Name: large-scale gravitaxis assay of caenorhabditis dauer larvae
Uploaded: 2022-05-31T15:00:00
Description: Watch this Scientific Journal Video about Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae at JoVE.com

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 t...

<ol>
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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. Addi...

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 t...

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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 ...

This high throughput assay can be used to observe robust gravitactic behavior in caenorhabditis dauer larvae. Worm behavior is observed over large distances compared with assays performed on a petri dish. This enhances the sensitivity of the assay and allows for detailed data analysis.Studying gravitaxis behavior can provide insight into how gravity sensation occurs in caenorhabditis, which has relevance for understanding the mammalian vestibular system. Creating the gravitaxis assay chambers, isolating dauers, and injecting the worms into the chambers takes practice. You can save time by isolating the dauers and creating the chambers a day in advance.Begin by setting up a Bunsen burner, one to two razor blades, pliers, tweezers, and a plastic cutting surface, and a fume hood. To make a chamber, gather two 5 milliliter serological pipettes, and with tweezers, remove the cotton plug from one pipette. Hold a razor blade using pliers over the Bunsen burner until hot.Use the heated blade to slice off the tapered end of the second pipette so that the entire pipette has a uniform diameter. Now, quickly bring the two modified pipette ends close to the flame and melt them slightly. Join these ends by pressing them together firmly, ensuring that the walls of both pipettes are continuous.If any gaps are visible after joining, break them apart and repeat this step. Prepare NGM with 4%agar according to standard worm procedures and fill each chamber by attaching it to a serological pipetter while the agar is still molten. To minimize any variations in the consistency of the agar due to uneven cooling, hold the pipetter parallel to the bench top while drawing up the agar.Then slowly draw the solution and seal the tip with parafilm before removing the chamber from the pipetter. Lay the chamber flat to cool and allow the agar to harden before moving. Once cooled, heat a three-millimeter hex key over a Bunsen burner and create a small plastic opening by firmly pressing it into the wall of each chamber about five millimeters to one side of the midline.Next, use a heated blade to remove the cotton and tapered ends of the chamber and seal the ends with paraffin film. 10 to 15 days before the experiment, chunk each strain onto two to three large NGM plates with OP50 bacteria and wrap the paraffin film. Collect worms by rinsing the lids and plates with M9 Buffer and pipetting the solution into 15-milliliter centrifuge tubes.Pellet the worms by spinning at 1600 x G for 30 to 60 seconds at room temperature and aspirate most of the M9 Buffer with a pipette or vacuum aspirator. Add seven milliliters of 1%sodium dodecyl sulfate solution to each worm pellet and leave the worms in it for 30 minutes. To allow aeration, keep rotating the tubes continuously during this time.Then, remove the detergent by rinsing it three to five times with M9.After adding five milliliters M9, add five milliliters of cold filtered 60%sucrose solution. Mix thoroughly and create a separation gradient by centrifuging at 1600 x G for five minutes at room temperature. Fill new 15-milliliter centrifuge tubes with two milliliters of M9 Buffer.Now crush the ends of glass Pasteur pipettes to widen the bore. And use these pipettes to transfer the top layer of the solution from the sucrose gradient to the new tubes. Rinse the dauers three to five times with M9 Buffer.Centrifuge dauers at 1600 x G for five minutes at room temperature and aspirate most of the M9 solution with a pipette or vacuum aspirator. Next, estimate the worm density by manually counting the number of worms in three separate 1-microliter droplets under a dissecting microscope, and use the average of these counts to approximate the number of worms per microliter. Now widen the tip bore by cutting the end of 10, 20, or 200-microliter pipette tips.Set the micropipette to a slightly larger volume than the intended aspiration volume. Aspirate approximately 1 to 2 microliters of the concentrated worm solution and allow a small amount of air to enter the tip. Gently push the pipette tip into the agar while depressing the micro pipette.This will create an injection site without clogging the tip. Release the worms into the agar and use paraffin film to seal the opening. To eliminate as many variables as possible once the chambers are placed within a dark Faraday cage at room temperature, label each chamber and hang it vertically within the Faraday cage.Test horizontal controls concurrently by laying some chambers flat within the same environmental conditions. Then use the vertically-oriented N2 worms as a positive control and horizontally-oriented chambers containing N2 dauers as an additional negative control against the experimental strains. After a few hours, the dauer worms begin to disperse from the start site and take several hours to reach either end of the chamber.Leave the chambers undisturbed during this time and score within 12 to 24 hours following injection. Next, remove the chambers one at a time and score gravitaxis by looking for live dauers under a dissecting microscope and marking their locations with ink. Do not score worms within 2.5 centimeters to either side of the injection site as these worms are not likely to be demonstrating a directional preference.Also, avoid scoring worms that appear dead or are trapped in the liquid and discard any chambers containing more than 50%dead or swimming worms. To quantify the results, use a marker to divide each half of the chamber into seven 3.5 centimeters, starting 2.5 centimeters away from the injection site. Use a manual tally counter to tally the number of worms observed in each section.In caenorhabditis briggsae, a large percentage of dauer worms showed migration toward the top of the chambers. However, the horizontal controls showed distribution around the center of the chambers, indicating negative gravitactic behavior. Also, the vertical distributions of caenorhabditis briggsae and caenorhabditis elegans did not show any difference suggesting that caenorhabditis briggsae dauers show a negative gravitaxis behavior similar to caenorhabditis elegans dauers.Paralytic agents such as sodium azide are not used in this protocol. Therefore, it is important to score the gravitaxis chambers as soon as they're removed from the Faraday cage. This assay led to the discovery of negative gravitaxis behavior in c.elegans.Through testing several mutant strains, it also revealed genetic and neural requirements for gravitaxis in worms.

