We thank Dr. Daniel Bumbarger for the predation behavior picture. This work was funded by the Max-Planck Society.

1. Mouth form Phenotyping
<ol>
	<li>Mouth Form Identification on Agarose Pads 
	Note: In order to visualize the nematode mouth morphs, immobilize the worms with a mild anesthetic treatment on agarose pads and observe as follows.
	<ol>
		<li>Grow and maintain nematode cultures such as P. pacificus on 6 cm standard nematode growth media (NGM) plates and feed on a bacterial lawn of E. coli OP50 19.</li>
		<li>Make agarose pads by first adding 0.06 g agarose to 3 ml H2O in a 15 ml tube to make 3 ml 2% agarose solution. This can be stored for up to a year at 4 &#176;C.</li>
		<li>Mix and melt the agarose thoroughly in a microwave or alternatively use a heat block set to &#62; 88&#176;C.</li>
		<li>Once fully melted, add 10 &#181;l of a 10 % sodium azide solution to the agarose and mix thoroughly. CAUTION: Dry sodium azide is reactive and all forms are toxic.</li>
		<li>Using a 1 ml micropipette, place a drop of no less than 300 &#181;l of the liquid agarose azide mix onto the middle of a standard glass microscope slide.</li>
		<li>Before the agar cools, quickly place a second microscope slide on top of the drop in order to flatten the agarose which forms a pad upon cooling. Repeat for as many pads as is required.</li>
		<li>Just prior to use, peel apart the glass microscope slides by sliding them off one another. Note: If agarose pads are prepared too far in advance they can be excessively dry and may damage the nematodes.</li>
		<li>To transfer worms to the anesthetic agarose pads, place a drop of M9 buffer (2 - 3 &#181;l) onto the center of the pad. Pick 2 - 3 young P. pacificus adults into the drop of M9 before placing a cover slip carefully over the pad. The P. pacificus nematodes will be immobilized in the agarose and ready to visualize.</li>
		<li>Transfer the microscope slide containing the anesthetized worm to an appropriate microscope and observe under 63X Nomarski optics. Categorize morph identities based on the following features: the presence of an additional sub-ventral tooth, enlarged dorsal tooth and wide mouth opening is indicative of a eurystomatous mouth morph animal, while the presence of a single dorsal tooth and narrower mouth opening indicates a stenostomatous animal (Figure 1). 
		Note: In order to maintain the health of the animal, worms should be maintained on the agar pad for no more than 5 min.</li>
		<li>After mouth morph identification, recover either eurystomatous or stenostomatous nematodes as required by removing the cover slip by gently sliding it off the agarose pad. Carefully pick the selected animals from the agarose pad (E. coli OP50 can be used on the pick to help make it stickier) onto fresh NGM plates. Allow recovery from the anesthetic until normal motile behavior has resumed upon which animals are ready for further predatory assays.</li>
	</ol>
	</li>
	<li>Rapid Mouth Phenotyping 
	Note: Alternatively, with more experience, mouth form type can be assayed without the need for any anesthetic treatment via a stereomicroscope with high magnification (150X).
	<ol>
		<li>Place nematodes on standard NGM plates with a bacterial lawn of E. coli OP50 onto the microscope viewing area.</li>
		<li>Detect differences in mouth size and width. Note: At this magnification no tooth like structures are observable therefore mouth morph identification is solely based on wide mouths versus narrow mouths.</li>
	</ol>
	</li>
</ol>
2. Bite Assay
Note: Biting assays permit a detailed predatory behavioral analysis.
<ol>
	<li>Grow and maintain nematode cultures on standard NGM plates (6 cm) and feed on a bacterial lawn of E. coli OP50 19.</li>
	<li>Make assay plates by growing a large quantity of selected prey nematode larvae such as C. elegans or alternatively an appropriate ecologically relevant prey. Note: Adult C. elegans are too large to be suitable prey so it is important to use the larval stage.
	<ol>
		<li>Maintain C. elegans or other potential prey species on standard NGM plates and feed on a bacterial lawn of E. coli OP50 until the population is freshly starved, resulting in an abundance of young L1 larvae. 
