I thank Dr. Maureen Barr, under whose supervision the dauer IL2 remodeling phenotype was initially characterized. Thanks to Dr. C. Britt Carlson for critical reading of the manuscript. Funding in the Barr lab was from the NIH (5R01DK59418) and postdoctoral fellowships from the USDA (2010-65106-20587) and the New Jersey Commission on Spinal Cord Research (CSCR12FEL004). Current funding for this work is from a University of Illinois Urbana-Champaign College of ACES FIRE grant.

1. Crude Dauer Pheromone Production
<ol>
	<li>Grow OP50 E. coli from a single colony O/N at 37 &#176;C while shaking in 100 ml of LB broth at 200 rpm. NOTE: This O/N culture can be made in advance and stored at 4 &#176;C.</li>
	<li>Grow N2 C. elegans on 15 - 20 6 cm Petri dishes with NGM agar (See recipe in Table 1) seeded with 50 &#181;l of OP50 E. coli until the bacteria is almost depleted19.</li>
	<li>Make 5x 250 ml of S media (See recipe in Table 1) in 1 L Erlenmeyer flasks19. NOTE: This solution can be made in advance.</li>
	<li>Grow a culture of OP50 E. coli in 4 L of LB broth O/N by inoculating with 1 ml of OP50 in E. coli and shaking at 37 &#176;C for 16 hr.</li>
	<li>Wash the nematodes from plates (step 1.2) with 1 ml of S basal solution (See recipe in Table 1) and transfer nematodes from 4-5 plates into each of 4 1 L Erlenmeyer flasks with the S media prepared in Step 1.3 (4-5 plates for each flask)19.</li>
	<li>Centrifuge the OP50 from step 1.4 at 650 x g for 10 min and remove supernatant. Resuspend each bacterial pellet in 10 ml of S basal solution. Transfer equal amounts of resuspended bacteria to each flask of nematodes.</li>
	<li>Place the flasks on an orbital shaker at room temperature (20 - 25 &#176;C) and 100 - 150 rpm for 4 - 5 days until the solution begins to clear, indicating a depletion of food.</li>
	<li>Once the food is depleted, grow another O/N culture of OP50 E. coli in 4x 1 L of LB broth. Centrifuge the bacteria at 650 x g for 10 min and add a pellet from one flask to each of the S media flasks with nematodes. Return the S media flasks to the orbital shaker, as in step 1.7, for an additional 3-4 days until the food is depleted again.</li>
	<li>Take a ~1 ml aliquot of solution from each flask and estimate the nematode concentration by sub-sampling 1 &#181;l from each 1 ml aliquot and counting the number of nematodes. NOTE: The nematode population should be 50,000 - 75,000 nematodes/ml. At this concentration the majority of nematodes are dauers.</li>
	<li>Discard any contaminated flasks. NOTE: Contamination frequently manifests as fungal hyphae seen during step 1.9.</li>
	<li>Centrifuge nematodes at 4,000 x g for 10 min at 4 &#176;C. Discard the nematode pellet and retain the supernatant. Alternatively, see the discussion for using these animals to make a low-potency pheromone.</li>
	<li>Filter the supernatant through #1 Whatman filter paper. NOTE: A vacuum flask will expedite this process.</li>
	<li>Further filter the supernatant through 0.45 &#181;m sterile vacuum filtration units. NOTE: Filtrate can be kept O/N at 4 &#176;C or proceed to 1.14.</li>
	<li>Add the filtrate to a 2 L beaker with a stir bar. Place in a fume hood on a heating stir plate. Bring to a boil while stirring and evaporate to approximately 200 ml. Transfer the solution to a 500 ml beaker. Continue boiling until approximately 50 ml are left. NOTE: Care should be taken during this step as it is possible for the solution to boil over. The color of the final 50 ml is a dark brown. At any time during the cooking process, the heat can be turned off and the filtrate stored O/N at RT or 4 &#176;C.</li>
	<li>Let cool and centrifuge for 5 min at 1,000 x g. Pour the supernatant into a 100 ml beaker and discard the pellet.</li>
	<li>Boil the supernatant down to a brown crust. Remove from heat and break up the crust with a spatula. Add sufficient 200 proof ethanol to cover the crust. Cover the beaker with Parafilm and let it sit O/N at RT. Pour off the ethanol into a clean beaker and put to the side. Add fresh ethanol to cover the crust. Repeat O/N extractions 3 times. Pool the extracts.</li>
	<li>Use a vacuum pump to dry the ethanol extracts. If a vacuum pump is not available, add the ethanol extract to a clean beaker and warm to 50 &#176;C while stirring in a fume hood. NOTE: Once the ethanol has evaporated a sticky residue of light brown color should remain. If heating is used, care should be taken to not burn the solution by heating for too long.</li>
