Name: swimming induced paralysis to assess dopamine signaling in caenorhabditis elegans
Uploaded: 2019-04-03T13:00:00.000+00:00
Duration: 7 min 36 s
Description: Watch this Scientific Journal Video about Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans at JoVE.com

The authors would like to thank Dr. Osama Refai from Dr. Randy Blakely&#39;s lab for guidance with the automated analysis of SWIP. This work was supported by funding from NIH R01 DA042156 to LC.

1. Preparation of Solutions and Media
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Prepare fresh sodium hypochlorite/NaOH solution by adding 1 mL of 5-6% sodium hypochlorite (bleach) and 180 &#181;L of 10 N NaOH to 3.8 mL of deionized water.</li>
	<li>Weigh 60 g sucrose and dissolve in autoclaved deionized water to a final volume of 100 mL to make 60% sucrose solution.</li>
	<li>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 &#176;C.</li>
	<li>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 &#181;L of the stock solution in 400 &#181;L of water to make 0.5 mM working solution.</li>
	<li>Prepare Nutrient growth media (NGM) plates
	<ol>
		<li>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 &#176;C, 15 PSI) for 1 hour using liquid cycle.</li>
		<li>Cool to and maintain the temperature at about 50 &#176;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).</li>
		<li>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 &#176;C in a box for up to 4 weeks.</li>
	</ol>
	</li>
	<li>Preparation of Lysogeny broth (LB) broth
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Preparation of NA22 bacterial plates
	<ol>
		<li>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 &#176;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 &#176;C on a shaking platform.</li>
		<li>To seed the plates, dispense 200 &#181;L 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 &#176;C.</li>
	</ol>
	</li>
	<li>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.</li>
</ol>
2. C. elegans Husbandry
NOTE: Culture wild-type N2C. elegans strain on Escherichia coli NA22 plates. The detailed culture methods are described below.
<ol>
	<li>Preparation of worm culture
	<ol>
		<li>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 &#176;C for 3-4 days. Under a stereo microscope, visually confirm the presence of gravid adults.</li>
	</ol>
	</li>
	<li>Preparation of synchronized population of worms
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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).</li>
		<li>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).</li>
		<li>Put a drop (2-50 &#181;L) 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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#181;L 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.</li>
		<li>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 &#176;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.</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>Manual method to test for SWIP
	<ol>
		<li>Aliquot 40 &#181;L 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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Automated analysis of SWIP
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Follow the script provided with the tracking software to obtain worm thrashing frequency and to generate heat maps from the worm thrashing data.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