		Note: Time for starvation is dependent on numerous environmental and experimental factors, including number of nematodes used to start the culture, the amount of E. coliOP50 added, and the ambient temperature.</li>
	</ol>
	</li>
	<li>Wash four or more freshly starved prey plates with M9 and pass the worm solution through two 20&#8197;&#181;m filters to remove all large animals and any remaining eggs before collection in a 15 ml tube. Only small larvae should remain in the solution.</li>
	<li>To form a larval pellet centrifuge the filtered prey at 377 x g for 1 min.</li>
	<li>Pipette 3&#8197;&#181;l of pure worm pellet onto a 6&#8197;cm NGM plate with no E. coliOP50 present and wait at least 30 min for larvae to spread out sufficiently to generate an assay plate. 
	Note: 3&#8197;&#181;l of pure worm pellet on the standard assay plates contain &#62; 3,000 prey larvae. This is sufficient to generate frequent contact between predators and prey.</li>
	<li>Screen predatory nematodes for the required mouth morph (protocol 1).</li>
	<li>Using standard worm picking techniques and a light stereomicroscope 19, transfer correctly classified predators on to the assay plate. Take care to transfer as little OP50 bacteria as possible to the assay plate when transferring predators in order to minimize bacterial contamination. Wait 15 min to allow the worm to recover from the stress of being transferred and check for wild-type motile behavior to ensure worms have not been damaged from the transfer. 
	Note: There is no need to starve P. pacificus, as they are highly efficient predators of other nematode larvae even while well fed upon bacteria.</li>
	<li>After recovery, observe the predator using a light stereomicroscope for 10 min. With this equipment, observe and characterize distinct feeding events such as biting, characterized by the predator restricting the movement of the prey; killing, where by an opening of the prey cuticle is detected; and feeding, categorized by an observable consumption of the prey innards (Figure 2A, B and Movie 1).</li>
	<li>Repeat the assay by screening and observing a minimum of 10 individual predatory nematodes to ensure accuracy.</li>
</ol>
3. Corpse Assay
Note: Corpse assays facilitate a more rapid quantification of predatory behavior.
<ol>
	<li>Grow and maintain nematode cultures on standard NGM plates and feed on a bacterial lawn of E. coli OP50 19. Generate triplicates of the assay plates mentioned previously (protocol 2.1 - 2.5).</li>
	<li>Screen predatory nematodes for the required mouth morph as described in protocol 1. Using standard worm picking techniques and a light stereomicroscope transfer 5 predatory nematodes with the required mouth morph to each assay plate. Leave the predators together with the prey for 2 hr.</li>
	<li>After 2 hr screen the assay plate for the presence of emptied corpses (Figure 2B and C). Identify corpses by the absence of motility along with obvious morphological defects including leaking innards or missing worm fragments.</li>
</ol>
4. Analysis of Pharyngeal and Tooth Movement
<ol>
	<li>Grow and maintain nematode cultures on standard NGM plates and feed on a bacterial lawn of E. coli OP50 19. Generate assay plates as mentioned previously (protocol 2.1 - 2.5). If standard 6 cm NGM plates do not fit between the objective and the microscope stage, use the lid of small 35 mm petri dishes containing 2 ml NGM as a suitable alternative.</li>
	<li>Screen predatory nematodes for the required mouth morph as described in protocol 1. Using standard worm picking techniques and a light stereomicroscope, transfer a single correctly classified predator on to the assay plate. Wait 15 min to allow the worm to recover from the stress of being transferred.</li>
	<li>Observe predatory animals on a microscope at 40 - 63X Normaski, with a high-speed camera (Movies 2 and 3). Record pharyngeal pumping and tooth movement over 15 sec, at 50 Hz in at least 20 animals to ensure accurate quantification. Replay recorded movies at the desired speed in order to count individual pumps and tooth events. 
	Note: Pumping is observed in the corpus, located at the middle of the pharynx, while tooth movement is detectable in the mouth opening and is only observed from the dorsal tooth.</li>
</ol>

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The authors declare no financial or competing interests.

