This research was supported by research grants from the National Institutes of Health to JHR (#R01 5R01HD081266 and #R01GM141493). Some strains were provided by the CGC, which is funded by the NIH Office of Research Infrastructure Programs (P40 OD010440). We would like to acknowledge Pradeep Joshi (UCSB) for his editorial input. Statistical consultation provided by the UCSB DATALAB.

The strains used in the present study are C. elegans (N2) and C. briggsae (AF16) (see Table of Materials). A mixed-sex population of dauers was used for each assay.
1. Chamber preparation
<ol>
	<li>Work in a fume hood. Set up the workspace with a Bunsen burner, 1-2 razorblades, pliers, tweezers, and a plastic cutting surface (see Table of Materials).</li>
	<li>For each chamber, gather two 5 mL serological pipettes. Remove the cotton plug from one pipette with tweezers. Hold a razorblade over the Bunsen burner until hot using pliers. Use the heated blade to slice off the tapered end of the second pipette so that the entire pipette has a uniform diameter (Figure 1A).</li>
	<li>Working quickly, bring the two modified pipette ends close to the flame, melting them slightly. Join these ends by pressing them firmly together, ensuring that the walls of both pipettes are continuous. If any gaps are visible after joining, break them apart and repeat this step (Figure 1A,B).</li>
</ol>
2. Filling the chambers with agar
<ol>
	<li>Prepare NGM with 4% agar according to standard worm procedures21. While the agar is still molten, fill each chamber by attaching it to a serological pipettor and slowly drawing up the solution. Seal the tip with paraffin film before removing the chamber from the pipettor. Lay the chamber flat (parallel to the benchtop) to cool. 
	&#8203;NOTE: Covering the flask with cling-wrap (see Table of Materials) after autoclaving helps prevent bubbles from forming in the solution18.
	<ol>
		<li>To minimize any variations in the consistency of the agar due to uneven cooling, hold the pipettor (see Table of Materials) parallel to the benchtop while drawing up the agar. Lay the chambers flat on the benchtop and allow the agar to harden before moving. 
		NOTE: The agar percentage and media content can be tailored to the experiment. 4% agar is stiffer than the roughly 2% concentration used for plate pouring and prevents worms from burrowing through the medium in our hands. However, other experimenters have observed burrowing behavior by adult worms in up to 9% agar18,19.</li>
	</ol>
	</li>
	<li>Once cooled, heat a 3 mm hex key (or another metal tool of the same size, see Table of Materials) over a Bunsen burner. Firmly press it into the wall of each chamber about 5 mm to one side of the midline, creating a small opening in the plastic (Figure 1C). Use a heated blade to remove the cotton and tapered ends of the chamber and seal these ends with paraffin film. 
	&#8203;NOTE: Once prepared, each chamber must be used within 1 day or kept at 4 &#176;C for several days. Cover any exposed openings with paraffin film to prevent the agar from drying out.</li>
</ol>
3. Isolating dauer larvae
<ol>
	<li>10-15 days prior to the experiment, chunk each strain onto 2-3 large NGM plates with OP50 bacteria and wrap with paraffin film. After 10-15 days, each plate must be fully starved with thousands of dauer larva present on the agar, walls, and lids. 
	NOTE: Make plates ahead of time using a thick OP50 recipe for maximum crowding (see Table of Materials). Dauer formation can also be induced through other methods, including the addition of pheromones or by growing at higher temperatures22.</li>
	<li>Collect worms by rinsing the lids and plates with M9 buffer and pipetting the solution into 15 mL centrifuge tubes. Spin at 1,600 x g for 30-60 s at room temperature and aspirate most of the M9 buffer with a pipette or vacuum aspirator.</li>
	<li>Isolate dauers by treating with 1% SDS followed by a 30% sucrose flotation gradient following the previously published report20.
	<ol>
		<li>Add 7 mL 1% SDS solution to each worm pellet. Leave the worms in SDS for 30 min; rotate the tubes continuously during this time to allow for aeration. Rinse 3-5x with M9 to remove the detergent.</li>
		<li>Next, add 5 mL M9 followed by 5 mL of cold, filtered 60% sucrose solution. Mix thoroughly, and then centrifuge at 1,600 x g for 5 min at room temperature to create a separation gradient.</li>
	</ol>
	</li>
