboratory wrapping film and wrap with a damp paper towel. Place in box and affix on a sticky pad in 20 &#176;C shaking incubator set at 200 rpm. Ensure that plates are stabilized in the box by adding an extra damp paper towel to ensure minimal movement of plates.</li>
	</ol>
	</li>
</ol>
4. Data collection and analysis
<ol>
	<li>Observe plates on day 5 under a dissecting microscope. Count the number of mobile and total number of D. dipsaci in DMSO solvent controls and drug-treated wells.
	<ol>
		<li>If the worms are relatively immobile, add 2 &#181;L of 1 M NaOH to a final concentration of 40 mM to the well to stimulate movement19, 20. 
		NOTE: After adding NaOH, worms will move instantly and need to be viewed within 5 min. The number of mobile worms and the total number of worms will be used to calculate the proportion of mobile worms. The length of the assay may change depending on the aim of the screen.</li>
	</ol>
	</li>
	<li>Calculate the proportion of mobile worms. In the D. dipsaci screens, wells that reproducibly yielded 0% mobile worms are categorized as strong hits. 
	NOTE: Prolonged exposure of plates on the stage of dissection microscopes is avoided because the light source can heat the plates and induce variable effects on the bioassay.</li>
</ol>

<ol>
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P., Hongli, S., Jingwu, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nematodes+threats+to+global+food+security.">Nematodes threats to global food security.</a> Acta Agriculturae Scandinavica, Section B - Soil &amp; Plant Science. 63 (5), 420-425 (2013).</li><li>Chen, Z. X., Chen, S. Y., Dickson, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nematicides:+Past+and+present+uses.">Nematicides: Past and present uses.</a> Nematology Vol 2, Nematode Management and Utilization. , 1179-1200 (2004).</li><li>Desaeger, J., Wram, C., Zasada, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+reduced-risk+agricultural+nematicides+-+and+review.">New reduced-risk agricultural nematicides - and review.</a> Journal of nematology. 52, (2020).</li><li>Donely, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+USA+lags+behind+other+agricultural+nations+in+banning+harmful+pesticides.">The USA lags behind other agricultural nations in banning harmful pesticides.</a> Environmental Health. 18 (1), 44 (2019).</li><li>Polansek, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monsanto+halts+launch+of+chemical+after+users+complain+of+rashes.">Monsanto halts launch of chemical after users complain of rashes.</a> Reuters. , (2017).</li><li>Sturhan, D., Brzeski, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stem+and+bulb+nematodes,+Ditylenchus+spp.">Stem and bulb nematodes, Ditylenchus spp.</a> Manual of agricultural nematology. , 423-464 (1991).</li><li>The European and Mediterranean Plant Protection Organization (EPPO). <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Data+Sheets+on+Quarantine+Pests;+Ditylenchus+dipsaci.+Prepared+by+CABI+and+EPPO+for+the+EU+under+Contract+90/399003.">Data Sheets on Quarantine Pests; Ditylenchus dipsaci. Prepared by CABI and EPPO for the EU under Contract 90/399003.</a> The European and Mediterranean Plant Protection Organization (EPPO). , (2021).</li><li>Quebec (Canada), Agriculture, P&#234;cherie et Alimentation Quebec. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=R&#233;seau+d'avertissements+phytosanitaires+(RAP).+Avertissement+Carotte,+C&#233;leri,+Laitue,+Oignon,+Poireau+[Warning+carrot+celery,+lettuce,+onion,+leek].">R&#233;seau d'avertissements phytosanitaires (RAP). Avertissement Carotte, C&#233;leri, Laitue, Oignon, Poireau [Warning carrot celery, lettuce, onion, leek].</a> Quebec (Canada), Agriculture, P&#234;cherie et Alimentation Quebec. , (2011).</li><li>Abawi, G. S., Moktan, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bloat+nematode+problem+on+garlic:+symptoms,+distribution,+and+management+guidelines.">Bloat nematode problem on garlic: symptoms, distribution, and management guidelines.</a> Department of Plant Pathology and Plant-microbe Biology, Geneva. , (2010).</li><li>Taylor, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+7.">Chapter 7.</a> Nematodes and other worms. , 143-234 (2019).</li><li>Storelli, A., Keiser, A., Eder, R., Jenni, S., Kiewnick, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+fluopyram+for+the+control+of+Ditylenchus+dipsaci+in+sugar+beet.">Evaluation of fluopyram for the control of Ditylenchus dipsaci in sugar beet.</a> Journal of nematology. 52, 1-10 (2020).</li><li>Oka, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nematicidal+activity+of+fluensulfone+against+some+migratory+nematodes+under+laboratory+conditions.">Nematicidal activity of fluensulfone against some migratory nematodes under laboratory conditions.</a> Pest Management Science. 70, 1850 (2014).</li><li>Poirier, S., Dauphinais, N., Van Der Heyden, H., V&#233;ronneau, P. Y., B&#233;lair, G., Gravel, V., Mimee, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Host+Range+and+Genetic+Characterization+of+Ditylenchus+dipsaci+Populations+from+Eastern+Canada.">Host Range and Genetic Characterization of Ditylenchus dipsaci Populations from Eastern Canada.</a> Plant disease. 103, 456-460 (2019).</li><li>Storelli, A., Kiewnick, S., Daub, M., 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=Virulence+and+pathogenicity+of+four+Ditylenchus+dipsaci+populations+on+sugar+beet.">Virulence and pathogenicity of four Ditylenchus dipsaci populations on sugar beet.</a> Eur J Plant Pathol. 161, 63-71 (2021).</li><li>K&#252;hnhold, V., Kiewnick, S., Sikora, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+an+in+vivo+bioassay+to+identify+sugar+beet+resistance+to+the+stem+nematode+Ditylenchus+dipsaci.">Development of an in vivo bioassay to identify sugar beet resistance to the stem nematode Ditylenchus dipsaci.</a> Nematology. 8 (5), 641-645 (2006).</li><li>Burns, A., Luciani, G., Musso, G. <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+is+a+useful+model+for+anthelmintic+discovery.">Caenorhabditis elegans is a useful model for anthelmintic discovery.</a> Nat Communications. 6, (2015).</li><li>Chen, S. Y., Dickson, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Technique+for+Determining+Live+Second-stage+Juveniles+of+Heterodera+glycines.">A Technique for Determining Live Second-stage Juveniles of Heterodera glycines.</a> Journal of nematology. 32, 117-121 (2000).</li><li>Xiang, N., Lawrence, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+In+Vitro+Techniques+for+Distinguishing+between+Live+and+Dead+Second+Stage+Juveniles+of+Heterodera+glycinesand+Meloidogyne+incognita.">Optimization of In Vitro Techniques for Distinguishing between Live and Dead Second Stage Juveniles of Heterodera glycinesand Meloidogyne incognita.</a> PLoS ONE. 11 (5), (2016).</li><li>US EPA. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluopyram.">Fluopyram.</a> US EPA. , (2016).</li><li>US EPA. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluopyram.">Fluopyram.</a> US EPA. , (2020).</li><li>US EPA. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxamyl+Facts.+Prevention,+Pesticides+and+Toxic+Substances.">Oxamyl Facts. Prevention, Pesticides and Toxic Substances.</a> US EPA. , (2000).</li></ol>