Nematodes provide a powerful system for understanding neurobiology and behavior with C. elegans thus far being the primary tool. However, numerous nematode species including P. pacificus display behaviors, which are absent or vary in complexities from the model organism C. elegans and therefore raise fascinating questions regarding the evolution and regulation of these behaviors. One such additional behavior found in many other nematode species including P. pacificus is the capacity to supplement their bacterial diet by engaging in predatory feeding 1, 20. We have therefore developed and described a detailed protocol for easy and rapid characterization of these previously unanalyzed predatory behaviors in nematodes.
Firstly, we have provided methods to screen for variations in feeding apparatus within the nematode mouth. The identification of the correct mouth type is an essential first step for successful predation assays as, at least within the Pristionchus genus only eurystomatous animals are capable of predatory feeding. It is best to identify mouth morphs with the &#34;rapid mouth phenotyping&#34; protocol described in protocol 1.2 as this method is much less invasive and therefore it is less likely that predatory behaviors may be perturbed. However, it is recommended to first become familiar with the different mouth structures by identification with anesthetized animals on agar pads (protocol 1.1).
Following identification of the desired mouth morph, we have described two assays for quantifying predatory feeding. These are a rapid, high throughput &#34;corpse assay&#34; (protocol 3) and a more time consuming but more in-depth behavioral analysis through the &#34;bite assay&#34; (protocol 2). Both of these protocols are highly flexible allowing for several modifications in order to optimize the assays depending on the experimental requirements. For bite assays using P. pacificus predators on C. elegans prey, observations of predatory behavioral interactions for a time window of 10 min was sufficient to quantify a significant amount of bites along with other feeding events. For &#34;corpse assays&#34; again utilizing P. pacificus predators on C. elegans prey, 5 predators for 2 hr produced easily quantifiable and consistent corpse numbers allowing for rapid behavioral analysis. However, it should be noted different species of predatory nematode move at different speeds, eat at different rates and generally demonstrate a large diversity in other behaviors 1. Additionally, different prey species may also be eaten at different rates for similar reasons. It is therefore recommended to optimize the assays based upon the nematode species tested both as predators and prey, and also for any differences in environmental conditions. During both &#34;bite&#34; and &#34;corpse&#34; assays it is critical that both prey and predators are healthy, as stressed or injured predators will not kill efficiently. In addition, fresh assay plates are essential as older plates can become dried out which adversely affects the health of the nematodes leading to erroneous assays. It is also hoped that future iterations of these predatory assays will be able to take advantage of recent advances in technology in order to automate much of the analysis as has been accomplished for investigating many behaviors observed in C. elegans21, 22. Currently problems are likely to arise in nematodes such as P. pacificus as they appear much more sensitive to contact, making isolation and immobilization in microfluidic chambers likely to abrogate predatory feeding. Overcoming this may prove challenging but would facilitate individual nematodes to be screened for subtler predatory behaviors.
Finally, we have also provided methods for examining the nematode feeding apparatus itself facilitating comparisons between predatory and bacterial feeding modes by quantifying the tooth and pharyngeal pumping kinetics using a high-speed camera (protocol 4). The quantification of pharyngeal pumping rates in C. elegans has been utilized to monitor feeding for many years 23, however, C. elegans lacks any form of mouth denticle and also lacks predatory behaviors. Through combining the quantification of pharyngeal pumping with that of tooth activity, any innervation of the teeth specific to predation can be also observed. Due to the magnification required to observe the tooth movement the animals often move out of the focal plane, thus it is usually only possible to observe the tooth for short time windows. Additionally, unlike C. elegans, the pharynx of P. pacificus does not continuously pump, rather it engages in spells of pumping and feeding. Therefore, for accurate pharyngeal pumping rates while feeding to be determined, it is important to record 15 sec of continuous feeding.
These methods presented here therefore provide the first framework for investigating predatory behaviors in nematode systems. Moreover, they may also be adaptable for use in investigating other interactions within the nematode ecosystem including the influence of additionally ecologically relevant organisms on predation including microorganisms, fungi and mites. Thus they provide a means to dissect how these predatory behaviors are regulated, how they may have evolved and also their ecological significance.

