Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans

Summary

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.

Abstract

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.

Introduction

Animals perform a variety of innate and complex behaviors that are mediated by different neurotransmitters coordinated by intricate signaling processes. The neurotransmitter dopamine (DA) mediates highly conserved behaviors across species, including learning, motor function and reward processing.

The soil nematode C. elegans, with a relatively simple and well mapped nervous system consisting of only 302 neurons, shows markedly complex behaviors, including many that are regulated by DA such as mating, learning, foraging, locomotion and egg laying¹. Among other features, short life cycle, ease of handling and the conservation of signaling molecules, highlight the advantages of using C. elegans as a model for studying the neural basis of conserved behaviors.

The hermaphrodite C. elegans contains eight dopaminergic neurons; In addition to these, the male contains six extra pairs for mating purposes. As in mammals, these neurons synthesize DA and express the DA transporter (DAT-1), a membrane protein found exclusively in dopaminergic neurons, which transports DA released in the synaptic cleft back into the dopaminergic neurons. Moreover, most of the proteins involved in each step of synthesis, packaging and release of DA are highly conserved between worms and humans and, like in mammals, DA modulates feeding behaviors and locomotion in C. elegans².

C. elegans crawls on solid surfaces and swims with a characteristic thrashing behavior in water. Interestingly, mutants lacking expression of DAT-1 (dat-1) crawl normally on solid surface but fail to sustain swimming when immersed in water. This behavior was termed swimming induced paralysis, or SWIP. Previous experiments demonstrated that SWIP, in part, is caused by an excess of DA in the synaptic cleft that ultimately overstimulates the D2-like postsynaptic receptors (DOP-3). Although originally identified in dat-1 knockout animals³, SWIP is also observed in wild-type animals treated with drugs that block the activity of DAT (e.g., imipramine⁴) and/or induce DA release (e.g., amphetamine⁵). On the other hand, pharmacological or genetic manipulations averting synthesis and release of DA and blocking DOP-3 receptor function prevent SWIP⁶. Taken together, these already published data have established SWIP as a reliable tool to study the behavioral effects caused by mutated proteins within dopaminergic synapses^3,4,7 and to be employed for forward genetic screens for the identification of novel regulatory pathways involved in DA signaling^{7,8,9,10,11,12}. Additionally, by providing an easily quantifiable readout of drug-induced behavior in living animals, SWIP enables the elucidation of mechanisms of action of drugs like amphetamine (AMPH) and azaperone at the dopaminergic synapses^5,6,13,14,15.

Protocols for performing the SWIP assays have been described before¹⁶. Here, we describe in detail the methodology and setup to perform the assay with the goal of providing a visual guide for the C. elegans community to effectively perform SWIP.