	<li>Fill new 15 mL centrifuge tubes with 2 mL of M9 buffer. Crush the ends of glass Pasteur pipettes to widen the bore and use these pipettes to transfer the top layer of the solution (containing isolated dauers) from the sucrose gradient to the new tubes. Rinse the dauers 3-5x with M9. 
	&#8203;NOTE: There will be thousands of isolated dauers in solution at this point. They may be used immediately or kept aerated by rotating in 5-7 mL M9 overnight.</li>
</ol>
4. Adding dauers to the chamber
<ol>
	<li>Centrifuge dauers at 1,600 x g for 5 min at room temperature and aspirate most of the M9 solution with a pipette or vacuum aspirator. To estimate worm density, manually count the number of worms in three separate 1 &#181;L droplets under a dissecting microscope. Use the average of these counts to approximate the number of worms per &#181;L. 
	NOTE: Some small volume pipettors may be unable to reach the bottom of a 15 mL centrifuge tube; in this case, a concentrated droplet of worms can be transferred first to a piece of paraffin film.</li>
	<li>Cut the end of a 10, 20, or 200 &#181;L pipette tip to widen the bore. Set the micropipette to a slightly larger volume than the intended aspiration volume. Aspirate approximately 1-2 &#181;L of the concentrated worm solution (ideally between 100-300 worms) and allow a small amount of air to enter the tip. 
	NOTE: Large numbers of worms (1,000+) in one chamber may increase the range of distances traveled due to overcrowding9.</li>
	<li>Gently push the pipette tip into the agar while depressing the micropipette. This will create an injection site in the agar without clogging the tip. Release the worms into the agar. Use paraffin film to seal the opening.</li>
</ol>
5. Running and scoring the assay
NOTE: Gravitaxis can be tested under various conditions that may affect behavior9.
<ol>
	<li>To eliminate as many variables as possible, place the chambers within a dark Faraday cage (see Table of Materials) at room temperature.</li>
	<li>Label each chamber and hang it vertically within the Faraday cage (perform this using labeling tape). Test horizontal controls concurrently by laying some chambers flat within the same environmental conditions. Test the experimental strains against vertically oriented N2 worms as a positive control. Use horizontally oriented chambers containing N2 dauers as an additional negative control. 
	NOTE: Horizontal and vertical assays must be performed in the same setting within the lab to minimize environmental variability between the two conditions. A large incubator may be used to house both Faraday cages.</li>
	<li>Dauers begin to disperse from the start site after a few hours and take several hours to reach either end of the chamber. Leave the chambers undisturbed during this time and score within 12-24 h following injection. 
	NOTE: Do not use a paralytic (such as sodium azide) to immobilize worms that have reached the ends of the chambers. Because the overall distribution of worms is being measured instead of a preference index (as in a two-choice assay), sodium azide is unnecessary and may alter the results.</li>
	<li>Remove and score gravitaxis chambers one at a time. Look for live dauers under a dissecting microscope and mark their locations in ink (Figure 1C,D). Do not score worms within 2.5 cm to either side of the injection site, as these worms are not likely to be demonstrating a directional preference.
	<ol>
		<li>Also, avoid scoring worms that appear dead or are trapped in the liquid. Discard any chambers containing &#62;50% dead or swimming worms. 
		NOTE: Once the chamber has been removed from the testing area, it needs to be scored promptly for accuracy. Dauers must only be visible between the agar&#39;s surface and the pipette&#39;s wall. If burrowing behavior is observed, consider increasing agar concentration in future assays.</li>
	</ol>
	</li>
	<li>To quantify the results, use a marker to divide each half of the chamber into seven 3.5 cm sections starting 2.5 cm away from the injection site (on some pipettes, 3.5 cm = 1 mL volume). Use a manual tally counter (see Table of Materials) to tally the number of worms observed in each section. 
	NOTE: There should be seven sections on each side of the origin, which can be numbered from -7 (bottom) to +7 (top). These numbers must correspond to the same relative locations based on how the chambers were constructed; agar is drawn from the -7 end to the +7 end in step 2.</li>
</ol>

