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>

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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. 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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><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&amp;amp;P Scientific</td><td>VP 522-100</td><td></td></tr><tr><td>Low retention pipette tips (200uL)</td><td>LABFORCE</td><td>1159M44</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&amp;amp;P Scientific</td><td>VP110A</td><td></td></tr><tr><td>pinner</td><td>V&amp;amp;P Scientific</td><td>VP381N</td><td></td></tr><tr><td>pinner rinse trays</td><td>V&amp;amp;P Scientific</td><td>VP 421</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

3.4K Views.  University of Toronto. Dit protocol maakt het mogelijk om nieuwe nematiciden te ontdekken om Ditylenchus dipsaci te bestrijden door middel van kweektechniek en chemische schermen met gemiddelde doorvoer. De eenvoud van deze techniek maakt het mogelijk om duizenden verbindingen per week te screenen om verbindingen met nematicidaal potentieel te identificeren. Deze techniek kan nieuwe verbindingen identificeren om plantparasitaire nematoden van gewassen te doden en daarom de wereldwijde voedselvoorziening te vergrote...

Video: Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

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 ...

Environment

High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp.

Root-knot nematodes (genus Meloidogyne) are obligate plant parasites. They are extremely polyphagous and considered one of the most economically important plant parasitic nematodes. The microscopic second-stage juvenile (J2), molted once in the egg, is the infective stage. The J2s hatch from the eggs, move freely in the soil within a film of water, and locate root tips of suitable plant species. After penetrating the plant root, they migrate towards the vascular cylinder where they establish a feeding site and initiate feeding using their stylets. The multicellular feeding site is comprised of several enlarged multinuclear cells called 'giant cells' which are formed from cells that underwent karyokinesis (repeated mitosis) without cytokinesis. Neighboring pericycle cells divide and enlarge in size giving rise to a typical gall or root knot, the characteristic symptom of root-knot nematode infection. Once feeding is initiated, J2s become sedentary and undergo three additional molts to become adults. The adult female lays 150-250 eggs in a gelatinous matrix on or below the surface of the root. From the eggs new infective J2s hatch and start a new cycle. The root-knot nematode life cycle is completed in 4-6 weeks at 26-28&deg;C.
Here we present the traditional protocol to infect plants, grown in pots, with root-knot nematodes and two methods for high-throughput assays. The first high-throughput method is used for plants with small seeds such as tomato while the second is for plants with large seeds such as cowpea and common bean. Large seeds support extended seedling growth with minimal nutrient supplement. The first high throughput assay utilizes seedlings grown in sand in trays while in the second assay plants are grown in pouches in the absence of soil. The seedling growth pouch is made of a 15.5 x 12.5cm paper wick, folded at the top to form a 2-cm-deep trough in which the seed or seedling is placed. The paper wick is contained inside a transparent plastic pouch. These growth pouches allow direct observation of nematode infection symptoms, galling of roots and egg mass production, under the surface of a transparent pouch. Both methods allow the use of the screened plants, after phenotyping, for crossing or seed production. An additional advantage of the use of growth pouches is the small space requirement because pouches are stored in plastic hanging folders arranged in racks.

High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp.

Two distinct methods to screen plants with root-knot nematodes are described. The described approaches include high-throughput screens with nematodes in a nondestructive manner facilitating the use of these plants in breeding programs.

Root-knot nematodes (genus Meloidogyne) are obligate plant parasites. They are extremely polyphagous and considered one of the most economically important ...