Nematodes with their small but complex nervous systems have proved powerful tools for understanding many aspects of neurobiology including behavior. Much of this research has focused on the model organism Caenorhabditis elegans in which a wealth of different behaviors have been successfully dissected and analyzed. These include mechanosensory 3, chemotactic 4, thermotactic 5,6 and magnetotactic 7 influencing mating 8,9, learning 10 and feeding behaviors 11. However, other more distantly related nematode species display behaviors which are not observed in the rhabditid C. elegans or alternatively show additional levels of complexity, which raises pertinent questions regarding their evolution and regulation. One such instance of this can be observed in the distantly related diplogastrid nematode Pristionchus pacificus, which displays much more complex feeding behaviors and rhythms than are observed in C. elegans 1. This is despite the two species sharing homologous pharyngeal neurons 12. Coinciding with these additional feeding behaviors, P. pacificus also displays an expanded dietary range, as they are avid predators, capable of supplementing their bacterial diet by also feeding on the larvae of other nematodes. Fortunately, P. pacificus has been developed as a model for comparative and integrative evolutionary biology and therefore many molecular and genetic tools are now available. These include a fully sequenced and annotated genome 13, molecular and genetic tools including transgenes 14 and CRISPR/Cas9 15,16 as well as a detailed and well annotated phylogeny 17 with over 25 closely related species including its newly discovered sister species. In addition, the ecology of numerous Pristionchus species including P. pacificus is well defined with many species having now been described sharing a necromenic association with scarab beetles, a host they frequently share with other nematode species 18. P. pacificus therefore provides an excellent model system with which to dissect the evolution of novel behaviors and their ecological significance.
In order to analyze predatory feeding behaviors in nematode species such as P. pacificus we developed several novel behavioral assays for easy observation and quantification of predatory actions. As P. pacificus displays a dimorphic mouth structure, which strongly influences predatory behavior, identification of the correct morphotype is essential 1,2. The narrow mouthed stenostomatous morph contains a single blunt dorsal tooth and does not engage in any predatory feeding. Alternatively, the wide mouthed eurystomatous morph includes a much larger claw shaped dorsal tooth and an additional opposing sub-ventral tooth, which together operate to efficiently open the cuticle of their prey. The ratio of the predatory eurystomatous to the non-predatory stenostomatous form varies among Pristionchus species and also within P. pacificus, however, the percentage of eurystomatous mouth morph in the P. pacificus wild type strain (PS312) is usually 70 - 90% 2. Additionally, the mouth form ratios can fluctuate depending on differing environmental influences (both known, including starvation and some small molecule signaling as well as unknown factors), thus correct identification and isolation of the predatory eurystomatous mouth form is essential for successful predatory assays.
Alongside the description of the predatory mouth form we have developed a &#34;bite assay&#34; for direct observation and quantification of predatory behaviors including biting, killing and feeding events. Here prey nematodes are isolated through filtering of newly starved cultures and exposed to predatory adult P. pacificus, which are observed together over a short time span. In addition, we have also developed a high throughput &#34;corpse assay&#34; to facilitate rapid screening of predatory behavior through indirect observation of predatory events. This takes advantage of the presence of larval corpses as a tool to screen for predation. Both assays provide easy and highly repeatable methods for observing and measuring predatory behavior in nematode species such as P. pacificus.