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The authors declare no competing interests.

Comparison with prior methods 
Unlike chemotaxis, gravitaxis in Caenorhabditis cannot be reliably observed using a traditional agar plate experimental design. A standard Petri dish is 150 mm in diameter, resulting in only 75 mm available in either direction for dauers to demonstrate gravitaxis preference. Although C. elegans&#39; orientational preference can be assayed in solution12, this method is low throughput as worms must be analyzed one at a time. Additionally, gravitactic preferences and behaviors may differ between worms floating in media versus crawling on a surface. For these reasons, we developed a high throughput assay with enhanced sensitivity that can be used to analyze gravitaxis behavior in crawling or climbing worms. Because C. elegans are sensitive to light and electromagnetic fields, both of which interfere with gravitactic preference9, these assays need to be performed without either stimulus. If a homemade Faraday cage is being used, checking the cage&#39;s strength and reinforcing it are necessary and recommended.
The assay chambers described in this protocol measure approximately 54 cm in length and are placed within Faraday cages to shield against light and electromagnetic fields. It was found that changing the &#34;arena&#34; for observing gravitaxis behavior has several advantages. First, it allows for detailed quantification and descriptive analysis. The relative distances traveled, and overall distributions of worms can be collected and compared instead of counting the number of negatively versus positively gravitactic worms. Second, the enclosed environment of an agar-filled pipette more closely replicates the conditions that may promote gravitaxis in the wild. As stated above, the dauer stage is a dispersal phase that allows escaping from unfavorable conditions6. Because Caenorhabditis live in compost, where guiding cues such as light may not be available, gravity may enable worms to navigate to the surface6,9. Two-dimensional plate-based assays are unlikely to replicate these conditions even if they are tested without other stimuli. Finally, this larger apparatus tests larger quantities of worms in a single assay.
Limitations and other considerations 
While this protocol effectively measures gravitactic preference in large numbers of dauers, it is impractical for small or single worm experiments due to the time and materials required to construct each chamber. By combining counts across trials rather than using a gravitaxis index, statistical power is increased because each worm is treated as an independent event. While this enhanced sensitivity is useful in differentiating gravitactic and non-gravitactic worms, it is important to note that C. elegans dauers exhibit social behaviors7,24 that could affect overall gravitaxis. We found that higher worm densities are correlated with a greater range in the distance traveled over the large-scale assay, though this effect is minimal when the total count is less than approximately 1,000 worms9. Interaction effects resulting from factors such as the total worms in each chamber should be monitored when analyzing the data. As with all behavioral assays, scoring should be blinded when possible.
Finding an average density of worms in the solution can minimize variability in the number of worms injected from chamber to chamber. Worms quickly settle in solution, so mixing the worms before pipetting, either by flicking the tubes or pipetting up and down with a wide-bore tip is important. Even when a consistent density is achieved, the number of worms added may vary because worms are likely to stick to the inside surface of the plastic pipette tips. Coating pipette tips with BSA or other solutions could minimize this effect. Worms must be injected with as little solution as possible; if too much liquid is added to the chamber, worms will become trapped in the solution and may even drift. For this reason, only live, crawling worms may be scored, and any chamber containing more than 50% dead or swimming worms must not be used. Chambers are to be removed from the Faraday cage one at a time and scored within 10-15 min for the greatest accuracy.
Potential applications 
This assay may be used to test the environmental, genetic, and cellular requirements for gravitaxis in a variety of Caenorhabditis strains. So far, light and electromagnetic fields have been identified as stimuli that interfere with gravitaxis9. However, C. elegans is sensitive to other stimuli, including temperature, volatile and non-volatile chemicals, texture, humidity, and sound, which influence their behavior25,26,27,28. Understanding how various sensory inputs are integrated within the nervous system is an outstanding question in neuroscience, particularly when integration occurs at the level of individual neurons9,29,30,31,32. Sensory integration is especially important in proprioception, which is itself an integrative modality that draws from multiple sensory cues1.
Understanding gravity sensation in Caenorhabditis has implications for human health. Millions of individuals suffer from vestibular dysfunction in the United States alone33, and many of these disorders have underlying genetic causes34,35. For this reason, the identification of gene candidates and targeted therapies is an active area of research36. Additionally, because vertebrate vestibular and auditory systems are closely linked developmentally and evolutionarily33,36,37, elucidating gravity sensation could also provide insight into hearing and hearing disorders.