The authors declare no conflicts of interest.

Critical steps 
Despite the protocol&#39;s simplicity, there are critical steps in the protocol that deserve additional attention to maximize the likelihood of success. First, overbleaching the seeds can disrupt their growth. Therefore, limiting the seeds&#39; time in the bleaching solution to 20 minutes or less is essential. Second, as previously noted by Storelli et al., the apparent health of the nematodes decreases over time when stored at 4 &#176;C16. Using the nematodes soon after their collection provides additional confidence that optimal screening conditions can be achieved. Should longer-term storage be necessary, ensure that the tube&#39;s lid is not tight to allow oxygen exchange. Third, ensuring that the peas grow on the NA plates for the suggested time enables the experimenter to judge which seeds are contaminated. Fourth, overgrowing the peas on the GA plates before adding the nematodes will weaken infection and reduce the nematode yield. Finally, many factors that are difficult to control can impact screening results. Therefore, it is essential to perform multiple independent replicate screens on different days and ideally with PPNs collected from different culture plates to ensure the reproducibility of results.
Limitations of method 
A limitation to the protocol is that it fails to synchronize the developmental stage of the collected worms, which range from juveniles to adults. Hence, strong hits revealed by any screen are likely effective at multiple stages. However, the protocol increases the risk of overlooking effective stage-specific hits. A second consideration is that in vitro screening should be considered the first step in a nematicide-discovery pipeline; soil-based assays are an excellent addition to a pipeline to test the translatability of the hits.
Significance and application of the protocol 
The protocols described herein are simple and easily replicated. Furthermore, this protocol has been successfully applied to other PPNs in the lab, including Pratylenchus penetrans, by making only slight modifications. Developing new and safe PPN control measures is essential to ensure global food security. This is especially true for species like D. dipsaci that are generally tolerant to a wide variety of currently acceptable chemical nematicides13,14. Hence, the protocols outlined here have the potential to make an important contribution to human health on a global scale.