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

Research

JoVE Journal

Behavior

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 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 ...

Real-Time Void Spot Assay

Normal voiding behavior is the result of the coordinated function of the bladder, the urethra, and the urethral sphincters under the proper control of the nervous system. To study voluntary voiding behavior in mouse models, researchers have developed the void spot assay (VSA), a method that measures the number and area of urine spots deposited on a filter paper lining the floor of an animal&#8217;s cage. Although technically simple and inexpensive, this assay has limitations when used as an end-point assay, including a lack of temporal resolution of voiding events and difficulties quantifying overlapping urine spots. To overcome these limitations, we developed a video-monitored VSA, which we call real-time VSA (RT-VSA), and which allows us to determine voiding frequency, assess voided volume and voiding patterns, and make measurements over 6 h time windows during both the dark and light phases of the day. The method described in this report can be applied to a wide variety of mouse-based studies that explore the physiological and neurobehavioral aspects of voluntary micturition in health and disease states.

This article describes a new method to study mouse voiding behavior by incorporating video monitoring in the conventional void spot assay. This approach provides temporal, spatial, and volumetric information on the voiding events and details of mouse behavior during the light and dark phases of the day.

Normal voiding behavior is the result of the coordinated function of the bladder, the urethra, and the urethral sphincters under the proper control of the ...

High-Throughput Assay to Measure &lt;em&gt;Caenorhabditis elegans&lt;/em&gt; Food Intake and Lifespan

Quantifying Food Intake in Caenorhabditis elegans by Measuring Bacterial Clearance

Feeding is an essential biological process for an organism&#39;s growth, reproduction, and survival. This assay aims to measure the food intake of Caenorhabditis elegans&#160;(C. elegans), an important parameter when studying the genetics of aging or metabolism. In most species, feeding is determined by measuring the difference between the amount of food provided and the amount left after a given time interval. The method presented here uses the same strategy to determine the feeding of C. elegans. It measures the amount of bacteria, the food source of C. elegans, cleared within 72 h. This method uses 96-well microtiter plates and has allowed the screening of hundreds of drugs for their ability to modulate food intake at a speed and depth not possible in other animal models. The strength of this assay is that it allows to measure feeding and lifespan simultaneously and directly measures the disappearance of food and, thus, is based on the same principles used for other organisms, facilitating species-to-species comparison.

Author Spotlight: Exploring the Relationship Between Lipotoxicity and HFpEF 

This protocol describes an assay to quantify Caenorhabditis elegans feeding rate based on measuring the clearance of bacteria in liquid culture.

Feeding is an essential biological process for an organism&#39;s growth, reproduction, and survival. This assay aims to measure the food intake of ...

Extraction and Quantification of Volatile Sex Pheromone Responses in C. elegans

Volatile Sex Pheromone Extraction and Chemoattraction Assay in Caenorhabditis elegans

Chemical communication is vital in organismal health, reproduction, and overall well-being. Understanding the molecular pathways, neural processes, and computations governing these signals remains an active area of research. The nematode Caenorhabditis elegans provides a powerful model for studying these processes as it produces a volatile sex pheromone. This pheromone is synthesized by virgin females or sperm-depleted hermaphrodites and serves as an attractant for males.
This protocol describes a detailed method for isolating the volatile sex pheromone from several C. elegans strains (WT strain N2, daf-22, and fog-2) and C. remanei. We also provide a protocol for quantifying the male chemotaxis response to the volatile sex pheromone. Our analysis utilizes measurements such as chemotaxis index (C.I.), arrival time (A.T.), and a trajectory plot to compare male responses under various conditions accurately. This method allows for robust comparisons between males of different genetic backgrounds or developmental stages. Furthermore, the experimental setup outlined here is adaptable to investigating other chemoattraction chemicals.

Author Spotlight: Examining Volatile Sex Pheromone Influence on Male C. elegans Behavior

This protocol establishes methods for extracting and quantifying responses to the volatile sex pheromone in C. elegans, providing tools to study chemical communication and navigation trajectory.

Chemical communication is vital in organismal health, reproduction, and overall well-being. Understanding the molecular pathways, neural processes, and ...

Preparation of OP50 for Caenorhabditis elegans Culture

To begin, inoculate a single OP50 colony in five milliliters of LB with ampicillin and amphotericin B and incubate for approximately six hours at 37 degrees Celsius in a bacterial shaker. Once the culture becomes cloudy, dilute the preinoculum culture of OP50 by 1 to 2, 000 in 250 milliliters of TB with ampicillin and glycerol. Then transfer the OP50 liquid culture into a sterile centrifugation tube.Centrifuge the tube for 15 minutes at 3, 100 G in a tabletop centrifuge at four degrees Celsius. Discard the supernatant and resuspend the OP50 pellet in sterile water before recentrifugation. After the second wash, discard the supernatant and resuspend the OP50 pellet in sterile water to a volume of 50 milliliters.Transfer it to a pre-weighed 50 milliliter conical tube. Centrifuge the tube for 20 minutes to pellet the OP50. At the end of the centrifugation, carefully remove all remaining supernatant, ensuring no water remains in the tube.Calculate the weight of the pellet by subtracting the weight of the empty centrifugation tube from the weight of the centrifugation tube with the pellet. Thoroughly resuspend the OP50 pellet in S-complete buffer to achieve a concentration of 100 milligrams per milliliter. Test the OP50 suspension on an agar plate to check for any contamination.Store the OP50 suspension in S-complete buffer at four degrees Celsius.