	<li>Dissolve residue in 10 ml of distilled sterile water. Filter sterilize this solution with a 0.22 &#181;m syringe filter and store at -20 &#176;C in 1 ml aliquots. NOTE: 1 ml of water is used for dissolving the residue for every 100 ml of the original liquid nematode culture. For example, if one of the liquid culture flasks is discarded due to contamination, then adjust the amount of water used to dissolve the residue.</li>
</ol>
2. Determination of Pheromone Dose Response
<ol>
	<li>Prepare dauer pheromone by making 6x 5 ml of NGM media with Noble agar, without peptone, and supplemented with 50 &#181;g/ml streptomycin sulfate and a range of dauer pheromone (see recipe in Table 1)4, 19. Mix components in a 13x100 mm glass culture tube, boil over a Bunsen burner, and then pour into 3 cm Petri dishes.</li>
	<li>Weigh a microcentrifuge tube. Add 1 ml of O/N culture of E. coli OP50 to the tube and centrifuge at ~17,000 x g in a microcentrifuge tube. Remove all of the supernatant. Weigh the tube again and determine the weight of the bacterial pellet. Resuspend the pellet with M9 buffer (See recipe in Table 1) for a final concentration of 4% (w/v). Aliquot 10 &#181;l of the bacterial suspension onto the center of each pheromone plate4. Store plates O/N at RT.</li>
	<li>Pick 10-15 gravid N2 wild-type hermaphrodites one day after the molt into adulthood onto the pheromone plates4. Store the plates at 25 &#176;C for 3 - 4 hr.</li>
	<li>Pick off all of the adult hermaphrodites. Record the number of eggs laid for each plate. Seal the plates with Parafilm and return to 25 &#176;C for 3 days17.</li>
	<li>Count the percentage of dauers (Figure 2). Note: For both methods it is important to note that dauers (and to a lesser extent non-dauers) will climb up the walls and onto the lid of Petri dishes. It is therefore important to examine all surfaces of the dish for nematodes.</li>
	<li>Fit the data to a dose-response curve and estimate the EC90 concentration of pheromone (sufficient pheromone to induce 90% of animals to form dauers) as shown in Figure 3. Note: We typically use a logistic regression model to estimate the EC90, but different statistical software packages may have alternative methods that produce similar estimates.</li>
</ol>
3. Live Imaging of Dauer IL2 Neurons&#8203;
<ol>
	<li>Prepare dauer pheromone plates as in protocol 2 using the EC90 value determined in step 2.6.</li>
	<li>Induce dauers using the methods in protocol 2 with a strain carrying an integrated IL2 reporter transgene (myIs13[Pklp-6::gfp], myIs14[Pklp-6::gfp], myIs16[Pklp-6::tdTomato] or qIs56[Plag-2::gfp])12, 20, 21. NOTE: If qIs56 is used, expression of GFP will not be visible in the IL2Qs until several hours into the dauer molt and some of the initial remodeling events will not be visible. Additionally, while the fluorescent signal from the Plag-2::gfp transgene is ultimately brighter in the IL2s during dauer than the Pklp-6 fluorescentreporters, the Plag-2::gfp transgene also expresses weakly in the head muscles which can mask the fine IL2 branches.</li>
	<li>Following egg-laying, remove the adult hermaphrodites, seal the plates with Parafilm and incubate for 34 hr at 25 &#176;C17. NOTE: The length of time to the dauer molt varies slightly among individuals. The most rapid period of dendrite arborization begins 4 - 5 hr following the onset of the dauer molt (39 - 44 hr post egg-laying at 25 &#176;C)12.</li>
	<li>The day before imaging, make up a solution of 10% (w/v) agarose in M9 buffer with dauer pheromone at the EC90 concentration. Aliquot approximately 100 &#181;l of agarose onto a microscope slide and quickly, but gently, place another slide on top to create a flat agarose surface. After the agarose has solidified, wrap the slides in saran wrap and store in a humidity chamber (pipette tip box with wet paper towels).</li>
	<li>After the 34 hr incubation period, examine pheromone plates for nematodes that have stopped pharyngeal pumping. Separate slides such that only one surface of one slide has agarose on it. Pick one or more nematodes onto a 10% agarose pad and 1 &#181;l of 0.1 micron polystyrene beads22. NOTE: With a well-synchronized population, the majority of the nematodes on the plate will be entering into the molt from L2d to dauer. However, there may be variation among individuals. It is therefore important to note additional features (stoma occlusion, pharyngeal remodeling) to estimate the amount of time the animals have been within the dauer molt (see representative results for example).</li>