Plant-parasitic nematodes (PPNs) are estimated to be responsible for the loss of 12.3% of global food production and cause an estimated 157 billion dollars in damage annually1,2,3. Unfortunately, the ability to control PPNs is waning because effective chemical nematicides have been banned or are facing escalating restrictions because of human safety and environmental concerns. This is primarily due to the poor nematode selectivity of previous generations of pesticides4. Over the last 25 years, six new chemical nematicides have been piloted or introduced into the market5. One of these has already been banned in Europe, and another has been discontinued while being investigated for its impact on human heatlh6,7. Hence, there is a pressing need for new nematicides that are highly selective for PPNs.
The stem and bulb nematode, Ditylenchus dipsaci (D. dipsaci) is an economically impactful PPN4. D. dipsaci infects nearly 500 plant species across 30 biological races and targets some of the most agriculturally important crops such rye, oats, garlic, onion, and leek8,9. For example, D. dipsaci has recently scourged garlic fields in Ontario and Quebec, resulting in losses of up to 90%10,11. Its geographical distribution is nearly ubiquitous and includes the Americas (including California and Florida), Europe, much of Asia (including China), and Oceania9. D. dipsaci is a migratory endoparasite that enters the stomata on leaves or wounds and lenticels where they release enzymes to break down the cell wall12. Compounding the impact of D. dipsaci on crops, the damage caused by the PPN makes the plant susceptible to secondary infection11. Unfortunately, D. dipsaci shows high tolerance levels to current nematicides compared to other nematode strains13,14.
This protocol describes the culturing of D. dipsaci, and its use in large-scale screens for small molecule candidate nematicides.&#160;Briefly, D. dipsaci populations are maintained and expanded on pea plants cultured in sterile Gamborg B-5 (GA) media15. Before growing seed sprouts on GA medium, the seeds must be sterilized through a series of washes and plated on nutrient agar (NA) to check for contamination. Seed sterilization is essential to detect bacterial and fungal contaminants that may be present. The non-contaminated seeds are then transferred to GA plates, where seed sprouts will grow in preparation for infection. The GA plates containing seed sprouts are infected with nematodes from a previous culture plate by transferring a piece of agar containing root tissue to the fresh plates. After 6-8 weeks, the nematodes are extracted from the GA media and are filtered through a coffee filter-lined funnel into a collection beaker. The nematodes can be used in various bioassays once a suitable number has been collected. The technique described in this protocol generates approximately 15,000 D. dipsaci per culture plate. Alternative protocols to cultivate D. dipsaci have been published16,17.
An in vitro small molecule screening assay based on previous work18&#160;is also described here.&#160;As a proxy of worm health, the mobility of 20 nematodes per well is examined after 5 days of small molecule exposure. To better visualize worm mobility, NaOH is added to increase the movement of live worms19,20.&#160;This protocol allows medium-throughput screening and provides valuable data to assess the nematicidal potential of small molecules. If a different nematode collection technique is used16,17, the small molecule screening methodology described herein can nevertheless be implemented.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 mL tube</td>
			<td>Sarstedt</td>
			<td>62.554.205</td>
			<td></td>
		</tr>
		<tr>
			<td>2 forceps</td>
			<td>Almedic</td>
			<td>7747-A10-108</td>
			<td></td>
		</tr>
		<tr>
			<td>2L beaker</td>
			<td>Pyrex</td>
			<td>CLS10002L</td>
			<td></td>
		</tr>
		<tr>
			<td>50mL beaker</td>
			<td>Pyrex</td>
			<td>CLS100050</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well plate</td>
			<td>Sarstedt</td>
			<td>83.3924</td>
			<td></td>
		</tr>
		<tr>
			<td>aluminium foil</td>
			<td>Alcan Plus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>bacteriological agar</td>
			<td>BioShop</td>
			<td>AGR001.5</td>
			<td></td>
		</tr>
		<tr>
			<td>coffee filter</td>
			<td>No name brand</td>
			<td>716</td>
			<td></td>
		</tr>
		<tr>
			<td>commercial bleach 6%</td>
			<td>Lavo Pro 6</td>
			<td>DIN102358107</td>
			<td></td>
		</tr>
		<tr>
			<td>disposable petri dishes (10cm x 1.5cm)</td>
			<td>Fisherbrand</td>
			<td>FB0875712</td>
			<td></td>
		</tr>
		<tr>
			<td>disposable petri dishes (10cm x 2.5cm)</td>
			<td>Sigma-Aldrich</td>
			<td>Z358762</td>
			<td></td>
		</tr>
		<tr>
			<td>dissecting scope</td>
			<td>Leica</td>
			<td>Leica MZ75</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-Aldrich</td>
			<td>472301-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Funnel</td>
			<td>VWR</td>
			<td>414004-270</td>
			<td></td>
		</tr>
		<tr>
			<td>Gamborg B5</td>
			<td>Sigma-Aldrich</td>
			<td>G5893-10L</td>
			<td></td>
		</tr>
		<tr>
			<td>glass petri dish</td>
			<td>VWR</td>
			<td>75845-546</td>
			<td></td>
		</tr>
		<tr>
			<td>glass slide</td>
			<td>MAGNA</td>
			<td>60-1200</td>
			<td></td>
		</tr>
		<tr>
			<td>Lint-Free Blotting Paper</td>
			<td>V&#38;P Scientific</td>
			<td>VP 522-100</td>
			<td></td>
		</tr>
		<tr>
			<td>mutlichannel pipette</td>
			<td>Eppendorf research plus</td>
			<td>3125000036</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Sigma-Aldrich</td>
			<td>S8045-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>nutrient agar</td>
			<td>Sigma-Aldrich</td>
			<td>70148-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>parafilm</td>
			<td>Bemis</td>
			<td>PM-996</td>
			<td></td>
		</tr>
		<tr>
			<td>pea seeds</td>
			<td>Ontario Seed Company</td>
			<td>D-1995-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>pin cleaning solutions</td>
			<td>V&#38;P Scientific</td>
			<td>VP110A</td>
			<td></td>
		</tr>
		<tr>
			<td>pinner</td>
			<td>V&#38;P Scientific</td>
			<td>VP381N</td>
			<td></td>
		</tr>
		<tr>
			<td>pinner rinse trays</td>
			<td>V&#38;P Scientific</td>
			<td>VP 421</td>
			<td></td>
		</tr>
		<tr>
			<td>pipette tips- low retention ? Reg. 200uL</td>
			<td>LABFORCE</td>
			<td>1159M44</td>
			<td></td>
		</tr>
		<tr>
			<td>reagent reservoir with lid for multichannel pipettes</td>
			<td>Sigma-Aldrich</td>
			<td>BR703459</td>
			<td></td>
		</tr>
		<tr>
			<td>shaking incubator</td>
			<td>New Brunswick Scientific</td>
			<td>I26</td>
			<td></td>
		</tr>
		<tr>
			<td>sterile surgical blade</td>
			<td>MAGNA</td>
			<td>sb21-100-a</td>
			<td></td>
		</tr>
		<tr>
			<td>stir bar</td>
			<td>Fisherbrand</td>
			<td>2109 - 1451359</td>
			<td></td>
		</tr>
		<tr>
			<td>sucrose</td>
			<td>BioShop</td>
			<td>SUC507.1</td>
			<td></td>
		</tr>
	</tbody>
</table>