Protocol

1. Preparation of Solutions and Media

1. Prepare M9 buffer by dissolving KH₂PO₄ 3.0 g (22.05 mM), Na₂HPO₄ 6.0 g (42.2 mM), and NaCl 5.0 g (85.5 mM) in 1 L of autoclaved deionized water. Add 1.0 mL of 1 M MgSO₄ (12 g in a final volume of 100 mL autoclaved deionized water) after autoclaving. Mix 100 mL of the resulting 10x M9 with 900 mL of autoclaved deionized water to make a 1x solution.
2. To make egg buffer, dissolve 6.896 g of NaCl (118 mM), 3.578 g of KCl (48 mM), 0.294 g of CaCl₂-2H₂O (2 mM), 0.406 g of MgCl₂-6H₂O (2 mM) and 5.958 g (25 mM) of HEPES in 1 L of autoclaved deionized water. Adjust pH to 7.3 using NaOH.
3. Prepare fresh sodium hypochlorite/NaOH solution by adding 1 mL of 5-6% sodium hypochlorite (bleach) and 180 µL of 10 N NaOH to 3.8 mL of deionized water.
4. Weigh 60 g sucrose and dissolve in autoclaved deionized water to a final volume of 100 mL to make 60% sucrose solution.
5. Dissolve 0.684 g of sucrose in 10 mL of autoclaved deionized water to make 200 mM sucrose. Check and adjust to the same an osmolarity using osmometer. Make 1 mL aliquots in 1.5 mL microcentrifuge tubes and freeze at -20 °C.
6. Weigh 0.184 g of AMPH (molecular weight 184.75 g/mol) and dissolve in 10 mL of deionized water to make a 100 mM stock solution. Mix 2 µL of the stock solution in 400 µL of water to make 0.5 mM working solution.
7. Prepare Nutrient growth media (NGM) plates
   1. Mix 3 g of NaCl (52.65 mM), 20 g of peptone, 25 g of bacto-agar and 975 mL of deionized water in a 2 L Erlenmeyer flask. Include a magnetic stir bar and autoclave (121 °C, 15 PSI) for 1 hour using liquid cycle.
   2. Cool to and maintain the temperature at about 50 °C by placing the flask on a heater while stirring. Add 0.5 mL of cholesterol (5 mg/mL in ethanol), 1 mL of 1 M MgSO₄, 1 mL of 1 M CaCl₂ and 25 mL of 1 M potassium phosphate buffer, pH 7.4 (108.3 g of KH₂PO₄, 35.6 g of K₂HPO₄, deionized water to 1 L).
   3. Pipette 25 mL each into 100 mm x 15 mm Petri plates and allow the media to solidify. Store the plates upside down at 4 °C in a box for up to 4 weeks.
8. Preparation of Lysogeny broth (LB) broth
   1. Dissolve 5 g of LB powder mix in 200 mL of deionized water in an Erlenmeyer flask. Autoclave for 30 minutes utilizing the liquid sterilization cycle. Allow the broth to cool down. Store at room temperature for 1-2 weeks.
9. Preparation of NA22 bacterial plates
   1. Use a sterile pipette tip or a sterile bacterial loop to streak an LB plate with a small volume of NA22 E. coli bacteria from glycerol stock and incubate the plate upside down in a 37 °C incubator overnight to grow isolated colonies. Pick and introduce a single colony into 200 mL of LB broth prepared in step 1.8 and let grow overnight at 37 °C on a shaking platform.
   2. To seed the plates, dispense 200 µL of bacterial culture onto the NGM plates prepared earlier in step 1.7 and spread with a sterile glass hockey stick. Let the plates dry overnight or longer under a hood and store upside down in an airtight box at 4 °C.
10. To make the eyelash/platinum tool to pick worms, glue a thick eyelash or a platinum filament into a glass Pasteur pipette using super glue. Cut the tip of the eyelash at an angle using a razor blade. Alternatively, a Bunsen burner can be used to melt the tip of the glass pipette around the platinum filament.

2. C. elegans Husbandry

NOTE: Culture wild-type N2C. elegans strain on Escherichia coli NA22 plates. The detailed culture methods are described below.