<table><tbody><tr><td>Nylon net filters (20 &amp;#181;m)</td><td>Merck Millipore Ltd</td><td>NY2004700</td><td>Used to filter worms just leaving larvae for use as prey.</td></tr><tr><td>PP Funnel for filter (54 mm)</td><td>Duran</td><td>292215003</td><td>Used to filter worms just leaving larvae for use as prey.</td></tr><tr><td>Small petri dish (35/10 mm)</td><td>Greiner Bio-One&amp;nbsp;</td><td>627102</td><td>For imaging on High speed camera</td></tr><tr><td>Zeiss SteREO Discovery V12</td><td></td><td></td><td>For mouth form identificaton</td></tr><tr><td>Axio-Imager A1</td><td></td><td></td><td>For mouth form identificaton</td></tr><tr><td>Glass Slides</td><td>Roth</td><td>H869</td><td></td></tr><tr><td>Cover Slips</td><td>Roth</td><td>657</td><td></td></tr><tr><td>Motion Scope M3 Highspeed camera</td><td>IDT</td><td></td><td>High speed camera</td></tr><tr><td>Video zoom 44 ENG 1/2&quot; 0.5X to 2.4X</td><td>Zeis</td><td>452984-0000-000</td><td>High speed camera zoom</td></tr><tr><td>Nematode Growth Medium (NGM) ingredients:</td><td></td><td></td><td></td></tr><tr><td>Agar</td><td>Roth</td><td>5210.2</td><td>CAS-Nr. 9002-18-0</td></tr><tr><td>Sodium chloride (NaCl)</td><td>Roth</td><td>3957</td><td>CAS-Nr. 7647-14-5</td></tr><tr><td>Bacto Tryptone</td><td>BD</td><td>211699</td><td>Lot 4316614</td></tr><tr><td>Calcium chloride dihydrate (CaCl&lt;sub&gt;2&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>C3306</td><td>CAS-Nr. 10035-04-8</td></tr><tr><td>Cholesterol from lanolin</td><td>Sigma-Aldrich</td><td>F 26732 00050</td><td>CAS-Nr. 57-88-5</td></tr><tr><td>Magnesium sulfate heptahydrate (MgSO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Merck</td><td>1,058,861,000</td><td>CAS-Nr. 10034-99-8</td></tr><tr><td>Potassium dihydrogen phosphate (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>ACROS organics</td><td>271080025</td><td>CAS-Nr. 7778-77-0</td></tr><tr><td>6 cm petri dish</td><td>Greiner Bio-One</td><td>628102</td><td></td></tr><tr><td>3.5 cm petri dish</td><td>Greiner Bio-One</td><td>627102</td><td></td></tr><tr><td>M9 ingredients:</td><td></td><td></td><td></td></tr><tr><td>Potassium dihydrogen phosphate (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>ACROS organics</td><td>271080025</td><td>CAS-Nr. 7778-77-0</td></tr><tr><td>Sodium hydrogen phosphate heptahydrate (NaHPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S9390-500G-D</td><td>CAS-Nr. 7782-85-6</td></tr><tr><td>Sodium chloride (NaCl)</td><td>Roth</td><td>3957</td><td>CAS-Nr. 7647-14-5</td></tr></tbody></table>