Immunology and Infection

Screening Cotton Genotypes for Reniform Nematode Resistance

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium hirsutum) varieties to improve nematode management. Most protocols involve extracting vermiform nematodes or eggs from the cotton root system or potting soil to determine population density or reproduction rate. These approaches are generally time-consuming with a small number of genotypes evaluated. An alternative approach is described here in which the root system is visually examined for nematode infection. The protocol involves inoculating cotton seedling 7 days after planting with vermiform nematodes and determining the number of females attached to the root system 28 days after inoculation. Data are expressed as the number of females per gram of fresh root weight to adjust for variation in root growth. The protocol provides an excellent method for evaluating host-plant resistance associated with the ability of the nematode to establish an infection site; however, resistance that hinders nematode reproduction is not assessed. As with other screening protocols, variation is commonly observed in nematode infection among individual genotypes within and between experiments. Data are presented to illustrate the range of variation observed using the protocol. To adjust for this variation, control genotypes are included in experiments. Nonetheless, the protocol provides a simple and rapid method to evaluate host-plant resistance. The protocol has been successfully used to identify resistant accessions from the G. arboreum germplasm collection and evaluate segregating populations of more than 300 individuals to determine the genetics of resistance. A vegetative propagation method for recovering plants for resistance breeding was also developed. After removal of the root system for nematode evaluation, the vegetative shoot is replanted to allow the development of a new root system. More than 95% of the shoots typically develop a new root system with plants reaching maturity.

Here, a protocol is presented for the rapid non-destructive screening of cotton genotypes for reniform nematode resistance. The protocol involves visually examining the roots of nematode-infected cotton seedlings to determine infection response. The vegetative shoot from each plant is then propagated to recover plants for seed production.

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium ...

Genetics

Establishing Fungal Entomopathogens as Endophytes: Towards Endophytic Biological Control

Beauveria bassiana is a fungal entomopathogen with the ability to colonize plants endophytically. As an endophyte, B. bassiana may play a role in protecting plants from herbivory and disease. This protocol demonstrates two inoculation methods to establish B. bassiana endophytically in the common bean (Phaseolus vulgaris), in preparation for subsequent evaluations of endophytic biological control. Plants are grown from surface-sterilized seeds for two weeks before receiving a B. bassiana treatment of 108 conidia/ml (or water) applied either as a foliar spray or a soil drench. Two weeks later, the plants are harvested and their leaves, stems and roots are sampled to evaluate endophytic fungal colonization. For this, samples are individually surface sterilized, cut into multiple sections, and incubated in potato dextrose agar media for 20 days. The media is inspected every 2-3 days to observe fungal growth associated with plant sections and record the occurrence of B. bassiana to estimate the extent of its endophytic colonization. Analyses of inoculation success compare the occurrence of B. bassiana within a given plant part (i.e. leaves, stems or roots) across treatments and controls. In addition to the inoculation method, the specific outcome of the experiment may depend on the target crop species or variety, the fungal entomopathogen species strain or isolate used, and the plant's growing conditions.

This protocol demonstrates two inoculation methods to introduce the fungal entomopathogen Beauveria bassiana as an endophyte in the common bean (Phaseolus vulgaris), in preparation for subsequent evaluations of endophytic biological control.

 Beauveria bassiana is a fungal entomopathogen with the ability to colonize plants endophytically. As an endophyte, B. bassiana may play a role in ...

Bioengineering

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 ...

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 ...

Culturing and Genetically Manipulating Entomopathogenic Nematodes

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic of their life cycle is the mutualistic association with the bacteria Photorhabdus and Xenorhabdus, respectively. The nematode parasites are able to locate and enter suitable insect hosts, subvert the insect immune response, and multiply efficiently to produce the next generation that will actively hunt new insect prey to infect. Due to the properties of their life cycle, entomopathogenic nematodes are popular biological control agents, which are used in combination with insecticides to control destructive agricultural insect pests. Simultaneously, these parasitic nematodes represent a research tool to analyze nematode pathogenicity and host anti-nematode responses. This research is aided by the recent development of genetic techniques and transcriptomic approaches for understanding the role of nematode secreted molecules during infection. Here, a detailed protocol on maintaining entomopathogenic nematodes and using a gene knockdown procedure is provided. These methodologies further promote the functional characterization of entomopathogenic nematode infection factors.

Entomopathogenic nematodes live in symbiosis with bacteria and together they successfully infect insects by undermining their innate immune system. To promote research on the genetic basis of nematode infection, methods for maintaining and genetically manipulating entomopathogenic nematodes are described.

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic ...

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. ...

