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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#956;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 &#956;m filter). Keep at 4 &#176;C</td>
		</tr>
		<tr>
			<td>AF16 C. briggsae 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&#34; L x 6&#34; W x 26&#34; 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 KH2PO4, 42 mM Na2HPO4, 86 mM NaCl</td>
		</tr>
		<tr>
			<td>N2 C. elegans 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

2.1K 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

Asymmetric Walkway: A Novel Behavioral Assay for Studying Asymmetric Locomotion

Behavioral assays are commonly used for the assessment of sensorimotor impairment in the central nervous system (CNS). The most sophisticated methods for quantifying locomotor deficits in rodents is to measure minute disturbances of unconstrained gait overground (e.g., manual BBB score or automated CatWalk). However, cortical inputs are not required for the generation of basic locomotion produced by the spinal central pattern generator (CPG). Thus, unconstrained walking tasks test locomotor deficits due to motor cortical impairment only indirectly. In this study, we propose a novel, precise foot-placement locomotor task that evaluates cortical inputs to the spinal CPG. An instrumented peg-way was used to impose symmetrical and asymmetrical locomotor tasks mimicking lateralized movement deficits. We demonstrate that shifts from equidistant inter-stride lengths of 20% produce changes in the forelimb stance phase characteristics during locomotion with preferred stride length. Furthermore, we propose that the asymmetric walkway allows for measurements of behavioral outcomes produced by cortical control signals. These measures are relevant for the assessment of impairment after cortical damage.

Here, we present a protocol to quantify precise stepping in rodents. Cortical and the spinal central pattern generator signals are required for precise foot-placement during obstructed locomotion. We report here the novel constrained walking task that directly examines precise stepping behavior.

Behavioral assays are commonly used for the assessment of sensorimotor impairment in the central nervous system (CNS). The most sophisticated methods for ...

The Treadmill Fatigue Test: A Simple, High-throughput Assay of Fatigue-like Behavior for the Mouse

Fatigue is a prominent symptom in many diseases and disorders and reduces quality of life for many people. The lack of clear pathogenesis and failure of current interventions to adequately treat fatigue in all patients leaves a need for new treatment options. Despite the therapeutic need and importance of preclinical research in helping identify promising novel treatments, few preclinical assays of fatigue are available. Moreover, the most common preclinical assay used to assess fatigue-like behavior, voluntary wheel running, is not suitable for use with some strains of mice, may not be sensitive to drugs that reduce fatigue, and has relatively low throughput. The current protocol describes a novel, non-voluntary preclinical assay of fatigue-like behavior, the treadmill fatigue test, and provides evidence of its efficacy in detecting fatigue-like behavior in mice treated with a chemotherapy drug known to cause fatigue in humans and fatigue-like behavior in animals. This assay may be a beneficial alternative to wheel running, as fatigue-like behavior and potential interventions can be assessed in a greater number of mice over a shorter time frame, thus permitting faster discovery of new therapeutic options.

Fatigue is a common, undertreated and frequently poorly-understood symptom in many diseases and disorders. New preclinical assays of fatigue may help to improve current understanding and future treatment of fatigue. To that end, the current protocol provides a novel means of measuring fatigue-like behavior in the mouse.

Fatigue is a prominent symptom in many diseases and disorders and reduces quality of life for many people. The lack of clear pathogenesis and failure of ...

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.

The nematode, Caenorhabditis elegans' compact nervous system of only 302 neurons underlies a diverse repertoire of behaviors. To facilitate the dissection ...