assaying predatory feeding behaviors in pristionchus and other nematodes

This protocol provides multiple methods for the analysis and quantification of predatory feeding behaviors in nematodes. Many nematode species including Pristionchus pacificus display complex behaviors, the most striking of which is the predation of other nematode larvae. However, as these behaviors are absent in the model organism Caenorhabditis elegans, they have thus far only recently been described in detail along with the development of reliable behavioral assays 1. These predatory behaviors are dependent upon phenotypically plastic but fixed mouth morphs making the correct identification and categorization of these animals essential. In P. pacificus there are two mouth types, the stenostomatous and eurystomatous morphs 2, with only the wide mouthed eurystomatous containing an extra tooth and being capable of killing other nematode larvae. Through the isolation of an abundance of size matched prey larvae and subsequent exposure to predatory nematodes, assays including both "corpse assays" and "bite assays" on correctly identified mouth morph nematodes are possible. These assays provide a means to rapidly quantify predation success rates and provide a detailed behavioral analysis of individual nematodes engaged in predatory feeding activities. In addition, with the use of a high-speed camera, visualization of changes in pharyngeal activity including tooth and pumping dynamics are also possible.

Following successful identification of the appropriate mouth morph in P. pacificus, clear differences between eurystomatous and stenostomatous animals can be detected (Figure 3) with only the eurystomatous animals engaging in killing behavior. In stenostomatous animals this behavior seems to be suppressed entirely. Furthermore, differences in the tooth activity and pharyngeal pumping of eurystomatous animals on bacteria and prey (Figure 4 and Movies 2 and 3) are also evident. While predatory feeding, the pumping rate is reduced below that observed during bacterial feeding and tooth movement is detected in a one to one ratio with the pharyngeal pumping. This is potentially indicative of key regulatory mechanisms modulating the behavioral response to differing diet.
<img alt="Figure 1" src="/files/ftp_upload/54404/54404fig1.jpg" /> 
Figure 1. P. pacificus has a Mouth Dimorphism which Influences Feeding Behavior. (A) The eurystomatous mouth form is capable of predation and has a wide mouth opening with a large claw shaped dorsal tooth (false-colored red) and (B) a large opposing hook shaped sub-ventral tooth (false-colored blue). (C) The stenostomatous mouth form is only able to feed on bacteria and has a narrower mouth opening with a flint shaped dorsal tooth (false-colored red) and (D) no sub-ventral tooth (*). Normaski images are&#160; 63X and scale bar represents 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/54404/54404fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54404/54404fig2.jpg" /> 
Figure 2. Predation Assays. (A) P. pacificus bites and kills the larvae of other nematodes such as C. elegans. (B) For biting assays, the number of bitesby predators (*) can be observed using a light stereomicroscope and successful killing and feeding events also recorded. Corpses are also clearly visible (circles). (C) For corpse assays, larval carcasses (arrows) can be easily identified compared to living larvae. Scale bar represents 1 mm in B and 150 &#181;m in C. <a href="https://www.jove.com/files/ftp_upload/54404/54404fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54404/54404fig3.jpg" /> 
Figure 3. Results of Bite and Corpse Assays on C. elegans Prey. (A) Biting behavior is only evident in the eurystomatous mouth form with this behavior not displayed in stenostomatous animals. Error bar represents standard deviation of 10 replicates. (B) Coinciding with no biting behavior evident from stenostomatous animals, corpse assays also reveal carcasses only on assay plates of eurystomatous animals. Error bar represents standard deviation of 5 replicates. <a href="https://www.jove.com/files/ftp_upload/54404/54404fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54404/54404fig4.jpg" /> 
Figure 4. Eurystomatous Pumping Rate and Tooth Movement during Predatory Feeding. Tooth movement is only observed while eurystomatous animals are engaged in predatory feeding. This also coincides with a reduction in pharyngeal pumping. Error bar represents standard deviation of 10 replicates. <a href="https://www.jove.com/files/ftp_upload/54404/54404fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/54404/54404movie1.jpg" /> 
<a href="https://www.jove.com/files/ftp_upload/54404/Movie_1_Killing_behaviour.mp4">Movie 1. Observation of Killing Behavior for Bite Assay using a Light Stereomicroscope. (Right click to download).</a> 
<img alt="Movie 2" src="/files/ftp_upload/54404/54404movie2.jpg" /> 
<a href="https://www.jove.com/files/ftp_upload/54404/Movie_2_HS_Camera_Killing.mp4">Movie 2. High-speed Camera Movie of P. pacificus killing C. elegans Larvae. (Right click to download).</a>
<img alt="Movie 3" src="/files/ftp_upload/54404/54404movie3.jpg" /> 
<a href="https://www.jove.com/files/ftp_upload/54404/Movie_3_HS_pumping_and_tooth.mp4">Movie 3. Zoomed in High-speed Camera of Tooth Movement During Predation. (Right click to download).</a>