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 worms while picking as well as timed sodium hypochlorite/NaOH treatment during lysis are critical steps, as trauma during picking (more common when using a platinum pick) or extended exposure of embryos to sodium hypochlorite/NaOH solution can cause permanent damage to the worms18 and thus compromise their ability to swim.
Sterile techniques should be followed to avoid contamination. Contamination of the agar plates compromises the health of the animals and thus alters their ability to swim. Slight difference in the percentage of animals exhibiting SWIP can be observed using agar plates seeded with different strains of E. coli bacteria (NA22, OP50, etc.). In our protocol, we use NA22 to yield a large number of worms.
Glass-spot plates, used during the SWIP assay, are preferred to plastic plates because they can be thoroughly washed, autoclaved and re-used when different types of drugs are tested.
Another important factor to be considered in a SWIP assay is the osmolarity of the liquid media in which the animals are tested19. In our protocol, we use sucrose to bring the osmolarity of water up to 200 mOsm/L which was previously shown to be an optimal condition for the animals (Blakely RD. personal communication). Water with a controlled osmolarity is preferred because it eliminates possible differences in water quality over multiple assays performed in different days, weeks or months. Notably, dat-1 and wild-type animals treated with AMPH do not exhibit SWIP if salty solutions (e.g., M9 solution) are used as control media.
The time required for most of the animals to exhibit SWIP can slightly change among different worm stages (L1-L4). For example, Masoudi et al. (2014) reported that after 5 min 80% of dat-1 L1 animals still swim, thus only 20% of animals exhibit SWIP. On the other hand, only 50% of L4 dat-1 animals still swim after 5 min20. Therefore, it is important that animals tested for SWIP are assayed at the same age. We have optimized our assay using always late L4 staged animals. Late L4 larvae have the advantage of being easily recognizable among the other larval stages because, at this stage, animals have reached their adult size and exhibit a characteristic thin line dividing the white spot in the center of their body that will later differentiate into a mature vulva. The time-span of the assay is also critical. For example, after 15 minutes 90% of L4 dat-1 mutants exhibit SWIP but at later time (30 min), only 60% of them exhibit SWIP20.
The automated SWIP assay eliminates human errors and improves high-throughput screening with respect to manual assays. However, tracking software programs are time consuming since they can only track a single worm at a time.
With respect to other C. elegans DA-dependent behaviors, SWIP is a less time-consuming type of assay. For instance, the basal slowing response2 is not an immediate-type of assay. In fact, worms need to be chronically fed with the drugs, and this could result in penetrant and off-target effects. Thus, it might not be as effective to screen for drugs.
One of the major applications of SWIP is to screen for various drugs that target the dopaminergic pathway. In cases where the drugs are not water soluble (e.g., mazindol), proper dilutions should be performed to achieve concentrations where the carrier solution is not toxic to the worms. Moreover, if different concentrations of the same drug are used5,13,14, dose-response curves can also be analyzed. For example, the rate of progression (slope of the curve of animals per minute, Figure 2) can be reported as function of the concentration and used to compare the effects of different drugs (e.g., AMPH vs cocaine).
When SWIP is used to investigate the mechanism of action of drugs at the dopaminergic synapses, attention should be paid if results are extended to other animals. For instance, imipramine, a specific inhibitor of the mammalian norepinephrine transporter (NET) has been used to induce DA-mediate SWIP in N2 animals4. C. elegans does not synthetize norepinephrine and consequently does not express NET. However, the C. elegans DA transporter shares homology with mammalian NET21. For this reason, drugs that are specific inhibitors of mammalian NET and have limited effects on mammalian DAT (e.g., imipramine) show high selectivity to C. elegans DAT. Thus, species selectivity might limit our ability to extrapolate findings from C. elegans to humans.
The most important factor to consider when designing SWIP assays is the inclusion of experiments proving that SWIP is mediated by the dopaminergic system. In fact, impaired swimming could be generated by factors other than genes related to the DA system (e.g., general defects in muscles contraction). To ensure that SWIP is indeed mediated by DA, protocols should include experiments performed with animals in which DA has been depleted. This can be achieved by either using knockout animals lacking expression of cat-2, the C. elegans homologue of the tyrosine hydroxylase, which is the rate-limiting enzyme for DA synthesis, or by pre-treating wild-type animals with reserpine, a drug that causes DA depletion from vesicles3. For instance, McDonald et al.3 showed that basal SWIP observed in dat-1 mutants was recovered when these animals were pre-treated with reserpine. This result suggests that SWIP is DA-mediated. On the other hand, using cat-2, dat-1 and mutants lacking expression of each of the dopaminergic receptors, Safratowich et al. (2014) demonstrated that the trace amine &#946;-phenylethylamine (&#946;PEA) induces SWIP within 1 minute of treatment independently from DA but by direct activation of the ligand-gated ion channel LGC-5514. The authors showed that &#946;PEA- and DA-induced SWIP are mechanistically different and they can be easily discriminated experimentally. In fact, while &#946;PEA-induced SWIP is DA- and dop-3-independent, it reaches maximal values within 1 min and rapidly decreases after 2 min14, DA-mediated SWIP is cat-2 and dop-3 dependent and is essentially zero after 1 minute (Figure 2). Thus, there is a large difference in the time required to reach maximal effects, and this allows to quickly discriminate between the two phenomena: 1) the fast &#946;PEA-induced SWIP within 1 minute obtained by direct activation of the LGC-55 channels and 2) the slow DA-mediated SWIP which occurs when a surplus of extracellular DA builds up over time (10-15 min) and the DA receptors DOP-3 are overstimulated3.
In conclusion, with the right set of experiments, which include the use of knockout animals for key player genes of the dopaminergic system (cat-2, dat-1, dop-3) and the use of drugs depleting DA storages (reserpine), SWIP has been successfully used to elucidate the mechanism of action of drugs like AMPH5,13, &#946;PEA14,15 and azaperone6.