culturing and screening the plant parasitic nematode ditylenchus dipsaci

Plant-parasitic nematodes (PPNs) destroy over 12% of global food crops every year, which equates to roughly 157 billion dollars (USD) lost annually. With a growing global population and limited arable land, controlling PPN infestation is critical for food production. Compounding the challenge of maximizing crop yields are the mounting restrictions on effective pesticides because of a lack of nematode selectivity. Hence, developing new and safe chemical nematicides is vital to food security. In this protocol, the culture and collection of the PPN species Ditylenchus dipsaci are demonstrated. D. dipsaci is both economically damaging and relatively resistant to most modern nematicides. The current work also explains how to use these nematodes in screens for novel small molecule nematicides and reports on data collection and analysis methodologies. The demonstrated pipeline affords a throughput of thousands of compounds per week and can be easily adapted for use with other PPN species such as Pratylenchus penetrans. The techniques described herein can be used to discover new nematicides, which may, in turn, be further developed into highly selective commercial products that safely combat PPNs to help feed an increasingly hungry world.

Independent replicates reveal reproducible hits 
To illustrate the expected variation between the replicate screens, the means and variation in sample mobility are plotted from three representative plates from a recent screen (Figure 1). Three replicates of the screen were performed on three different days. All three plates have negative (solvent-only) controls (darker bars), and samples within the set contained established nematicides. The remaining compounds are from a custom library currently being characterized in the Roy lab. Compared to similar screens done with C. elegans with the same molecules (SC, JK, PJR, unpublished results), the hit rate with D. dipsaci is significantly lower, and many drug plates exhibit no activity (Figure 1A). Some plates have fully reproducible hits (Figure 1B,C), while others vary in activity (Figure 1C). Relative to other species screened (not shown), D. dipsaci shows less variability in its response to compounds. Regardless, replicates are considered necessary in the identification of reproducible hits.
Nematicides vary in their ability to immobilize Ditylenchus dipsaci 
Of the seven characterized nematicides tested against D. dipsaci, only Fluopyram exhibits robust activity in the assay described herein (Figure 1C). This is consistent with previous work showing that D. dipsaci is tolerant of nematicides13,14. Fluopyram inhibits complex II of the electron transport chain in a nematode-selective manner and is a commercial nematicide used to control a variety of PPNs, including Rotylenchulus reniformis and Meloidogyne incognita4,21,22. Fluopyram and one of the nematicides that lacked activity at the concentration tested (Oxamyl)23 were investigated in more detail through a dose-response analysis with D. dipsaci. Fluopyram induces an apparent dose-dependent effect on D. dipsaci mobility with an EC50 of 9.3 &#956;M (with a 95% confidence interval between 8.2 to 10.5 &#956;M) (Figure 2A,B). This result is expected based on published in vitro results from Storelli et al., 202013. Oxamyl has no significant effect on mobility up to a concentration of 120 &#181;M (p = 0.3632, unpaired t-test) (Figure 2C,D).
Sodium hydroxide improves assay sensitivity 
In contrast to the near-continuous swimming activity of C. elegans in liquid culture18, D. dipsaci animals are dramatically less mobile. This is not uncommon among parasitic nematodes in culture16. To help distinguish &#39;resting&#39; worms from sick worms, 40 mM of NaOH is used at the assay endpoint to stimulate movement in those individuals who are capable (Figure 3)19,20. This technique allows for the clear identification of small molecules that immobilize worms, given that all worms in negative control wells move and yield an exceptionally low background of false positives in the screen.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63438/63438fig01v2.jpg" /> 
Figure 1: Examples of screen output. Three biological replicates (open circles) with D. dipsaci were performed against the small molecules (60 &#181;M) in each of the three plates shown. All three plates have 0.6% of DMSO solvent-only controls (darker bars). Except where otherwise noted with solvent controls or known nematicides, the wells of the three plates contain relatively uncharacterized drug-like compounds purchased from vendors (see Burns et al., 2015)18. (A) A 96 well plate from the small molecule library lacks any molecule with observable bioactivity against D. dipsaci. (B) A 96 well plate from the small molecule library has a single molecule that reproducibly disrupts D. dipsaci mobility. (C) A 96 well drug plate containing the characterized nematicides fluopyram (fluo), iprodione (ipro), abamectin (abam), fluensulfone (flue), tioxazafen (tiox), oxamyl (oxam), and wact-11. The error bars represent the standard error of the mean. <a href="https://www.jove.com/files/ftp_upload/63438/63438fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63438/63438fig02.jpg" /> 