1. Preparation of worm culture
   1. To make a starter culture of worms, cut a small piece of agar from a plate containing well-fed animals and transfer it onto a NA22 E. coli bacteria plate prepared in step 1.9 using a sterile spatula. Incubate plates at 20 °C for 3-4 days. Under a stereo microscope, visually confirm the presence of gravid adults.
2. Preparation of synchronized population of worms
   1. Collect gravid adults from at least 2 plates by dispensing autoclaved deionized water all around the plate using a squirt bottle. Gently swirl the plate to dislodge the worms and collect the worms into a 15 mL polystyrene conical tube using a disposable plastic pipette.
   2. Spin down the tube in a centrifuge at 140 x g for 2 min to pellet the worms, then aspirate off supernatant using a vacuum pump or with built in laboratory vacuum.
   3. Resuspend and wash the worms by filling the tube with autoclaved deionized water and mix and centrifuge at 140 x g for 2 min. Aspirate the supernatant and repeat this last step two more times or until worms are clear from bacteria (water appears clear when mixed with the worms).
   4. Add 5 mL of freshly made sodium hypochlorite/NaOH solution (step 1.3) to the worm pellet and rapidly mix using a vortex. Incubate the tube on a rocker for about 4-8 min. The time of incubation with sodium hypochlorite/NaOH solution fluctuates between 4-8 minutes based on the quality of the stock sodium hypochlorite solution (bleach).
   5. Put a drop (2-50 µL) of solution containing worms on a glass microscope slide and check every 2 min under the microscope for worm lysis. When about 70% of the worms are lysed and eggs are released, fill the tube with egg buffer prepared in step 1.2 and immediately centrifuge for 1 min at 140 x g to pellet the embryos and worm carcasses.
   6. Aspirate the supernatant and wash the pellet 3 more times by filling the tube each time with egg buffer. Spin down at 140 x g for 1 min and remove the supernatant each time. The pellet turns white at the end of washes.
   7. After the final wash, separate the embryos from the dead carcasses in 30% sucrose solution. Add 5 mL of autoclaved deionized water to the pellet, resuspend and add 5 mL of 60% sucrose prepared in step 1.4. Mix thoroughly and centrifuge at 160 x g for 6 min.
   8. Use a glass Pasteur pipette to transfer the embryos floating at the upper meniscus into a fresh 15 mL conical tube. Do not take more than 3-4 mL. To remove any remaining sucrose, wash the embryos 3 times with autoclaved water by centrifuging at 140 x g for 3 min, removing the supernatant and resuspending the pellet (and filling the tube) each time.
   9. Repeat the washes with 1x M9 buffer. After the final wash, resuspend the pellet in 10 mL of M9. Leave the tubes on a shaker overnight (no more than 14 h) for the eggs to hatch into L1 larvae. Worms will remain in L1 larval stage due to lack of food.
   10. Wash the L1 larvae 3 times with autoclaved water to remove any pheromones released by the larvae by centrifuging at 140 x g for 2 min. Resuspend the larvae in 1 mL of water. Make a 1:10 dilution of the worms in water, pipette a 10 µL drop on a glass slide, put a coverslip on and count the number of worms under a stereoscope. Repeat this twice and average the results.
   11. Pipette the volume of worms that corresponds to about 1,000 worms onto an NA22 plate (that was previously brought to room temperature) by placing small drops on the plate. Leave the plate half-open until the drop dries out. Then cover the plate and incubate upside down in 20 °C incubator for about 44-48 h or until the worms reach late L4 stage, as confirmed visually under a stereomicroscope. Now the worms are ready to be tested for SWIP.

3. SWIP

NOTE: We describe the manual method of assessing SWIP in wild-type worms treated with AMPH. We also briefly discuss the tracking of worms and further analysis of worm kinetics using an automated worm tracker and a tracking software which were previously described by Hardaway et al.¹⁰.

1. Manual method to test for SWIP
   1. Aliquot 40 µL of 200 mOsm/L sucrose solution either with or without 0.5 mM AMPH into a glass spot plate. Under the stereoscope, pick 8-10 late-L4 stage worms with an eyelash or platinum pick and submerge the pick in the plate containing the solution until worms move off the pick and swim into the solution. Note the number of worms picked into the well, start the timer, observe and record the number of worms exhibiting SWIP at each minute mark.
   2. Copy the raw data into a spreadsheet and calculate the percent of worms paralyzed by dividing the number of worms paralyzed at each minute by total number of worms tested throughout the assay and multiply by 100. Copy the percent values into any graphing and statistical software and plot the data with percent values on the Y axis and time on X axis using the XY graph format.
   3. Perform two-way ANOVA followed by post-hoc analysis (e.g., Bonferroni post-test) to test for statistical significance among control, AMPH groups and time of treatment.
2. Automated analysis of SWIP
   1. Perform automated analysis on a single worm at a time. The protocol to set up camera, the worm tracker software and script to run the tracking software analysis are described in detail in Hardaway et al.¹⁶.
   2. Briefly, place a single late L4 stage hermaphrodite into a glass spot plate utilizing an eyelash pick, as described in the manual method in section 3.1.2. Record swimming videos of one worm at the time and use the worm tracker software to calculate the frequency of body bends.
   3. Follow the script provided with the tracking software to obtain worm thrashing frequency and to generate heat maps from the worm thrashing data.