	<li>Determine the orientation (left/right, dorsal/ventral) of the nematode. Capture Z-stack images of an IL2Q dendrite at 100x magnification with the lowest possible fluorescence intensity every 15 min or as needed for the specific experiment. Continue capturing images until the nematode has separated from the L2d cuticle and immobilization is impossible or if the animal recovers from dauer (pharynx begins pumping). When not actively imaging, place the slide in a humidity chamber. NOTE: We use an epifluorescent microscope with a motorized stage and differential interference contrast (DIC) optics.</li>
</ol>

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The authors have nothing to disclose

An examination of dauer-specific neuroplasticity and other dauer-associated morphological changes requires controlled and reliable formation of dauers either through genetic mutation (i.e., daf-c mutants) or by exposure of wild-type animals to pheromone. While the use of a genetic mutation to induce dauers is convenient, it may confound results. Therefore data collected with daf-c mutations should be subsequently confirmed with wild-type animals induced to enter the dauer stage by exposure to dauer pheromone.
The production of crude dauer pheromone is relatively simple and variations of this protocol are published18. With the possible exception of contamination, the method presented here is very reliable. To produce the extract, we use a standard method of culturing large densities of nematodes in S media liquid culture19. There is a large amount of flexibility in the quantity of initial nematode inoculum added to the liquid culture. Starting with larger numbers of nematodes will expedite the procedure. However, it is important to ensure that all of the initial nematode inoculum is free of contamination. Both fungal and bacterial contamination may arise during growth in the liquid media. Fungal contamination can be easily detected by the presence of hyphae. Bacterial contamination is more difficult to assess. Typically a bacterial contaminant may be observed if the liquid culture does not clear soon after the listed incubation periods. There is flexibility in the total nematode population density needed to produce sufficient pheromone. For example, a concentration of 25,000 nematodes/ml is sufficient to produce a pheromone effective at inducing dauers. An alternative method consists of transferring the nematodes from a liquid culture to a small volume of M9 buffer (~50 ml) followed by an O/N incubation at RT on an orbital shaker and subsequent purification of the pheromone by the methods described herein. This alternative is effective, but produces a pheromone with a higher EC50 (i.e., less potent).
As each batch of crude pheromone may differ in potency, it is important to test a subsample for effectiveness in inducing dauer. For many applications a single bioassay with one replicate for each pheromone concentration is sufficient to provide a general estimate of potency. However, there can be significant variation between independently performed dose-response bioassays. Therefore, if experiments require a delicate balance between dauer formation and non-dauer development, it will be necessary to perform 2 - 3 replicate bioassays to determine an accurate effective dose. In the future, as the importance of ascarosides in nematode biology is more widely recognized, it may be possible to purchase synthesized dauer pheromone at reasonable cost.
The finding of dauer-specific neuronal remodeling in the IL2s provides a platform for studying the molecular mechanisms of stress-induced neuroplasticity. To identify the mechanism by which specific genes regulate neuroplasticity, it may be necessary to follow the remodeling process in real-time. The method of IL2 in vivo imaging builds upon previous C. elegans methods22, 24, 25. One obstacle to imaging during dauer formation is the possibility of dauer recovery. The easiest solution is to initially use a daf-c mutation. Confirmation of any results can then be performed in a non-daf-c background using the methods described herein.
Singh and Sulston noted several obstacles for the in vivo imaging of dauer formation25. These included newly formed dauers escaping off the microscope slide and uncontrolled movement following radial shrinkage. The use of microsphere beads has eliminated the possibility of escape22. However the radial shrinkage that occurs approximately 10 hr into the dauer molt still creates difficulties for subsequent imaging.