Sensing Earth&#39;s gravitational pull is crucial for many organisms&#39; orientation, movement, coordination, and balance. However, the molecular mechanisms and neurocircuitry of gravity sensation are poorly understood compared with other senses. In animals, gravity sensation interacts with and can be outcompeted by other stimuli to influence behavior. Visual cues, proprioceptive feedback, and vestibular information can be integrated to generate a sense of body awareness relative to an animal&#39;s surroundings1,2. Conversely, gravitactic preference can be altered in the presence of other stimuli3,4,5. Therefore, gravitactic behavior is ideal for studying gravity sensation and understanding the nervous system&#39;s complex sensory integration and decision-making.
C. elegans is an especially useful model organism for studying gravitaxis because of its polyphenic lifecycle. When exposed to stressors during development, including heat, overcrowding, or a lack of food, C. elegans larvae develop into dauers, which are highly stress-resistant6. As dauers, worms perform characteristic behaviors, such as nictation, in which worms &#34;stand&#34; on their tails and wave their heads, that may facilitate dispersal to better habitats7. Gravitaxis assays of C. elegans and C. japonica suggest that dauer larvae negatively gravitax, and that this behavior is more readily observed in dauers than in adults8,9. Testing gravitaxis in other Caenorhabditis strains may reveal natural variation in gravitactic behavior.
Mechanisms for gravity sensation have been characterized in Euglena, Drosophila, Ciona, and various other species using gravitaxis assays3,10,11. Meanwhile, gravitaxis studies in Caenorhabditis initially provided mixed results. A study of C. elegans orientational preference found that worms orient with their heads down in solution, suggesting positive gravitactic preference12. Meanwhile, although C. japonica dauers were identified early on as being negatively gravitactic8, this behavior has only recently been described in C. elegans9. Several challenges arise in developing a representative gravitaxis assay in worms. Caenorhabditis strains are maintained on agar plates; for this reason, behavioral assays typically use agar plates as part of their experimental design13,14,15. The earliest reported gravitaxis assay in Caenorhabditis was performed by standing a plate on its side at a 90&#176; angle to the horizontal control plate8. However, gravitaxis behavior is not always robust under these conditions. While adult worms can be assayed for orientational preference in solution12, this directional preference may also be context-dependent, leading to different behaviors if the worms are crawling rather than swimming. Additionally, C. elegans is sensitive to other stimuli, including light and electromagnetic fields16,17, which interfere with their responses to gravity9. Therefore, an updated gravitaxis assay that shields against other environmental variables is important for dissecting the mechanisms of this sensory process.
In the present protocol, an assay for observing Caenorhabditis gravitaxis is described. The setup for this study is based in part on a method developed to study neuromuscular integrity18,19. Dauer larvae are cultured and isolated using standard procedures20. They are then injected into chambers made from two 5 mL serological pipettes filled with agar. These chambers can be oriented vertically or horizontally and placed within a dark Faraday cage for 12-24 h to shield against light and electromagnetic fields. The location of each worm in the chambers is recorded and compared with the vertical taxis of a reference strain such as C. elegans N2.