Figure 2: Example of positive and negative results using mobility assay. (A) Examples of the terminal phenotypes after the exposure of D. dipsaci to increasing concentrations of fluopyram after 5 days before addition of NaOH. (B) A dose-response analysis of the movement of D. dipsaci after 5 days of exposure to the indicated concentrations of fluopyram. (C,D) The same as A,B, except for oxamyl. Each graph shows trials done on two separate days with three replicates each day. The y-axis of each graph indicates the fraction mobile of the worms in each well calculated as the number of animals moving relative to the total number of animals in the well. The error bars on both graphs represent the standard error of the mean. The scale bar represents 1 mm. <a href="https://www.jove.com/files/ftp_upload/63438/63438fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63438/63438fig03.jpg" /> 
Figure 3: Mobility of D. dipsaci after adding 40 mM of NaOH. The effect of NaOH addition to D. dipsaci samples in the presence of solvent-only control (A), tioxazafen (B), or fluopyram (C) after 5 days of co-incubation. The scale bar represents 1 mm. <a href="https://www.jove.com/files/ftp_upload/63438/63438fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci at JoVE.com

Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

The present protocol describes a reliable and straightforward method for culturing, collecting, and screening Ditylenchus dipsaci.

Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

Plant-parasitic nematodes (PPNs) destroy over 12% of global food crops every year, which equates to roughly 157 billion dollars (USD) lost annually. With ...

culturing-screening-plant-parasitic-nematode-ditylenchus

Research

JoVE Journal

Biology

3.3K Views.  University of Toronto. The present protocol describes a reliable and straightforward method for culturing, collecting, and screening Ditylenchus dipsaci. 

Video: Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression analyses by RNA interference and stable transgenesis remain limited in many species due to the particular anatomy and molecular cellular biology of the organism. For example, outside of the crown group Caenorhabditis that includes Caenorhabditis elegans3, stably transmitted transgenic lines in non-Caenorhabditis species have not been reported in this specious phylum (Nematoda), with the exception of Strongyloides stercoralis4 and Pristionchus pacificus5. To facilitate the expanding role of P. pacificus in the study of development, evolution, and behavior6-7, we describe here the current methods to use microinjection for making transgenic animals and gene knock down by RNAi. Like the gonads of C. elegans and most other nematodes, the gonads of P. pacificus is syncitial and capable of incorporating DNA and RNA into the oocytes when delivered by direct microinjection. Unlike C. elegans however, stable transgene inheritance and somatic expression in P. pacificus requires the addition of self genomic DNA digested with endonucleases complementary to the ends of target transgenes and coinjection markers5. The addition of carrier genomic DNA is similar to the requirement for transgene expression in Strongyloides stercoralis4 and in the germ cells of C. elegans. However, it is not clear if the specific requirement for the animals' own genomic DNA is because P. pacificus soma is very efficient at silencing non-complex multi-copy genes or that extrachromosomal arrays in P. pacificus require genomic sequences for proper kinetochore assembly during mitosis. The ventral migration of the two-armed (didelphic) gonads in hermaphrodites further complicates the ability to inject both gonads in individual worms8. We also demonstrate the use of microinjection to knockdown a dominant mutant (roller,tu92) by injecting double-stranded RNA (dsRNA) into the gonads to obtain non-rolling F1 progeny. Unlike C. elegans, but like most other nematodes, P. pacificus PS312 is not receptive to systemic RNAi via feeding and soaking and therefore dsRNA must be administered by microinjection into the syncitial gonads. In this current study, we hope to describe the microinjection process needed to transform a Ppa-egl-4 promoter::GFP fusion reporter and knockdown a dominant roller prl-1 (tu92) mutant in a visually informative protocol.