In vivo and In vitro Infection of Potato Roots with Plant Parasitic Nematodes for the Assessment of Induced Structural Changes

Soil-dwelling plant parasitic nematodes (PPNs) are important potato pests that cause lesions and/or change plant roots structure, leading to reduced crop fitness and productivity. Research on the cellular and subcellular mechanisms of PPNs infection and development can resort to field plants or seedlings under greenhouse conditions. Field studies are more representative of natural environments but are subjected to the unpredictability of environmental conditions that can heavily influence research outcomes. Greenhouse studies allow higher control over environmental variables and higher safety against contaminants or pathogens. However, in some hosts, genetic diversity becomes an important factor of variability and influences the host-parasite complex response. We have developed in vitro co-cultures of transgenic roots with PPNs as a reliable alternative that occupies less space, requires less time to obtain, and is free from contamination or from host genetic variability. Co-cultures are obtained by introducing aseptic PPNs to host in vitro transgenic roots. They can be maintained indefinitely, which makes them excellent support for keeping collections of reference PPNs. In the present work, a protocol is detailed for the controlled infection of in vivo potato roots with the root lesion nematode and for establishing in vitro co-cultures of potato transgenic roots with the root-knot nematode. The in vitro co-cultures provided a laboratory proxy for the natural potato infection condition and produced nematode life stages irrespective of season or climate conditions. Additionally, the methodology used for structural analysis is detailed using histochemistry and optical microscopy. The acid fuchsin dye is used to follow nematode attack sites on roots, while differential staining with Periodic acid-Schiff (PAS) and toluidine blue O highlights nematode structures in potato internal root tissue.

The protocol describes infection of Solanum tuberosum roots with plant parasitic nematodes under in vivo greenhouse conditions and potato in vitro transgenic roots for histochemical analysis of root structure through optical microscopy.

Soil-dwelling plant parasitic nematodes (PPNs) are important potato pests that cause lesions and/or change plant roots structure, leading to reduced crop ...

Screening of Lentil Fields for Presence of Fusarium Wilt and Root Rot in T&#252;rkiye under Terrestrial Climate

Lentil is an important self-pollinated legume crop plant. Its production is limited by various biotic factors, especially fungal agents causing the wilt and root rot complex. The study aimed to understand the regional epidemiology and etiology of phytopathogenic fungal agents to develop control strategies against soilborne Fusarium spp. This study investigated 83 lentil sowing localities in Yozgat province for wilt, root and crown rot diseases caused by common Fusarium species during 2022 and 2023. Symptomatic lentil plants were collected for fungal isolation and identification. The Fusarium isolates were grouped according to colony morphology and cultured on PDA medium. Moreover, genomic DNAs obtained from Fusarium isolates were analyzed using PCR and compared with other Fusarium isolates registered in the NCBI GenBank. Genetic relationships among Fusarium isolates were determined using the Maximum Parsimony (MP) method in the Mega 11 program. The results, mean incidence and disease severity rate of wilt and root rot diseases in Yozgat province were determined to be 16.9% and 38.6%, respectively. Fusarium isolates were found in 95.4% of the samples. There was 99.5% to 100% nucleotide sequence homogeneity among F. oxysporum, F. culmorum, F. graminearum, F. acuminatum and F. solani isolates, and the most isolated species was F. oxysporum. The MP dendrogram of Fusarium isolates was divided into two main branches, the first branch included all F. solani isolates. The second main branch included other Fusarium species isolated in the present study and in NCBI GenBank. The study suggests periodic local surveys to determine the frequency of Fusarium wilt for suppression in lentils. Timely suppression of Fusarium-based damages is strongly suggested to control the disease and conserve the lentil production system.

A study in Yozgat province found that biotic factors, such as fungal diseases like wilt and root rot, limit lentil production. Fusarium isolates were found in 95.4% of samples, suggesting periodic local surveys and regular monitoring for sustainable technology development and effective control strategies.

Lentil is an important self-pollinated legume crop plant. Its production is limited by various biotic factors, especially fungal agents causing the wilt ...

Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

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