<table><tbody><tr><td>1% Sodium Dodecyl Sulfate solution</td><td></td><td></td><td>From stock 10% (w/v) SDS in DI water</td></tr><tr><td>15 mL Centrifuge tubes</td><td>Falcon</td><td>14-959-53A</td><td></td></tr><tr><td>3 mm Hex key</td><td></td><td></td><td>Other similar sized metal tools may be used</td></tr><tr><td>4% Agar in Normal Growth Medium (NGM) - 1 L</td><td></td><td></td><td>Prior to autoclaving: 3 g NaCl, 40 g Agar, 2.5 g Peptone, 2 g Dextrose, 10 mL Uracil (2 mg/mL), 500 &amp;mu;L Cholesterol (10 mg/mL), 1 mL CaCl2, 962 mL DI water; After autoclaving: 24.5 mL Phosphate Buffer, 1 mL 1 MgSO4 (1 M), 1 mL Streptomycin (200 mg/mL)</td></tr><tr><td>5 mL Serological pipettes</td><td>Fisherbrand</td><td>S68228C</td><td>Polystyrene, not borosilicate glass</td></tr><tr><td>60% Cold sucrose solution</td><td></td><td></td><td>60% sucrose (w/v) in DI water; sterilize by filtration (0.45 &amp;mu;m filter). Keep at 4 &amp;deg;C</td></tr><tr><td>AF16 &lt;em&gt;C. briggsae&lt;/em&gt; or other experimental strain</td><td></td><td></td><td>Available from the CGC (Caenorhabditis Genetics Center)</td></tr><tr><td>Bunsen burner</td><td></td><td></td><td></td></tr><tr><td>Cling-wrap</td><td>Fisherbrand</td><td>22-305654</td><td></td></tr><tr><td>Clinical centrifuge</td><td></td><td></td><td></td></tr><tr><td>Disposable razor blades</td><td>Fisherbrand</td><td>12-640</td><td></td></tr><tr><td>Faraday cage</td><td></td><td></td><td>Can be constructed using cardboard and aluminum foil; 30&quot; L x 6&quot; W x 26&quot; H or larger</td></tr><tr><td>Ink markers</td><td></td><td></td><td>Sharpie or other brand for marking on plastic</td></tr><tr><td>Labeling tape</td><td>Carolina</td><td>215620</td><td></td></tr><tr><td>M9 buffer</td><td></td><td></td><td>22 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 42 mM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 86 mM NaCl</td></tr><tr><td>N2 &lt;em&gt;C. elegans&lt;/em&gt; strain</td><td></td><td></td><td>Available from the CGC (Caenorhabditis Genetics Center)</td></tr><tr><td>NGM plates with OP50</td><td></td><td></td><td>1.7% (w/v) agar in NGM (see description: 4% agar in NGM). Seed with OP50</td></tr><tr><td>Paraffin film</td><td>Bemis</td><td>13-374-10</td><td></td></tr><tr><td>Plastic cutting board</td><td></td><td></td><td></td></tr><tr><td>Pliers</td><td></td><td></td><td></td></tr><tr><td>Rotating vertical mixer</td><td>BTLab SYSTEMS</td><td>BT913</td><td>With 22 x 15 mL tube bar</td></tr><tr><td>Serological pipettor</td><td>Corning</td><td>357469</td><td></td></tr><tr><td>Stereo Microscope</td><td>Laxco</td><td>S2103LS100</td><td></td></tr><tr><td>Tally counter</td><td>ULINE</td><td>H-7350</td><td></td></tr><tr><td>Thick NGM/agar plate media - 1 L</td><td></td><td></td><td>See 4% Agar in NGM recipe; replace 40 g Agar with 20 g Agar</td></tr><tr><td>Tweezers</td><td></td><td></td><td></td></tr></tbody></table>

large-scale gravitaxis assay of caenorhabditis dauer larvae

Gravity sensation is an important and relatively understudied process. Sensing gravity enables animals to navigate their surroundings and facilitates movement. Additionally, gravity sensation, which occurs in the mammalian inner ear, is closely related to hearing - thus, understanding this process has implications for auditory and vestibular research. Gravitaxis assays exist for some model organisms, including Drosophila. Single worms have previously been assayed for their orientation preference as they settle in solution. However, a reliable and robust assay for Caenorhabditis gravitaxis has not been described. The present protocol outlines a procedure for performing gravitaxis assays that can be used to test hundreds of Caenorhabditis dauers at a time. This large-scale, long-distance assay allows for detailed data collection, revealing phenotypes that may be missed on a standard plate-based assay. Dauer movement along the vertical axis is compared with horizontal controls to ensure that directional bias is due to gravity. Gravitactic preference can then be compared between strains or experimental conditions. This method can determine molecular, cellular, and environmental requirements for gravitaxis in worms.