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Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

Behavioral assays provide powerful tools for understanding neuronal function. Here we present several protocols for quantifying predatory feeding behavior found in the model nematode Pristionchus pacificus and its relatives. Additionally, we provide methods for analyzing predatory feeding adaptations including mouth structures and teeth.

Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

This protocol provides multiple methods for the analysis and quantification of predatory feeding behaviors in nematodes. Many nematode species including ...

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9.7K Views.  Max-Planck Institute for Developmental Biology. Behavioral assays provide powerful tools for understanding neuronal function. Here we present several protocols for quantifying predatory feeding behavior found in the model nematode Pristionchus pacificus and its relatives. Additionally, we provide methods for analyzing predatory feeding adaptations including mouth structures and teeth. 

Video: Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

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.

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Automated Analysis of a Nematode Population-based Chemosensory Preference Assay

The nematode, Caenorhabditis elegans' compact nervous system of only 302 neurons underlies a diverse repertoire of behaviors. To facilitate the dissection of the neural circuits underlying these behaviors, the development of robust and reproducible behavioral assays is necessary. Previous C. elegans behavioral studies have used variations of a "drop test", a "chemotaxis assay", and a "retention assay" to investigate the response of C. elegans to soluble compounds. The method described in this article seeks to combine the complementary strengths of the three aforementioned assays. Briefly, a small circle in the middle of each assay plate is divided into four quadrants with the control and experimental solutions alternately placed. After the addition of the worms, the assay plates are loaded into a behavior chamber where microscope cameras record the worms' encounters with the treated regions. Automated video analysis is then performed and a preference index (PI) value for each video is generated. The video acquisition and automated analysis features of this method minimizes the experimenter's involvement and any associated errors. Furthermore, minute amounts of the experimental compound are used per assay and the behavior chamber's multi-camera setup increases experimental throughput. This method is particularly useful for conducting behavioral screens of genetic mutants and novel chemical compounds. However, this method is not appropriate for studying stimulus gradient navigation due to the close proximity of the control and experimental solution regions. It should also not be used when only a small population of worms is available. While suitable for assaying responses only to soluble compounds in its current form, this method can be easily modified to accommodate multimodal sensory interaction and optogenetic studies. This method can also be adapted to assay the chemosensory responses of other nematode species.

We present a behavior recording setup and protocol that enables automated analysis of the nematode, Caenorhabditis elegans' preference for soluble compounds in a population-based assay. This article describes the construction of a behavior chamber, the behavioral assay protocol, and video analysis software usage.

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Behavior

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

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

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

Genetics

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

Stable isotopic profiling has long permitted sensitive investigations of the metabolic consequences of genetic mutations and/or pharmacologic therapies in cellular and mammalian models. Here, we describe detailed methods to perform stable isotopic profiling of intermediary metabolism and metabolic flux in the nematode, Caenorhabditis elegans. Methods are described for profiling whole worm free amino acids, labeled carbon dioxide, labeled organic acids, and labeled amino acids in animals exposed to stable isotopes either from early development on nematode growth media agar plates or beginning as young adults while exposed to various pharmacologic treatments in liquid culture. Free amino acids are quantified by high performance liquid chromatography (HPLC) in whole worm aliquots extracted in 4% perchloric acid. Universally labeled 13C-glucose or 1,6-13C2-glucose is utilized as the stable isotopic precursor whose labeled carbon is traced by mass spectrometry in carbon dioxide (both atmospheric and dissolved) as well as in metabolites indicative of flux through glycolysis, pyruvate metabolism, and the tricarboxylic acid cycle. Representative results are included to demonstrate effects of isotope exposure time, various bacterial clearing protocols, and alternative worm disruption methods in wild-type nematodes, as well as the relative extent of isotopic incorporation in mitochondrial complex III mutant worms (isp-1(qm150)) relative to wild-type worms. Application of stable isotopic profiling in living nematodes provides a novel capacity to investigate at the whole animal level real-time metabolic alterations that are caused by individual genetic disorders and/or pharmacologic therapies.