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

In model organisms, transgenesis can manipulate gene functions while RNAi can knockdown specific mRNA transcripts 1-2. This protocol aims to illustrate the techniques needed to introduce stably transmitted DNA and transient double stranded RNA into the necromenic nematode Pristionchus pacificus for studies in evolutionary, developmental, and behavioral biology.

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression ...

Fluorescence-microscopy Screening and Next-generation Sequencing: Useful Tools for the Identification of Genes Involved in Organelle Integrity

This protocol describes a fluorescence microscope-based screening of Arabidopsis seedlings and describes how to map recessive mutations that alter the subcellular distribution of a specific tagged fluorescent marker in the secretory pathway. Arabidopsis is a powerful biological model for genetic studies because of its genome size, generation time, and conservation of molecular mechanisms among kingdoms. The array genotyping as an approach to map the mutation in alternative to the traditional method based on molecular markers is advantageous because it is relatively faster and may allow the mapping of several mutants in a really short time frame. This method allows the identification of proteins that can influence the integrity of any organelle in plants. Here, as an example, we propose a screen to map genes important for the integrity of the endoplasmic reticulum (ER). Our approach, however, can be easily extended to other plant cell organelles (for example see1,2), and thus represents an important step toward understanding the molecular basis governing other subcellular structures.

A fundamental quest in cell biology is to define the mechanisms that underlie the identity of the organelles that make eukaryotic cells. Here we propose a method to identify the genes responsible for the morphological and functional integrity of plant organelles using fluorescence microscopy and next-generation sequencing tools.

This protocol describes a fluorescence microscope-based screening of Arabidopsis seedlings and describes how to map recessive mutations that alter the ...

Electroantennographic Bioassay as a Screening Tool for Host Plant Volatiles

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host plant.1,2 When a host plant is an important agronomic commodity feeding damage by insect pests can inflict serious economic losses to growers. Accordingly, kairomones can be used as attractants to lure or confuse these insects and, thus, offer an environmentally friendly alternative to pesticides for insect control.3 Unfortunately, plants can emit a vast number volatiles with varying compositions and ratios of emissions dependent upon the phenology of the commodity or the time of day. This makes identification of biologically active components or blends of volatile components an arduous process. To help identify the bioactive components of host plant volatile emissions we employ the laboratory-based screening bioassay electroantennography (EAG). EAG is an effective tool to evaluate and record electrophysiologically the olfactory responses of an insect via their antennal receptors. The EAG screening process can help reduce the number of volatiles tested to identify promising bioactive components. However, EAG bioassays only provide information about activation of receptors. It does not provide information about the type of insect behavior the compound elicits; which could be as an attractant, repellent or other type of behavioral response. Volatiles eliciting a significant response by EAG, relative to an appropriate positive control, are typically taken on to further testing of behavioral responses of the insect pest. The experimental design presented will detail the methodology employed to screen almond-based host plant volatiles4,5 by measurement of the electrophysiological antennal responses of an adult insect pest navel orangeworm (Amyelois transitella) to single components and simple blends of components via EAG bioassay. The method utilizes two excised antennae placed across a "fork" electrode holder. The protocol demonstrated here presents a rapid, high-throughput standardized method for screening volatiles. Each volatile is at a set, constant amount as to standardize the stimulus level and thus allow antennal responses to be indicative of the relative chemoreceptivity. The negative control helps eliminate the electrophysiological response to both residual solvent and mechanical force of the puff. The positive control (in this instance acetophenone) is a single compound that has elicited a consistent response from male and female navel orangeworm (NOW) moth. An additional semiochemical standard that provides consistent response and is used for bioassay studies with the male NOW moth is (Z,Z)-11,13-hexdecadienal, an aldehyde component from the female-produced sex pheromone.6-8

A method to rapidly screen host plant volatiles by measurement of the electrophysiological response of adult navel orangeworm (Amyelois transitella) antennae to single components and blends via electroantennographic analysis is demonstrated.

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host ...