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 synapses3,4,7 and to be employed for forward genetic screens for the identification of novel regulatory pathways involved in DA signaling7,8,9,10,11,12. 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 synapses5,6,13,14,15.
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.

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

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.

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 &#181;L of 200 mOsm/L sucrose (control solution) or sucrose with 0.5 mM AMPH and tested for SWIP.
When animals stop swimming (i.e., exhibit SWIP), they quickly sink to the bottom of the well and do not move. Therefore, the discrimination between animals that still swim on the surface of the water versus the ones that are steady at the bottom of the well is very straightforward. Most of the worms tested in control solution swim continuously for at least 10 minutes, whereas under AMPH treatment, the number of animals exhibiting SWIP progressively increases. The maximal percentage of animals exhibiting SWIP is proportional to the concentration of AMPH used1,5,13. When DAT-1 knockout (dat-1) worms are tested in control solution, 40-70% worms exhibit SWIP within 10 minutes4,13. This result is comparable to the percentage of paralyzed animals measured in wild-type animals treated with 0.5 mM AMPH (Figure 2).
Worms exposed to either sucrose or sucrose containing AMPH do not show SWIP in the first minute of observation (Figure 2). However, while worms treated with sucrose continue to swim for 10 minutes, worms treated with AMPH start to exhibit SWIP after 2 minutes of treatment and after 10 minutes, 66 &#177; 3% animals show SWIP (Figure 2).
For the automated analysis, videos of worms under control or AMPH treatments are recorded one worm at a time using a video recording software. A computer tracking software is used to track worm thrashing and the resulting data are imported into and analyzed with the software suit. Samples of heat maps of animals exposed to control or AMPH, displaying actively moving animals in red and paralyzed worms in green, are shown in Figure 3.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59243/59243fig1.jpg" /> 
Figure 1: Assay set up for SWIP. Gravid adult wild-type (N2) worms were lysed with sodium hypochlorite/NaOH treatment to release embryos. The embryos were allowed to hatch and develop into synchronized L1 larvae in M9 buffer for 14 h on a shaker and then plated on an NGM plate seeded with NA22 bacteria. After 42-48 h late L4 stage larvae were visually identified under the stereoscope and picked with an eyelash pick into a spot plate with or without amphetamine in control sucrose solution and scored for SWIP either manually or through automated analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59243/59243fig1large.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/59243/59243fig2.jpg" /> 
Figure 2: Amphetamine-induced SWIP using manual assay. Worms in sucrose or sucrose with 0.5 mM amphetamine (AMPH) were visually scored for SWIP behavior every minute using a stereoscope. The percent of animals exhibiting SWIP was calculated by dividing the number of paralyzed worms by the total number of worms assayed for each time point, and then multiplying the result by 100. The percent of worms exposed to AMPH (blue squares) showing SWIP increases overtime, while the untreated worms (red circles) continue to swim during the 10-minute window. N represents the number of animals tested in each group. Error bars indicate standard error of means (SEM). Statistical significance was assessed by performing two-way ANOVA with Bonferroni multiple comparison test (p &#60; 0.0001). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59243/59243fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59243/59243fig3.jpg" /> 
Figure 3: Amphetamine-induced SWIP via automated analysis. Videos of worms in sucrose or sucrose with 0.5 mM AMPH were recorded using a camera mounted on a stereoscope. Swimming videos of individual worms were tracked with a tracking software and analyzed using tracking software suit. Heat maps were generated from the data where red areas show the worms that are actively moving, and green areas indicate paralyzed worms. Each experimental group is representative of 6 animals. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59243/59243fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans at JoVE.com

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.

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) ...

swimming-induced-paralysis-to-assess-dopamine-signaling

Nanyang Technological University

Research

JoVE Journal

Behavior

7.7K Views.  Florida Atlantic University, John D MacArthur Campus. Het algemene doel van deze procedure is om aan te tonen Zwemmen-Geïnduceerde Verlamming of SWIP, een gedragstest om dopamine signalering in C.elegans te beoordelen. SWIP maakt het mogelijk om genen binnen of gerelateerd aan de dopaminerge synapsis snel te identificeren. Deze methode maakt het mogelijk het onderzoek van de effecten geproduceerd door een overmaat aan extracellulaire dopamine die wordt veroorzaakt door bijvoorbeeld drugs of door mutaties bij de dopamine transporter.Om er...