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

Stable isotopic profiling by gas chromatography mass spectrometric analysis of intermediary metabolic flux is described in the nematode, Caenorhabditis elegans. Methods are detailed for assessing isotopic enrichment in carbon dioxide, organic acids, and amino acids following isotope exposure either during development on agar plates or during adulthood in liquid culture.

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A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

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We have developed an assay that measures a nematode&#39;s ability to cross an aversive barrier (i.e. copper) then reach a food source over a prolonged period of time. This protocol builds upon previous work by integrating multiple variables in a manner that allows for continued data collection as the organisms shift towards an increasingly starved condition. Moreover, this assay permits an increased sample size so that larger populations of nematodes can be simultaneously evaluated.
Organisms defective for the ability to detect or respond to copper immediately cross the chemical barrier, while wild type nematodes are initially repelled. As wild type worms are increasingly starved, they begin to cross the barrier and reach the food source. We designed this assay to evaluate a mutant that is incapable of responding to diverse environmental cues, including food sensation or detection of aversive chemicals. When evaluated via this protocol, the defective organisms immediately crossed the barrier, but were also incapable of detecting a food source. Hence, these mutants repeatedly cross the chemical barrier despite temporarily reaching a food source. This assay can straightforwardly test populations of worms to evaluate potential pathway defects related to aversion and starvation.

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

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Neuroscience

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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.

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Laboratory Protocol for Genetic Gut Content Analyses of Aquatic Macroinvertebrates Using Group-specific rDNA Primers

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Most common gut content analyses of macroinvertebrates are visual. Requiring intense knowledge about morphological diversity of prey organisms, they miss soft bodied prey and, due to strong comminution of prey, are nearly impossible for some organisms, including amphipods. We provide detailed, novel genetic approaches for macroinvertebrate prey identification in the diet of amphipods.

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

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

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

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

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

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

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

Investigating Collective Behaviors in Nematodes Using Dog Food Agar Medium

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Animals exhibit dynamic collective behaviors, as observed in flocks of birds, schools of fish, and crowds of humans. The collective behaviors of animals have been investigated in the fields of both biology and physics. In the laboratory, researchers have used various model animals such as the fruit fly and zebrafish for approximately a century, but it has remained a major challenge to study large-scale complex collective behavior orchestrated by these genetically tractable model animals. This paper presents a protocol to create an experimental system of collective behaviors in Caenorhabditis elegans. The propagated worms climb on the lid of the Petri plate and show collective swarming behavior. The system also controls worm interactions and behaviors by changing the humidity and light stimulation. This system allows us to examine the mechanisms underlying collective behaviors by changing environmental conditions and examining the effects of individual-level locomotion on collective behaviors using mutants. Thus, the system is useful for future research in both physics and biology.

Author Spotlight: Collective Behavioral Analysis of the Nematode, Caenorhabditis elegans

Here, a system is reported for studying the collective behaviors of nematodes by culturing them in bulk using dog food agar medium. This system allows researchers to propagate large numbers of dauer worms and can be applied to Caenorhabditis elegans and other related species.

Animals exhibit dynamic collective behaviors, as observed in flocks of birds, schools of fish, and crowds of humans. The collective behaviors of animals ...

Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

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