Comparing gravitaxis across species 
Following the procedure outlined above, C. briggsae dauer gravitaxis can be compared with C. elegans gravitaxis and horizontal controls. The vertical distribution (maroon) of C. briggsae dauers is skewed toward the tops of the chambers, with a large percentage of worms reaching +7 (Figure 2A). Contrasted with horizontal controls (aqua), in which dauers are distributed in a roughly bell-shaped curve around the center of the chambers, this trend indicates negative gravitactic behavior. These data can be compared with C. elegans gravitaxis performed on the same experimental days (Figure 2B).
A Kruskal-Wallis test followed by Dunn&#39;s test with Bonferroni correction for multiple comparisons determined any significant differences between assays9,23. In this experiment, 1,108 C. briggsae dauers were scored across three vertical chambers from two independent experimental days. These worms migrated significantly upward than horizontal controls (p &#60; 0.001; 1,639 dauers in three horizontal chambers over 2 experimental days). Moreover, the vertical C. briggsae and vertical C. elegans distributions did not differ (p &#62; 0.05; 386 C. elegans dauers in two vertical chambers over 2 experimental days). These results suggest that C. briggsae dauers show a negative gravitaxis behavior similar to that of C. elegans dauers.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64062/64062fig01.jpg" /> 
Figure 1: Gravitaxis assay chamber setup. Photos depicting steps of gravitaxis assay chamber preparation. (A) Individual 5 mL pipettes were prepared for fusion into a single chamber. Removal of cotton plug indicated with black arrowhead; removal of tip indicated with a white arrowhead. The tubes used in this study have an inner diameter of 6 mm and are each 34.5 cm long (prior to cutting). (B) Completed chamber prior to addition of NGM agar. Pipettes have been fused by first melting over a Bunsen burner. (C) Example chambers filled with NGM agar. The injection site is indicated with an arrow and enlarged in the inset. Worms have been marked in ink, and lines are drawn to distinguish distances along with the chamber as well as to facilitate scoring. (D) View of chambers in their entirety (taken after scoring). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64062/64062fig01large.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/64062/64062fig02.jpg" /> 
Figure 2: Gravitaxis in C. briggsae vs. C. elegans. Results of gravitaxis in C. briggsae and C. elegans. (A) Histogram of C. briggsae gravitaxis. Movement in vertical chambers (maroon) is compared with horizontal controls (aqua). Distance from the injection point (-7 being the furthest below, +7 furthest above) is plotted as the percent of total worms assayed (x-axis). No data are plotted for the origin (+0) as worms are not counted within 2.5 cm to either side of the injection site. N = 1,639 worms across three tubes in the horizontal condition; vertical N = 1,108 worms across three tubes. (B) Boxplots comparing the distribution of C. briggsae horizontal and vertical worms versus C. elegans vertical worms. C. briggsae sample sizes are given above. C. elegans vertical N = 386 worms across two tubes. Means are indicated with white diamonds; vertical conditions are in maroon and are labeled &#34;V&#34;, and horizontal conditions (&#34;H&#34;) are in aqua. Comparisons were made using the Kruskal-Wallis test, followed by the Dunn&#39;s test with Bonferroni correction for multiple comparisons. No stars = p &#62; 0.05, **** = p &#60; 0.0001. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64062/64062fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae at JoVE.com

Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae

The present protocol outlines methods for conducting a large-scale gravitaxis assay with Caenorhabditis dauer larvae. This protocol allows for better detection of gravitaxis behavior compared with a plate-based assay.

Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae

Gravity sensation is an important and relatively understudied process. Sensing gravity enables animals to navigate their surroundings and facilitates ...

large-scale-gravitaxis-assay-of-caenorhabditis-dauer-larvae

Nanyang Technological University

Research

JoVE Journal

Behavior

2.2K Views.  University of California. The present protocol outlines methods for conducting a large-scale gravitaxis assay with Caenorhabditis dauer larvae. This protocol allows for better detection of gravitaxis behavior compared with a plate-based assay.

Video: Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages for identifying biologically active chemical structures that can modify complex phenotypes such as lifespan. Described here is a simple protocol for producing hundreds of 96-well culture plates with fairly consistent numbers of C. elegans in each well. Next, we specified how to use these cultures to screen thousands of chemicals for effects on the lifespan of the nematode C. elegans. This protocol makes use of temperature sensitive sterile strains, agar plate conditions, and simple animal handling to facilitate the rapid and high throughput production of synchronized animal cultures for screening.

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Here we describe a simple protocol for rapidly producing hundreds of nematode growth media agar, 96-well culture plates with consistent numbers of Caenorhhabditis elegans per well. These cultures are useful for the phenotypic screening of whole organisms. We focus here on using these cultures to screen chemicals for pro-longevity effects.

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages ...

Biochemistry

Measuring Caenorhabditis elegans Life Span on Solid Media

Aging is a degenerative process characterized by a progressive deterioration of cellular components and organelles resulting in mortality. The nematode Caenorhabditis elegans has emerged as a principal model used to study the biology of aging. Because virtually every biological subsystem undergoes functional decline with increasing age, life span is the primary endpoint of interest when considering total rate of aging. In nematodes, life span is typically defined as the number of days an animal remains responsive to external stimuli. Nematodes can be propagated either in liquid media or on solid media in plates, and techniques have been developed for measuring life span under both conditions. Here we present a generalized protocol for measuring life span of nematodes maintained on solid nematode growth media and fed a diet of UV-killed bacteria. These procedures can easily be adapted to assay life span under various common conditions, including a diet consisting of live bacteria, dietary restriction, and RNA interference.

