Large-Scale Gravitaxis Assay of Caenorhabditis Dauer Larvae

Summary

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.

Abstract

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.

Introduction

Sensing Earth's gravitational pull is crucial for many organisms' 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's surroundings^1,2. Conversely, gravitactic preference can be altered in the presence of other stimuli^3,4,5. Therefore, gravitactic behavior is ideal for studying gravity sensation and understanding the nervous system'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 "stand" 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 adults^8,9. 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 assays^3,10,11. 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 design^13,14,15. The earliest reported gravitaxis assay in Caenorhabditis was performed by standing a plate on its side at a 90° 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 fields^16,17, 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 integrity^18,19. 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.

Protocol

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

1. 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).
2. 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).
3. 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).

2. Filling the chambers with agar

1. 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.
   ​NOTE: Covering the flask with cling-wrap (see Table of Materials) after autoclaving helps prevent bubbles from forming in the solution¹⁸.
   1. 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% agar^18,19.
2. 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.
   ​NOTE: Once prepared, each chamber must be used within 1 day or kept at 4 °C for several days. Cover any exposed openings with paraffin film to prevent the agar from drying out.

3. Isolating dauer larvae

1. 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.
2. 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.
3. Isolate dauers by treating with 1% SDS followed by a 30% sucrose flotation gradient following the previously published report²⁰.
   1. 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.
   2. 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.
4. 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.
   ​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.

4. Adding dauers to the chamber

1. 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 µL droplets under a dissecting microscope. Use the average of these counts to approximate the number of worms per µL.
   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.
2. Cut the end of a 10, 20, or 200 µL pipette tip to widen the bore. Set the micropipette to a slightly larger volume than the intended aspiration volume. Aspirate approximately 1-2 µL 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.
3. 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.

5. Running and scoring the assay

NOTE: Gravitaxis can be tested under various conditions that may affect behavior⁹.

1. To eliminate as many variables as possible, place the chambers within a dark Faraday cage (see Table of Materials) at room temperature.
2. 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.
3. 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.
4. 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.
   1. Also, avoid scoring worms that appear dead or are trapped in the liquid. Discard any chambers containing >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's surface and the pipette's wall. If burrowing behavior is observed, consider increasing agar concentration in future assays.
5. 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.

Results

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...

Discussion

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' orientational preference can be assayed in solution¹², this method is low throughput as worms must be analyzed one at a time. Addi...

Materials

Name	Company	Catalog Number	Comments
1% Sodium Dodecyl Sulfate solution			From stock 10% (w/v) SDS in DI water
15 mL Centrifuge tubes	Falcon	14-959-53A
3 mm Hex key			Other similar sized metal tools may be used
4% Agar in Normal Growth Medium (NGM) - 1 L			Prior to autoclaving: 3 g NaCl, 40 g Agar, 2.5 g Peptone, 2 g Dextrose, 10 mL Uracil (2 mg/mL), 500 μL 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)
5 mL Serological pipettes	Fisherbrand	S68228C	Polystyrene, not borosilicate glass
60% Cold sucrose solution			60% sucrose (w/v) in DI water; sterilize by filtration (0.45 μm filter). Keep at 4 °C
AF16 C. briggsae or other experimental strain			Available from the CGC (Caenorhabditis Genetics Center)
Bunsen burner
Cling-wrap	Fisherbrand	22-305654
Clinical centrifuge
Disposable razor blades	Fisherbrand	12-640
Faraday cage			Can be constructed using cardboard and aluminum foil; 30" L x 6" W x 26" H or larger
Ink markers			Sharpie or other brand for marking on plastic
Labeling tape	Carolina	215620
M9 buffer			22 mM KH2PO4, 42 mM Na2HPO4, 86 mM NaCl
N2 C. elegans strain			Available from the CGC (Caenorhabditis Genetics Center)
NGM plates with OP50			1.7% (w/v) agar in NGM (see description: 4% agar in NGM). Seed with OP50
Paraffin film	Bemis	13-374-10
Plastic cutting board
Pliers
Rotating vertical mixer	BTLab SYSTEMS	BT913	With 22 x 15 mL tube bar
Serological pipettor	Corning	357469
Stereo Microscope	Laxco	S2103LS100
Tally counter	ULINE	H-7350
Thick NGM/agar plate media - 1 L			See 4% Agar in NGM recipe; replace 40 g Agar with 20 g Agar
Tweezers