Using RNA-mediated Interference Feeding Strategy to Screen for Genes Involved in Body Size Regulation in the Nematode C. elegans

Double-strand RNA-mediated interference (RNAi) is an effective strategy to knock down target gene expression1-3. It has been applied to many model systems including plants, invertebrates and vertebrates. There are various methods to achieve RNAi in vivo4,5. For example, the target gene may be transformed into an RNAi vector, and then either permanently or transiently transformed into cell lines or primary cells to achieve gene knockdown effects; alternatively synthesized double-strand oligonucleotides from specific target genes (RNAi oligos) may be transiently transformed into cell lines or primary cells to silence target genes; or synthesized double-strand RNA molecules may be microinjected into an organism. Since the nematode C. elegans uses bacteria as a food source, feeding the animals with bacteria expressing double-strand RNA against target genes provides a viable strategy6. Here we present an RNAi feeding method to score body size phenotype. Body size in C. elegans is regulated primarily by the TGF- &beta; - like ligand DBL-1, so this assay is appropriate for identification of TGF-&beta; signaling components7. We used different strains including two RNAi hypersensitive strains to repeat the RNAi feeding experiments. Our results showed that rrf-3 strain gave us the best expected RNAi phenotype. The method is easy to perform, reproducible, and easily quantified. Furthermore, our protocol minimizes the use of specialized equipment, so it is suitable for smaller laboratories or those at predominantly undergraduate institutions.

Using RNA-mediated Interference Feeding Strategy to Screen for Genes Involved in Body Size Regulation in the Nematode C. elegans

We demonstrate how to use the RNAi feeding technique to knock down target genes and score body size phenotype in C. elegans. This method could be used for a large scale screen to identify potential genetic components of interest, such as those involved in body size regulation by DBL-1/TGF-&beta; signaling.

Double-strand RNA-mediated interference (RNAi) is an effective strategy to knock down target gene expression1-3. It has been applied to many model systems ...

Culturing of Human Nasal Epithelial Cells at the Air Liquid Interface

In vitro models using human primary epithelial cells are essential in understanding key functions of the respiratory epithelium in the context of microbial infections or inhaled agents. Direct comparisons of cells obtained from diseased populations allow us to characterize different phenotypes and dissect the underlying mechanisms mediating changes in epithelial cell function. Culturing epithelial cells from the human tracheobronchial region has been well documented, but is limited by the availability of human lung tissue or invasiveness associated with obtaining the bronchial brushes biopsies. Nasal epithelial cells are obtained through much less invasive superficial nasal scrape biopsies and subjects can be biopsied multiple times with no significant side effects. Additionally, the nose is the entry point to the respiratory system and therefore one of the first sites to be exposed to any kind of air-borne stressor, such as microbial agents, pollutants, or allergens. 
Briefly, nasal epithelial cells obtained from human volunteers are expanded on coated tissue culture plates, and then transferred onto cell culture inserts. Upon reaching confluency, cells continue to be cultured at the air-liquid interface (ALI), for several weeks, which creates more physiologically relevant conditions. The ALI culture condition uses defined media leading to a differentiated epithelium that exhibits morphological and functional characteristics similar to the human nasal epithelium, with both ciliated and mucus producing cells. Tissue culture inserts with differentiated nasal epithelial cells can be manipulated in a variety of ways depending on the research questions (treatment with pharmacological agents, transduction with lentiviral vectors, exposure to gases, or infection with microbial agents) and analyzed for numerous different endpoints ranging from cellular and molecular pathways, functional changes, morphology, etc. 
In vitro models of differentiated human nasal epithelial cells will enable investigators to address novel and important research questions by using organotypic experimental models that largely mimic the nasal epithelium in vivo.

Nasal epithelial cells, obtained through superficial scrape biopsy of human volunteers, are expanded and transferred onto tissue culture inserts. Upon reaching confluency, cells are grown at air liquid interface, yielding cultures of ciliated and non-ciliated cells. Differentiated nasal epithelial cell cultures provide viable experimental models for studying the respiratory mucosa.

In vitro models using human primary  epithelial cells are essential in understanding key functions of the respiratory  epithelium in the context of ...

Identification of Post-translational Modifications of Plant Protein Complexes

Plants adapt quickly to changing environments due to elaborate perception and signaling systems. During pathogen attack, plants rapidly respond to infection via the recruitment and activation of immune complexes. Activation of immune complexes is associated with post-translational modifications (PTMs) of proteins, such as phosphorylation, glycosylation, or ubiquitination. Understanding how these PTMs are choreographed will lead to a better understanding of how resistance is achieved.
Here we describe a protein purification method for nucleotide-binding leucine-rich repeat (NB-LRR)-interacting proteins and the subsequent identification of their post-translational modifications (PTMs). With small modifications, the protocol can be applied for the purification of other plant protein complexes. The method is based on the expression of an epitope-tagged version of the protein of interest, which is subsequently partially purified by immunoprecipitation and subjected to mass spectrometry for identification of interacting proteins and PTMs.
This protocol demonstrates that: i). Dynamic changes in PTMs such as phosphorylation can be detected by mass spectrometry; ii). It is important to have sufficient quantities of the protein of interest, and this can compensate for the lack of purity of the immunoprecipitate; iii). In order to detect PTMs of a protein of interest, this protein has to be immunoprecipitated to get a sufficient quantity of protein.

We describe here a protocol for the purification and characterization of plant protein complexes. We demonstrate that by immunoprecipitating a single protein within a complex, so we can identify its post-translational modifications and its interacting partners.