Measuring Caenorhabditis elegans Life Span on Solid Media

In this article we present a general protocol for measuring life span of nematodes maintained on solid media with UV-killed bacterial food.

Aging is a degenerative process characterized by a progressive deterioration of cellular components and organelles resulting in mortality. The nematode ...

Biology

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

Caenorhabditis elegans (C. elegans) has been and remains a valuable model organism to study developmental biology, aging, neurobiology, and genetics. The large body of work on C. elegans makes it an ideal candidate to integrate into large-population, whole-animal studies to dissect the complex biological components and their relationships with another organism. In order to use C. elegans in collaborative -omics research, a method is needed to generate large populations of animals where a single sample can be split and assayed across diverse platforms for comparative analyses.
Here, a method to culture and collect an abundant mixed-stage C. elegans population on a large-scale culture plate (LSCP) and subsequent phenotypic data is presented. This pipeline yields sufficient numbers of animals to collect phenotypic and population data, along with any data needed for -omics experiments (i.e., genomics, transcriptomics, proteomics, and metabolomics). In addition, the LSCP method requires minimal manipulation to the animals themselves, less user preparation time, provides tight environmental control, and ensures that handling of each sample is consistent throughout the study for overall reproducibility. Lastly, methods to document population size and population distribution of C. elegans life stages in a given LSCP are presented.

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

To use Caenorhabditis elegans (C. elegans) in omics research, a method is needed to generate large populations of worms where a single sample can be measured across platforms for comparative analyses. Here, a method to culture C. elegans populations on large-scale culture plates (LSCPs) and to document population growth is presented.

Caenorhabditis elegans (C. elegans) has been and remains a valuable model organism to study developmental biology, aging, neurobiology, and genetics. The ...

Genetics

Surveying Low-Cost Methods to Measure Lifespan and Healthspan in Caenorhabditis elegans

The discovery and development of Caenorhabditis elegans as a model organism was influential in biology, particularly in the field of aging. Many historical and contemporary studies have identified thousands of lifespan-altering paradigms, including genetic mutations, transgenic gene expression, and hormesis, a beneficial, low-grade exposure to stress. With its many advantages, including a short lifespan, easy and low-cost maintenance, and fully sequenced genome with homology to almost two-thirds of all human genes, C. elegans has quickly been adopted as an outstanding model for stress and aging biology. Here, several standardized methods are surveyed for measuring lifespan and healthspan that can be easily adapted into almost any research environment, especially those with limited equipment and funds. The incredible utility of C. elegans is featured, highlighting the capacity to perform powerful genetic analyses in aging biology without the necessity of extensive infrastructure. Finally, the limitations of each analysis and alternative approaches are discussed for consideration.

Surveying Low-Cost Methods to Measure Lifespan and Healthspan in Caenorhabditis elegans

Caenorhabditis elegans serve as an excellent model system with robust and low-cost methods for surveying healthspan, lifespan, and resilience to stress.

The discovery and development of Caenorhabditis elegans as a model organism was influential in biology, particularly in the field of aging. Many ...

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

Caenorhabditis elegans (C. elegans) is a well-established model organism used across a range of basic and biomedical research. Within the nematode research community, there is a need for an affordable and effective way to maintain large, age-matched populations of C. elegans. Here, we present a methodology for mechanically sorting and cleaning C. elegans. Our aim is to provide a cost-effective, efficient, fast, and simple process to obtain animals of uniform sizes and life stages for their use in experiments. This tool, the Caenorhabditis Sieve, uses a custom-built lid system that threads onto common conical lab tubes and sorts C. elegans based on body size. We also demonstrate that the Caenorhabditis Sieve effectively transfers animals from one culture plate to another allowing for a rapid sorting, synchronizing, and cleaning without impacting markers of health, including motility and stress-inducible gene reporters. This accessible and innovative tool is a fast, efficient, and non-stressful option for maintaining C. elegans populations.

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

The current protocol includes a methodology for the sorting and cleaning of age-matched populations of Caenorhabditis elegans. It uses a simple, inexpensive, and efficient custom-made tool to obtain a large experimental population of nematodes for research.

Caenorhabditis elegans (C. elegans) is a well-established model organism used across a range of basic and biomedical research. Within the nematode ...