Phenotypic plasticity, including developmental and behavioral plasticity, is important for adaptations to adverse environmental conditions and is considered a driving force in evolution1. C. elegans is a model organism frequently used for studies in development and neurobiology. Under ideal conditions, C. elegans develops through four larval stages prior to entering the adult reproductive stage. However, under conditions of low food availability and high population densities, C. elegans can undergo a developmental switch and enter into a long-lived and stress-resistant dauer stage2. Dauers display several morphological and behavioral differences from non-dauers which likely mediate this stress resistance. The environmental cues and genetic pathways regulating dauer entry have been extensively described3, 4, 5, 6, 7. However, the molecular mechanisms controlling individual phenotypic changes that occur during dauer formation are less well understood.
The examination of dauer-specific phenotypes requires the production of dauer animals. This can be accomplished in three ways: 1) Picking from a starved culture of C. elegans, 2) Use of dauer formation constitutive (daf-c) mutants, 3) Induction of dauer formation through purified pheromone. It is relatively simple to pick dauers from a standard NGM plate with a C. elegans population that has exhausted its bacterial food-supply. In our hands, a standard NGM plate originally seeded with 50 &#181;l of OP50 Escherichia coli, allowed to grow for 3 days&#160;and subsequently inoculated with 3 - 4 adult hermaphrodites kept at 25 &#176;C will exhaust the food supply within one week and produce several thousand dauers within an additional 3 days (in addition to numerous starved non-dauers). However, this method is unsatisfactory for examining processes occurring during the molt into dauer. It is difficult to determine which of the several thousand animals on a typical starved plate are entering into dauer. Furthermore, use of a starved plate offers no possibility for synchronization. The use of daf-c mutants allows for synchronization and the reliable generation of dauers. However, there is no guarantee that a particular daf-c mutation will not affect other dauer-specific phenotypes alone or in combination with additional mutations.
The presence of a dauer pheromone was first implied by Cassada and Russell and later demonstrated by Golden and Riddle. The dauer pheromone has since been chemically characterized2, 8, 9, 10. The purified pheromone consists of a complex blend of ascarosides, which in different forms regulate both behavioral and developmental processes9, 11. Use of a crude dauer pheromone extract allows for reliable controlled induction of synchronized dauers in a wild-type background.
The nervous system and associated glial cells undergo remodeling during dauer larva formation12, 13, 14. Some of these changes are irreversible, such as the remodeling of glia cells during dauer13, 14. However other remodeling events are reversible upon a return to favorable environmental conditions. For example, a set of four IL2Q neurons undergo rapid and reversible neuronal remodeling during dauer formation including: dendrite arborization, a switch from single dendritic to multidendritic neurons and axon remodeling12. The methods herein allow for the reliable in vivo imaging of stress-induced neuroplasticity in a model organism. The methods presented for the production and testing of crude dauer pheromone are previously described and compiled from several sources4, 8, 15, 16, 17, 18.

<table border="0" cellpadding="0" cellspacing="0" width="873">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:259px;">N2 C. elegans Bristol strain</td>
			<td style="width:152px;">Caenorhabditis Genetics Center</td>
			<td style="width:212px;"></td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">PT2660 myIs13[Pklp-6::gfp] III</td>
			<td style="width:152px;">Upon request from Schroeder lab</td>
			<td></td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">PT2762 myIs14[Pklp-6::gfp] V</td>
			<td style="width:152px;">Upon request from Schroeder lab</td>
			<td></td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">JK2868 qIs56[Plag-2::gfp] V</td>
			<td style="width:152px;">Caenorhabditis Genetics Center</td>
			<td></td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">OP50 E. coli</td>
			<td style="width:152px;">Caenorhabditis Genetics Center</td>
			<td></td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Petri dishes</td>
			<td style="width:152px;">Tritech Research</td>