A Simple and Low-cost Assay for Measuring Ambulation in Mouse Models of Muscular Dystrophy

Measuring functional outcomes in the treatment of muscular dystrophy is an essential aspect of preclinical testing. The assessment of voluntary ambulation in mouse models is a non-invasive and reproducible activity assay that is directly analogous to measures of patient ambulation such as the 6-minute walk test and related mobility scores. Many common methods for testing mouse ambulation speed and distance are based on the open field test, where an animal&#39;s free movement within an arena is measured over time. One major downside to this approach is that commercial software and equipment for high-resolution motion tracking is expensive and may require transferring mice to specialized facilities for testing. Here, we describe a low-cost, video-based system for measuring mouse ambulation that utilizes free and open-source software. Using this protocol, we demonstrate that voluntary ambulation in the dystrophin-null mdx mouse model for Duchenne muscular dystrophy (DMD) is decreased relative to wild-type mouse activity. In mdx mice expressing the utrophin transgene, these activity deficits are not observed and the total distance traveled is indistinguishable from wild-type mice. This method is effective for measuring changes in voluntary ambulation associated with dystrophic pathology, and provides a versatile platform that can be readily adapted to diverse research settings.

This protocol describes a flexible, low-cost system for measuring mouse ambulation in an open field activity assay. We show that a 6-minute ambulation assay based on this system detects a decrease in voluntary movement in mdx mice, and accurately distinguishes improvement in a muscle-specific rescue of these animals.

Measuring functional outcomes in the treatment of muscular dystrophy is an essential aspect of preclinical testing. The assessment of voluntary ambulation ...

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen distribution. It is also possible that aberrant behavior of oxygen-sensing neurons could induce hypoxia-like behavior in the presence of normal oxygen levels. In D. melanogaster, development at low oxygen levels results in inhibition of growth and sluggish behavior during the larval phases. However, these established manifestations of oxygen deficit overlap considerably with the phenotypes of many mutations that regulate growth, stress responses or locomotion. As result, there is currently no assay available to identify i) cellular hypoxia induced by a mutation or ii) hypoxia-like behavior when induced by abnormal neuronal behavior.
We have recently identified two distinctive behaviors in D. melanogaster larvae that occur at normal oxygen levels in response to internal detection of hypoxia. First, at all stages, such larvae avoid burrowing into food, often straying far away from a food source. Second, tunneling into a soft substratum, which normally occurs during the wandering third instar stage is completely abolished if larvae are hypoxic. The assay described here is designed to detect and quantitate these behaviors and thus to provide a way to detect hypoxia induced by internal damage rather than low external oxygen. Assay plates with an agar substratum and a central plug of yeast paste are used to support animals through larval life. The positions and state of the larvae are tracked daily as they proceed from first to third instar. The extent of tunneling into the agar substratum during wandering phase is quantitated after pupation using NIH ImageJ. The assay will be of value in determining when hypoxia is a component of a mutant phenotype and thus provide insight into possible sites of action of the gene in question.

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

The protocol describes a simple assay to identify Drosophila melanogaster larvae that are experiencing hypoxia under normal atmospheric oxygen levels. This protocol allows hypoxic larvae to be distinguished from other mutants that show overlapping phenotypes such as sluggishness or slow growth.

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen ...

Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans

The swimming assay described in this protocol is a valid tool to identify proteins regulating the dopaminergic synapses. Similar to mammals, dopamine (DA) controls several functions in C. elegans including learning and motor activity. Conditions that stimulate DA release (e.g., amphetamine (AMPH) treatments) or that prevent DA clearance (e.g., animals lacking the DA transporter (dat-1) which are incapable of reaccumulating DA into the neurons) generate an excess of extracellular DA ultimately resulting in inhibited locomotion. This behavior is particularly evident when animals swim in water. In fact, while wild-type animals continue to swim for an extended period, dat-1 null mutants and wild-type treated with AMPH or inhibitors of the DA transporter sink to the bottom of the well and do not move. This behavior is termed "Swimming Induced Paralysis" (SWIP). Although the SWIP assay is well established, a detailed description of the method is lacking. Here, we describe a step-by-step guide to perform SWIP. To perform the assay, late larval stage-4 animals are placed in a glass spot plate containing control sucrose solution with or without AMPH. Animals are scored for their swimming behavior either manually by visualization under a stereoscope or automatically by recording with a camera mounted on the stereoscope. Videos are then analyzed using a tracking software, which yields a visual representation of thrashing frequency and paralysis in the form of heat maps. Both the manual and automated systems guarantee an easily quantifiable readout of the animals' swimming ability and thus facilitate screening for animals bearing mutations within the dopaminergic system or for auxiliary genes. In addition, SWIP can be used to elucidate the mechanism of action of drugs of abuse such as AMPH.

Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans

Swimming induced paralysis (SWIP) is a well-established behavioral assay used to study the underlying mechanisms of dopamine signaling in Caenorhabditis elegans (C. elegans). However, a detailed method to perform the assay is lacking. Here, we describe a step-by-step protocol for SWIP.

The swimming assay described in this protocol is a valid tool to identify proteins regulating the dopaminergic synapses. Similar to mammals, dopamine (DA) ...

Calcium Imaging in Freely Behaving Caenorhabditis elegans with Well-Controlled, Nonlocalized Vibration

Nonlocalized mechanical forces, such as vibrations and acoustic waves, influence a wide variety of biological processes from development to homeostasis. Animals cope with these stimuli by modifying their behavior. Understanding the mechanisms underlying such behavioral modification requires quantification of neural activity during the behavior of interest. Here, we report a method for calcium imaging in freely behaving Caenorhabditis elegans with nonlocalized vibration of specific frequency, displacement, and duration. This method allows the production of well-controlled, nonlocalized vibration using an acoustic transducer and quantification of evoked calcium responses at single-cell resolution. As a proof of principle, the calcium response of a single interneuron, AVA, during the escape response of C. elegans to vibration is demonstrated. This system will facilitate understanding of neural mechanisms underlying behavioral responses to mechanical stimuli.

Calcium Imaging in Freely Behaving Caenorhabditis elegans with Well-Controlled, Nonlocalized Vibration

Reported here is a system for calcium imaging in freely behaving Caenorhabditis elegans with well-controlled, nonlocalized vibration. This system allows researchers to evoke nonlocalized vibrations with well-controlled properties at nano-scale displacement and to quantify calcium currents during responses of C. elegans to the vibrations.

Nonlocalized mechanical forces, such as vibrations and acoustic waves, influence a wide variety of biological processes from development to homeostasis. ...

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila larvae, customized metal nickel-titanium alloy (nitinol) filaments. These mechanical probes are similar to the von Frey filaments used in vertebrates to measure mechanical nociception. Here, we demonstrate how to make and calibrate these mechanical probes and how to generate a full behavioral dose-response from subthreshold (innocuous or non-noxious range) to suprathreshold (low to high noxious range) stimuli. To demonstrate the utility of the probes, we investigated tissue damage-induced hypersensitivity in Drosophila larvae. Mechanical allodynia (hypersensitivity to a normally innocuous mechanical stimulus) and hyperalgesia (exaggerated responsiveness to a noxious mechanical stimulus) have not yet been established in Drosophila larvae. Using mechanical probes that are normally innocuous or probes that typically elicit an aversive behavior, we found that Drosophila larvae develop mechanical hypersensitization (both allodynia and hyperalgesia) after tissue damage. Thus, the mechanical probes and assay that we illustrate here will likely be important tools to dissect the fundamental molecular/genetic mechanisms of mechanical hypersensitivity.

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

The goal of this protocol is to show how to perform an improved assay for mechanical nociception in Drosophila larvae. We use the assay here to demonstrate that mechanical hypersensitivity (allodynia and hyperalgesia) exists in Drosophila larvae.

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila ...