Developmental Biology

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

Neuroscience

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their locomotory patterns upon detection of diverse environmental stimuli and can modulate their suite of behavioral responses in response to starvation conditions. Nematodes typically exhibit a decreased aversive response when removed from a food source for over 30 min. Observation of behavioral changes in response to a changing nutritional status can provide insight into the mechanisms that regulate the transition from a well-fed to starved state.
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

Here, we present a Caenorhabditis elegans-specific assay designed to evaluate changes in copper aversion behavior and the ability to locate a common food source, as the organism progresses from a well-fed to starved nutritional state.

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their ...

C. elegans Chemotaxis Assay

Many organisms use chemotaxis to seek out food sources, avoid noxious substances, and find mates. Caenorhabditis elegans has impressive chemotaxis behavior.
 The premise behind testing the response of the worms to an odorant is to place them in an area and observe the movement evoked in response to an odorant. Even with the many available assays, optimizing worm starting location relative to both the control and test areas, while minimizing the interaction of worms with each other, while maintaining a significant sample size remains a work in progress 1-10. The method described here aims to address these issues by modifying the assay developed by Bargmann et al.1. A Petri dish is divided into four quadrants, two opposite quadrants marked "Test" and two are designated "Control". Anesthetic is placed in all test and control sites. The worms are placed in the center of the plate with a circle marked around the origin to ensure that non-motile worms will be ignored. Utilizing a four-quadrant system rather than one 2 or two 1 eliminates bias in the movement of the worms, as they are equidistant from test and control samples, regardless of which side of the origin they began. This circumvents the problem of worms being forced to travel through a cluster of other worms to respond to an odorant, which can delay worms or force them to take a more circuitous route, yielding an incorrect interpretation of their intended path. This method also shows practical advantages by having a larger sample size and allowing the researcher to run the assay unattended and score the worms once the allotted time has expired.

C. elegans Chemotaxis Assay

A method of quantitatively evaluating the chemotactic response of Caenorhabditis elegans is described. A chemotactic index (CI) was employed as a way to precisely evaluate the response of worms to certain targets, and serve as a platform of comparison between strains and compounds of interest.

Many organisms use chemotaxis to seek out food sources, avoid noxious substances, and find mates.  Caenorhabditis elegans has impressive chemotaxis ...

Automating Aggregate Quantification in Caenorhabditis elegans

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This increased attention has called for the diversification and improvement of animal models capable of reproducing disease phenotypes observed in humans with PCDs. Though murine models have proven invaluable, they are expensive and are associated with laborious, low-throughput methods. Use of the Caenorhabditis elegans nematode model to study PCDs has been justified by its relative ease of maintenance, low cost, and rapid generation time, which allow for high-throughput applications. Additionally, high conservation between the C. elegans and human genomes makes this model an invaluable discovery tool. Nematodes that express fluorescently tagged tissue-specific polyglutamine (polyQ) tracts exhibit age- and polyQ length-dependent aggregation characterized by fluorescent foci. Such reporters are often employed as proxies to monitor changes in proteostasis across tissues. Manual aggregate quantification is time-consuming, limiting experimental throughput. Furthermore, manual foci quantification can introduce bias, as aggregate identification can be highly subjective. Herein, a protocol consisting of worm culturing, image acquisition, and data processing was standardized to support high-throughput aggregate quantification using C. elegans that express intestine-specific polyQ. By implementing a C. elegans-based image processing pipeline using CellProfiler, an image analysis software, this method has been optimized to separate and identify individual worms and enumerate their respective aggregates. Though the concept of automation is not entirely unique, the need to standardize such procedures for reproducibility, elimination of bias from manual counting, and increase throughput is high. It is anticipated that these methods can drastically simplify the screening process of large bacterial, genomic, or drug libraries using the C. elegans model.

Automating Aggregate Quantification in Caenorhabditis elegans

The following protocol describes the development and optimization of a high-throughput workflow for worm culturing, fluorescence imaging, and automated image processing to quantify polyglutamine aggregates as an assessment of changes in proteostasis.

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This ...

Investigating Collective Behaviors in Nematodes Using Dog Food Agar Medium

The Large-Scale Cultivation of Nematodes to Study Their Collective Behaviors

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 ...

Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae

Summary

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Chapters in this video

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Measuring Caenorhabditis elegans Life Span on Solid Media

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

Surveying Low-Cost Methods to Measure Lifespan and Healthspan in Caenorhabditis elegans

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

C. elegans Chemotaxis Assay

Automating Aggregate Quantification in Caenorhabditis elegans

The Large-Scale Cultivation of Nematodes to Study Their Collective Behaviors