Video: Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans

Assaying &beta;-amyloid Toxicity using a Transgenic C. elegans Model

Accumulation of the &beta;-amyloid peptide (A&beta;) is generally believed to be central to the induction of Alzheimer's disease, but the relevant mechanism(s) of toxicity are still unclear. A&beta; is also deposited intramuscularly in Inclusion Body Myositis, a severe human myopathy. The intensely studied nematode worm Caenorhabditis elegans can be transgenically engineered to express human A&beta;. Depending on the tissue or timing of A&beta; expression, transgenic worms can have readily measurable phenotypes that serve as a read-out of A&beta; toxicity. For example, transgenic worms with pan-neuronal A&beta; expression have defects is associative learning (Dosanjh et al. 2009), while transgenic worms with constitutive muscle-specific expression show a progressive, age-dependent paralysis phenotype (Link, 1995; Cohen et al. 2006). One particularly useful C. elegans model employs a temperature-sensitive mutation in the mRNA surveillance system to engineer temperature-inducible muscle expression of an A&beta; transgene, resulting in a reproducible paralysis phenotype upon temperature upshift (Link et al. 2003). Treatments that counter A&beta; toxicity in this model [e.g., expression of a protective transgene (Hassan et al. 2009) or exposure to Ginkgo biloba extracts (Wu et al. 2006)] reproducibly alter the rate of paralysis induced by temperature upshift of these transgenic worms. Here we describe our protocol for measuring the rate of paralysis in this transgenic C. elegans model, with particular attention to experimental variables that can influence this measurement.

Assaying &#946;-amyloid Toxicity using a Transgenic C. elegans Model

The intensely studied nematode worm Caenorhabditis elegans can be transgenically engineered to express the human &beta;-amyloid peptide (A&beta;). Induced expression of A&beta; in C. elegans muscle leads to a rapid, reproducible paralysis phenotype that can be used to monitor treatments that modulate A&beta; toxicity.

Accumulation of the &beta;-amyloid peptide (A&beta;) is generally believed to be central to the induction of Alzheimer's disease, but the relevant ...

Neuroscience

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and pathways as well as its ease of cultivation. Several C. elegans disease models have been reported, including neurodegenerative disorders such as Parkinson's disease (PD), which involves the degeneration of dopaminergic (DA) neurons 1. Both transgenes and neurotoxic chemicals have been used to induce DA neurodegeneration and consequent movement defects in worms, allowing for investigations into the basis of neurodegeneration and screens for neuroprotective genes and compounds 2,3.
Screens in lower eukaryotes like C. elegans provide an efficient and economical means to identify compounds and genes affecting neuronal signaling. Conventional screens are typically performed manually and scored by visual inspection; consequently, they are time-consuming and prone to human errors. Additionally, most focus on cellular level analysis while ignoring locomotion, which is an especially important parameter for movement disorders.
We have developed a novel microfluidic screening system (Figure 1) that controls and quantifies C. elegans' locomotion using electric field stimuli inside microchannels. We have shown that a Direct Current (DC) field can robustly induce on-demand locomotion towards the cathode ("electrotaxis") 4. Reversing the field's polarity causes the worm to quickly reverse its direction as well. We have also shown that defects in dopaminergic and other sensory neurons alter the swimming response 5. Therefore, abnormalities in neuronal signaling can be determined using locomotion as a read-out. The movement response can be accurately quantified using a range of parameters such as swimming speed, body bending frequency and reversal time.
Our work has revealed that the electrotactic response varies with age. Specifically, young adults respond to a lower range of electric fields and move faster compared to larvae 4. These findings led us to design a new microfluidic device to passively sort worms by age and phenotype 6.
We have also tested the response of worms to pulsed DC and Alternating Current (AC) electric fields. Pulsed DC fields of various duty cycles effectively generated electrotaxis in both C. elegans and its cousin C. briggsae 7. In another experiment, symmetrical AC fields with frequencies ranging from 1 Hz to 3 KHz immobilized worms inside the channel 8.
Implementation of the electric field in a microfluidic environment enables rapid and automated execution of the electrotaxis assay. This approach promises to facilitate high-throughput genetic and chemical screens for factors affecting neuronal function and viability.