Plants adapt quickly to changing environments due to elaborate perception and signaling systems. During pathogen attack, plants rapidly respond to ...

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. ...

Identification of Novel Regulators of Plant Transpiration by Large-Scale Thermal Imaging Screening in Helianthus Annuus

Plant adaptation to biotic and abiotic stresses is governed by a variety of factors, among which the regulation of stomatal aperture in response to water deficit or pathogens plays a crucial role. Identifying small molecules that regulate stomatal movement can therefore contribute to understanding the physiological basis by which plants adapt to their environment. Large-scale screening approaches that have been used to identify regulators of stomatal movement have potential limitations: some rely heavily on the abscisic acid (ABA) hormone signaling pathway, therefore excluding ABA-independent mechanisms, while others rely on the observation of indirect, long-term physiological effects such as plant growth and development. The screening method presented here allows the large-scale treatment of plants with a library of chemicals coupled with a direct quantification of their transpiration by thermal imaging. Since evaporation of water through transpiration results in leaf surface cooling, thermal imaging provides a non-invasive approach to investigate changes in stomatal conductance over time. In this protocol, Helianthus annuus seedlings are grown hydroponically and then treated by root feeding, in which the primary root is cut and dipped into the chemical being tested. Thermal imaging followed by statistical analysis of cotyledonary temperature changes over time allows for the identification of bioactive molecules modulating stomatal aperture. Our proof-of-concept experiments demonstrate that a chemical can be carried from the cut root to the cotyledon of the sunflower seedling within 10 minutes. In addition, when plants are treated with ABA as a positive control, an increase in leaf surface temperature can be detected within minutes. Our method thus allows the efficient and rapid identification of novel molecules regulating stomatal aperture.

Identification of Novel Regulators of Plant Transpiration by Large-Scale Thermal Imaging Screening in Helianthus Annuus

We provide a method for identifying modulators of foliar transpiration by large-scale screening of a compound library.

Plant adaptation to biotic and abiotic stresses is governed by a variety of factors, among which the regulation of stomatal aperture in response to water ...

Detecting Virus and Salivary Proteins of a Leafhopper Vector in the Plant Host

Insect vectors horizontally transmit many plant viruses of agricultural importance. More than one-half of plant viruses are transmitted by hemipteran insects that have piercing-sucking mouthparts. During viral transmission, the insect saliva bridges the virus-vector-host because the saliva vectors viruses, and the insect proteins, trigger or suppress the immune response of plants from insects into plant hosts. The identification and functional analyses of salivary proteins are becoming a new area of focus in the research field of arbovirus-host interactions. This protocol provides a system to detect proteins in the saliva of leafhoppers using the plant host. The leafhopper vector Nephotettix cincticeps infected with rice dwarf virus (RDV) serves as an example. The vitellogenin and major outer capsid protein P8 of RDV vectored by the saliva of N. cincticeps can be detected simultaneously in the rice plant that N. cincticeps feeds on. This method is applicable for testing the salivary proteins that are transiently retained in the plant host after insect feeding. It is believed that this system of detection will benefit the study of hemipteran-virus-plant or hemipteran-plant interactions.

This protocol demonstrates how to use the plant host to detect salivary proteins of leafhopper and plant viral proteins released by leafhopper vectors.

Insect vectors horizontally transmit many plant viruses of agricultural importance. More than one-half of plant viruses are transmitted by hemipteran ...

A Robust Method for the Large-Scale Production of Spheroids for High-Content Screening and Analysis Applications

High-content screening (HCS) and high-content analysis (HCA) are technologies that provide researchers with the ability to extract large-scale quantitative phenotypic measurements from cells. This approach has proved powerful for deepening our understanding of a wide range of both fundamental and applied events in cell biology. To date, the majority of applications for this technology have relied on the use of cells grown in monolayers, although it is increasingly realized that such models do not recapitulate many of the interactions and processes that occur in tissues. As such, there has been an emergence in the development and use of 3-dimensional (3D) cell assemblies, such as spheroids and organoids. Although these 3D models are particularly powerful in the context of cancer biology and drug delivery studies, their production and analysis in a reproducible manner suitable for HCS and HCA present a number of challenges. The protocol detailed here describes a method for the generation of multicellular tumor spheroids (MCTS), and demonstrates that it can be applied to three different cell lines in a manner that is compatible with HCS and HCA. The method facilitates the production of several hundred spheroids per well, providing the specific advantage that when used in a screening regime, data can be obtained from several hundred structures per well, all treated in an identical manner. Examples are also provided, which detail how to process the spheroids for high-resolution fluorescence imaging and how HCA can extract quantitative features at both the spheroid level as well as from individual cells within each spheroid. This protocol could easily be applied to answer a wide range of important questions in cell biology.

This protocol details a method for the production of three different types of spheroids in a manner that makes them suitable for large-scale high-content screening and analysis. In addition, examples are presented showing how they can be analyzed at spheroid and individual cell levels.