Results

We present an example of SWIP assay induced by AMPH treatment. Figure 1 shows a schematic representation of the assay setup as described above. For the manual assay, about 8-10 age synchronized late L4 stage worms are collected with an eyelash or platinum pick and placed into a glass spot plate filled with 40 µL of 200 mOsm/L sucrose (control solution) or sucrose with 0.5 mM AMPH and tested for SWIP.

Discussion

Here, we describe a step-by-step protocol to perform a behavioral assay, SWIP, in C. elegans. This protocol is simple and straightforward with no major technical hurdles making this assay very user friendly. Nevertheless, there are some critical aspects that need to be considered in order to effectively perform the assay.

Care should be taken to ensure that the worms used for the assay are well fed, since dietary restriction affects SWIP¹⁷. Gentle handling of w...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Aluminum foil	Reynolds wrap	1091835
Amphetamine	Sigma	51-63-8
Autoclave
Bacterial Incubator	New Brunswick scientific	M1352-0000
Bacteriological grade, Agar	Lab Scientific, Inc	A466
Bacto (TM) Peptone	BD	REF 211677
Calcium Chloride (dihydrate)	Sigma-Aldrich	C3881
Camera	Thorlabs	U-CMAD3
Centrifuge	Eppendorf 5810R 15amp	E215059
Cholesterol	Sigma-Aldrich	57-88-5
Deionized water	Millipore	Z00QSV0WW	Milli-Q
Depression glass spot plate	Corning	Corning, Inc. 722085
Erlenmeyer flask	ThermoFisher	4103-0250PK
Eye lash
Glass slide	Fisherbrand	12-550-15
Graphing and statistical software	Prism		Graphpad 5
HEPES	Sigma-Aldrich	RB=H3375 & H7006
Hypochlorite	Hawkins	Sodium Hypochlorite 4-6%, USP" 1 gal
LB Broth, Miller	Fisher	BP1426
Magnesium Chloride (Hexahydrate)	Sigma-Aldrich	RB=M0250	500 g
Magnesium sulfate (heptahydrate)	Sigma-Aldrich	M1880
Magnetic stir bar	Fisherbrand	16-800-510
Microcentrifuge tubes	ThermoFisher	69715
NA 22 bacteria	CGC
Nystatin	Sigma	1400-61-9
Osmometer	Advanced Instruments, Inc	Model 3320
Pasteur Pipettes	Fisherbrand	13-678-20A
Petriplates	Falcon	351007
pH Meter	Orion VersaStar Pro	IS-68X591202-B 0514
Polystrine conical tubes	Falcon	352095
Potassium Chloride	Sigma-Aldrich	P9541
Potassium dihydrogen phosphate	Sigma-Aldrich	7778-77-0
Potassium Phosphate - DIBASIC	Sigma-Aldrich	P-8281
Potassium Phosphate - MONOBASIC	Sigma-Aldrich	P0662
Serological pipettes	VWR	10ml=89130-898
Shaker	Reliable Scientific	55S 12x16
Sodium Chloride	Fisher	RB=BP358-1
Sodium dihydrogen Phosphate	Fisher	RB=S381
Spreadsheet	MS office		Microsoft Excel
Stereo Microscope	Zeiss	Model tlb3. 1 stemi2000
Sterile Pipette tips	Various	02-707-400
Sucrose	Sigma-Aldrich	RB=S5016
Superglue	Loctite	1647358 .14 oz.
SwimR sofware	10.18129/B9.bioc.SwimR
Tracker 2	Worm Tracker 2.0	www.mrc-lmb.cam.ac.uk/wormtracker/
Video recording software	Virtualdub	http://www.virtualdub.org/