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

A semi-automated micro-electro-fluidic method to induce on-demand locomotion in Caenorhabditis elegans is described. This method is based on the neurophysiologic phenomenon of worms responding to mild electric fields (&#x201C;electrotaxis&#x201D;) inside microfluidic channels. Microfluidic electrotaxis serves as a rapid, sensitive, low-cost, and scalable technique to screen for factors affecting neuronal health.

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and ...

Bioengineering

Paradigms for Pharmacological Characterization of C. elegans Synaptic Transmission Mutants

The nematode, Caenorhabditis elegans, has become an expedient model for studying neurotransmission. C. elegans is unique among animal models, as the anatomy and connectivity of its nervous system has been determined from electron micrographs and refined by pharmacological assays. In this video, we describe how two complementary neural stimulants, an acetylcholinesterase inhibitor, called aldicarb, and a gamma-aminobutyric acid (GABA) receptor antagonist, called pentylenetetrazole (PTZ), may be employed to specifically characterize signaling at C. elegans neuromuscular junctions (NMJs) and facilitate our understanding of antagonistic neural circuits.
Of 302 C. elegans neurons, nineteen GABAergic D-type motor neurons innervate body wall muscles (BWMs), while four GABAergic neurons, called RMEs, innervate head muscles. Conversely, thirty-nine motor neurons express the excitatory neurotransmitter, acetylcholine (ACh), and antagonize GABA transmission at BWMs to coordinate locomotion. The antagonistic nature of GABAergic and cholinergic motor neurons at body wall NMJs was initially determined by laser ablation and later buttressed by aldicarb exposure. Acute aldicarb exposure results in a time-course or dose-responsive paralysis in wild-type worms. Yet, loss of excitatory ACh transmission confers resistance to aldicarb, as less ACh accumulates at worm NMJs, leading to less stimulation of BWMs. Resistance to aldicarb may be observed with ACh-specific or general synaptic function mutants. Consistent with antagonistic GABA and ACh transmission, loss of GABA transmission, or a failure to negatively regulate ACh release, confers hypersensitivity to aldicarb. Although aldicarb exposure has led to the isolation of numerous worm homologs of neurotransmission genes, aldicarb exposure alone cannot efficiently determine prevailing roles for genes and pathways in specific C. elegans motor neurons. For this purpose, we have introduced a complementary experimental approach, which uses PTZ.
Neurotransmission mutants display clear phenotypes, distinct from aldicarb-induced paralysis, in response to PTZ. Wild-type worms, as well as mutants with specific inabilities to release or receive ACh, do not show apparent sensitivity to PTZ. However, GABA mutants, as well as general synaptic function mutants, display anterior convulsions in a time-course or dose-responsive manner. Mutants that cannot negatively regulate general neurotransmitter release and, thus, secrete excessive amounts of ACh onto BWMs, become paralyzed on PTZ. The PTZ-induced phenotypes of discrete mutant classes indicate that a complementary approach with aldicarb and PTZ exposure paradigms in C. elegans may accelerate our understanding of neurotransmission. Moreover, videos demonstrating how we perform pharmacological assays should establish consistent methods for C. elegans research.

This video demonstrates how to employ two neural stimulants, aldicarb and pentylenetetrazole (PTZ), in complementary ways to study synaptic function in the nematode, C. elegans. This complementary approach may also be used to shed light on evolutionarily conserved mechanisms for modulating neuronal synchrony and has implications for epilepsy and seizures.

The nematode, Caenorhabditis elegans, has become an expedient model for studying neurotransmission. C. elegans is unique among animal models, as the ...