Comparative Analysis of Experimental Methods to Quantify Animal Activity in Caenorhabditis elegans Models of Mitochondrial Disease

Caenorhabditis elegans is widely recognized for its central utility as a translational animal model to efficiently interrogate mechanisms and therapies of diverse human diseases. Worms are particularly well-suited for high-throughput genetic and drug screens to gain deeper insight into therapeutic targets and therapies by exploiting their fast development cycle, large brood size, short lifespan, microscopic transparency, low maintenance costs, robust suite of genomic tools, mutant repositories, and experimental methodologies to interrogate both in vivo and ex vivo physiology. Worm locomotor activity represents a particularly relevant phenotype that is frequently impaired in mitochondrial disease, which is highly heterogeneous in causes and manifestations but collectively shares an impaired capacity to produce cellular energy. While a suite of different methodologies may be used to interrogate worm behavior, these vary greatly in experimental costs, complexity, and utility for genomic or drug high-throughput screens. Here, the relative throughput, advantages, and limitations of 16 different activity analysis methodologies were compared that quantify nematode locomotion, thrashing, pharyngeal pumping, and/or chemotaxis in single worms or worm populations of C. elegans at different stages, ages, and experimental durations. Detailed protocols were demonstrated for two semi-automated methods to quantify nematode locomotor activity that represent novel applications of available software tools, namely, ZebraLab (a medium-throughput approach) and WormScan (a high-throughput approach). Data from applying these methods demonstrated similar degrees of reduced animal activity occurred at the L4 larval stage, and progressed in day 1 adults, in mitochondrial complex I disease (gas-1(fc21)) mutant worms relative to wild-type (N2 Bristol) C. elegans. This data validates the utility for these novel applications of using the ZebraLab or WormScan software tools to quantify worm locomotor activity efficiently and objectively, with variable capacity to support high-throughput drug screening on worm behavior in preclinical animal models of mitochondrial disease.

Comparative Analysis of Experimental Methods to Quantify Animal Activity in Caenorhabditis elegans Models of Mitochondrial Disease

This study presents protocols for two semi-automated locomotor activity analysis approaches in C. elegans complex I disease gas-1(fc21) worms, namely, ZebraLab (a medium-throughput assay) and WormScan (a high-throughput assay) and provide comparative analysis among a wide array of research methods to quantify nematode behavior and integrated neuromuscular function.

Caenorhabditis elegans is widely recognized for its central utility as a translational animal model to efficiently interrogate mechanisms and therapies of ...

A High Throughput Microplate Feeder Assay for Quantification of Consumption in Drosophila

Quantifying food intake in Drosophila is used to study the genetic and physiological underpinnings of consumption-associated traits, their environmental factors, and the toxicological and pharmacological effects of numerous substances. Few methods currently implemented are amenable to high throughput measurement. The Microplate Feeder Assay (MFA) was developed for quantifying the consumption of liquid food for individual flies using absorbance. In this assay, flies consume liquid food medium from select wells of a 1536-well microplate. By incorporating a dilute tracer dye into the liquid food medium and loading a known volume into each well, absorbance measurements of the well acquired before and after consumption reflect the resulting change in volume (i.e., volume consumed). To enable high throughput analysis with this method, a 3D-printed coupler was designed that allows flies to be sorted individually into 96-well microplates. This device precisely orients 96- and 1536-well microplates to give each fly access to up to 4 wells for consumption, thus enabling food preference quantification in addition to regular consumption. Furthermore, the device has barrier strips that toggle between open and closed positions to allow for controlled containment and release of a column of samples at a time. This method enables high throughput measurements of consumption of aqueous solutions by many flies simultaneously. It also has the potential to be adapted to other insects and to screen consumption of nutrients, toxins, or pharmaceuticals.

A High Throughput Microplate Feeder Assay for Quantification of Consumption in Drosophila

The microplate feeder assay offers an economical, high throughput method for quantifying liquid food consumption in Drosophila. A 3D-printed device connects a 96-well microplate in which flies are housed to a 1536-well microplate from which flies consume a feeding solution with a tracer dye. The solution volume decline is measured spectrophotometrically.

Quantifying food intake in Drosophila is used to study the genetic and physiological underpinnings of consumption-associated traits, their environmental ...