Biology

Measuring Caenorhabditis elegans Life Span on Solid Media

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

Measuring Caenorhabditis elegans Life Span on Solid Media

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

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

Measuring Caenorhabditis elegans Sensitivity to the Acetylcholine Receptor Agonist Levamisole

At the neuromuscular junction (NMJ), the binding of the excitatory neurotransmitter acetylcholine (ACh) to postsynaptic receptors leads to muscle contraction. As in vertebrate skeletal muscle, cholinergic signaling in the body wall muscles of the model organism Caenorhabditis elegans is required for locomotion. Exposure to levamisole, a pharmacological agonist of one class of ACh receptors on the body wall muscles, causes time-dependent paralysis of wild-type animals. Altered sensitivity to levamisole suggests defects in signaling at the NMJ or muscle function. Here, a protocol for a liquid levamisole assay performed on C. elegans grown in 24-well plates is presented. Vigorous swimming of the animals in liquid allows for the assessment and quantitation of levamisole-induced paralysis in hundreds of worms over a one-hour time period without requiring physical manipulation. This procedure can be used with both wild-type and mutants that have altered sensitivity to levamisole to demonstrate the functional consequences of altered signaling at the NMJ.

Measuring Caenorhabditis elegans Sensitivity to the Acetylcholine Receptor Agonist Levamisole

The present protocol describes an assay to determine response to levamisole, a pharmacological agonist of one class of Caenorhabditis elegans acetylcholine receptors. In this liquid levamisole swim assay, researchers visually observe and quantitate the time-dependent paralysis of animals cultivated in 24-well plates.

At the neuromuscular junction (NMJ), the binding of the excitatory neurotransmitter acetylcholine (ACh) to postsynaptic receptors leads to muscle ...

Genetics

Automated Analysis of C. elegans Swim Behavior Using CeleST Software

Dissecting the neuronal and neuromuscular circuits that regulate behavior remains a major challenge in biology. The nematode Caenorhabditis elegans has proven to be an invaluable model organism in helping to tackle this challenge, from inspiring technological approaches,&#160;building the human brain connectome, to actually shedding light on the specific molecular drivers of basic functional patterns. The bulk of the behavioral studies in C. elegans have been performed on solid substrates. In liquid, animals exhibit behavioral patterns that include movement at a range of speeds in 3D, as well as partial body movements, such as a posterior curl without anterior shape change, which introduce new challenges for quantitation. The steps of a simple procedure, and use of a software that enables high-resolution analysis of C. elegans swim behavior, are presented here. The software, named CeleST, uses a specialized computer program that tracks multiple animals simultaneously and provides novel measures of C. elegans locomotion in liquid (swimming). The measures are mostly grounded in animal posture and based on mathematics used in computer vision and pattern recognition, without computational requirements for threshold cut-offs. The software tool can be used to both assess overall swimming prowess in hundreds of animals from combined small batch trials and to reveal novel phenotypes even in well-characterized genetic mutants. The preparation of specimens for analysis with CeleST is simple and low-tech, enabling wide adaptation by the scientific community. Use of the computational approach described here should therefore contribute to the greater understanding of behavior and behavioral circuits in the C. elegans model.

Automated Analysis of C. elegans Swim Behavior Using CeleST Software

An efficient and simple methodology for computer-based analysis of nematode swimming behavior in liquid is described. The method requires little to no investment for C. elegans laboratories. The hardware used is standard, and the computer software for the behavioral analysis (CeleST) is an open source one.

Dissecting the neuronal and neuromuscular circuits that regulate behavior remains a major challenge in biology. The nematode Caenorhabditis elegans has ...

Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis

The neurodegenerative disease amyotrophic lateral sclerosis (ALS) features progressive loss of motor neurons accompanied by muscle weakness and motor impairment that worsens with time. While considerable advances have been made in determining genetic drivers of ALS for a subset of patients, the majority of cases have an unknown etiology. Further, the mechanisms underlying motor neuron dysfunction and degeneration are not well understood; therefore, there is an ongoing need to develop and characterize representative models to study these processes. Caenorhabditis elegans can adapt their movement to the physical constraints of their surroundings, with two primary movement paradigms studied in a laboratory environment- crawling on a solid surface and swimming in liquid. These represent a complex interplay between sensation, motor neurons, and muscles. C. elegans models of ALS can exhibit impairment in one or both of these movement paradigms. This protocol describes two sensitive assays for evaluating motility in C. elegans: an optimized radial locomotion assay measuring crawling on a solid surface and an automated method for tracking and analyzing swimming in liquid (thrashing). In addition to the characterization of baseline motor impairment of ALS models, these assays can detect suppression or enhancement of the phenotypes from genetic or small molecule interventions. Thus, these methods have utility for studying ALS&#160;models and any C. elegans strain that exhibits altered motility.

Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis

This protocol describes two sensitive assays for discriminating among mild, moderate, and severe motor impairment in C. elegans models of amyotrophic lateral sclerosis, with general utility for C. elegans strains, with altered motility.

The neurodegenerative disease amyotrophic lateral sclerosis (ALS) features progressive loss of motor neurons accompanied by muscle weakness and motor ...

Quantifying Tissue-Specific Proteostatic Decline in Caenorhabditis elegans

The ability to maintain proper function and folding of the proteome (protein homeostasis) declines during normal aging, facilitating the onset of a growing number of age-associated diseases. For instance, proteins with polyglutamine expansions are prone to aggregation, as exemplified with the huntingtin protein and concomitant onset of Huntington&#8217;s disease. The age-associated deterioration of the proteome has been widely studied through the use of transgenic Caenorhabditis elegans expressing polyQ repeats fused to a yellow fluorescent protein (YFP). This polyQ::YFP transgenic animal model facilitates the direct quantification of the age-associated decline of the proteome through imaging the progressive formation of fluorescent foci (i.e., protein aggregates) and subsequent onset of locomotion defects that develop as a result of the collapse of the proteome. Further, the expression of the polyQ::YFP transgene can be driven by tissue-specific promoters, allowing the assessment of proteostasis across tissues in the context of an intact multicellular organism. This model is highly amenable to genetic analysis, thus providing an approach to quantify aging that is complementary to lifespan assays. We describe how to accurately measure polyQ::YFP foci formation within either neurons or body wall muscle during aging, and the subsequent onset of behavioral defects. Next, we highlight how these approaches can be adapted for higher throughput, and potential future applications using other emerging strategies for C. elegans genetic analysis.

Quantifying Tissue-Specific Proteostatic Decline in Caenorhabditis elegans

Proteostatic decline is a hallmark of aging, facilitating the onset of neurodegenerative diseases. We outline a protocol to quantifiably measure proteostasis in two different Caenorhabditis elegans tissues through heterologous expression of polyglutamine repeats fused to a fluorescent reporter. This model allows rapid in vivo genetic analysis of proteostasis.

The ability to maintain proper function and folding of the proteome (protein homeostasis) declines during normal aging, facilitating the onset of a ...

Swimming-Induced Paralysis (SWIP) Assay: A Method to Quantify Dopamine-Mediated Locomotion in C. elegans

Source: Kudumala, S., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/59243/swimming-induced-paralysis-to-assess-dopamine-signaling">Swimming Induced Paralysis to Assess Dopamine Signaling in&#160;Caenorhabditis elegans</a>.&#160;J. Vis. Exp.&#160;(2019).
Here, we introduce an assay to measure SWIP, a behavioral phenotype of&#160;C. elegans&#160;that occurs during vigorous motion like swimming.&#160; The example protocol discusses both manual and automated analysis approaches.

Source: Kudumala, S., et al. Swimming Induced Paralysis to Assess Dopamine Signaling in&#160;Caenorhabditis elegans.&#160;J. Vis. Exp.&#160;(2019).
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JoVE Encyclopedia of Experiments

Caenorhabditis elegans (worm)

Evaluating Motor Impairment in C. elegans Models Using the Radial Locomotion Assay

Source: Currey, H. N., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/62699">Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis.</a> J. Vis. Exp. (2021).
This video demonstrates the use of a radial locomotion assay to evaluate motor dysfunction in C. elegans expressing either wild-type or ALS-mutant human TDP-43 protein. Worms with varying levels of wild-type or mutant TDP-43 expression are allowed to crawl, and their movement is measured by the radial distance traveled from the central point on the assay plate. The assay distinguishes between mild, moderate, and severe motor impairments based on the distance traveled by the different C. elegans strains.

Source: Currey, H. N., et al. Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis. J. Vis. Exp. (2021).
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Swimming Induced Paralysis to Assess Dopamine Signaling in Caenorhabditis elegans

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