			<td>60 mm T3308, 35 mm T3501</td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="132">
			<td height="132" style="height:132px;">NGM agar</td>
			<td style="width:152px;">Various</td>
			<td></td>
			<td style="width:251px;">3 g NaCl, 17 g agar, 2.5 g peptone in 975 mL H2O autoclaved cooled to 55&#176;C followed by 1 mL of 5 mg/ml cholesterol in ethanol, 25 mL of 1 M KPO4 buffer (pH 6.0), 1 mL of 1 M CaCl2, 1 mL of 1 M MgSO4</td>
		</tr>
		<tr height="356">
			<td height="356" style="height:356px;">S Media</td>
			<td style="width:152px;">Various</td>
			<td></td>
			<td style="width:251px;">S media consists of S basal solution (1.46 g NaCl, 0.25g K2HPO4, 1.5 g KH2PO4, 0.25 ml of 5 mg/ml cholesterol in ethanol and H2O to 250 mL. Autoclaved.) To the S basal solution add 2.5 mL of potassium citrate pH 6.0 (2 g citric acid monohydrate, 29.4 g tri-potassium citrate monohydrate, H2O to 100 mL, autoclaved), 2.5 mL of Trace Metals Solution (1.86 g disodium EDTA, 0.69 g FeSO4&#160;&#8226;7 H2O, 0.2 g MnCl2&#8226;4 H2O, 0.29 g ZnSO4&#160;&#8226;7 H2O, 0.025 g CuSO4&#160;&#8226;5 H2O, H2O to 1 liter. Autoclaved and stored in the dark), 0.75 mL of 1 M CaCl2, and 0.75 mL of 1 M MgSO4.</td>
		</tr>
		<tr height="392">
			<td height="392" style="height:392px;width:259px;">Dauer pheromone plates for dose-response assay</td>
			<td style="width:152px;">Various</td>
			<td></td>
			<td style="width:251px;">Mix 0.085 g Noble agar, 0.5 mL of 0.513 M NaCl and 5 &#181;L of 5 mg/ml cholesterol dissolved in ethanol in each of 6x 15 mL culture tubes. Add sufficient crude pheromone to each tube to create a range of concentrations. Add H2O to bring the volume to 4.725 mL.&#160; Boil over a Bunsen burner until the agar is dissolved and then cool to 55 &#176;C. Add 125 &#181;L 1 M KPO4 buffer (pH 6.0), 50 &#181;l 0.1 M CaCl2, 50 &#181;L 0.1 M MgSO4 and 50 &#181;l of 5 mg/ml streptomycin sulfate to each tube.&#160; Pour the contents of each tube into a 35 mm diameter Petri dish. Let the agar solidify at room temperature overnight (if longer, store in enclosed container to prevent excessive drying).</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Microsphere beads 0.1 micron</td>
			<td style="width:152px;">Polysciences Inc.</td>
			<td>00876-15</td>
			<td style="width:251px;"></td>
		</tr>
		<tr height="68">
			<td height="68" style="height:68px;">M9 buffer</td>
			<td style="width:152px;">Various</td>
			<td></td>
			<td style="width:251px;">Mix 3 g KH2PO4, 6 g Na2HPO4, 5 g NaCl, 1 ml 1 M MgSO4, H2O to 1 liter. Sterilize by autoclaving</td>
		</tr>
	</tbody>
</table>

in vivo imaging of dauer-specific neuronal remodeling in c. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

The production of crude dauer pheromone results in a yellow liquid (Figure 1)&#160;that is both heat and cold stable. Aliquots of crude pheromone are stored at -20 &#176;C indefinitely with no obvious loss in activity. Following the production of pheromone, a single dose-response bioassay is sufficient for a rough estimate of potency. However, for most experiments it is necessary to repeat the bioassay 2 - 3 times and take an average from each experiment. Representative results from two pheromone bioassays are given in Figure 3. From these data an EC50 and EC90 can be calculated.
In addition to resistance to sodium dodecyl sulfate2, dauers can be recognized by several morphological features including an occluded stoma, lateral alae, refractile bodies in the anterior hypodermis and a shrunken pharynx (Figure 2 and Figure&#160;4A&#160;for L2 larvae). Dauers also have several distinctive behavioral characteristics including: a propensity to remain behaviorally quiescent, the presence of an occasional dispersal strategy called nictation, suppression of head foraging movements, suppression of pharyngeal pumping and the induction of excretory duct pulsing2, 5, 23. The behavioral characteristics show variability among individuals.
The molt into dauer takes approximately 12 hr from the end of the pre-dauer L2 stage (L2d)4. This period corresponds with stereotypical growth of the IL2Q arbors. The first noticeable event of the molt into dauer is the suppression of pharyngeal pumping. Suppression of pumping is common to all molts in C. elegans. However, suppression of pharyngeal pumping continues throughout the dauer stage and ends approximately 4 hr following a return of the animal to favorable environmental conditions. During the initial period of pharyngeal pumping suppression, a layer of cuticle forms over the stoma. These changes occur during the first hour following the onset of the molt into dauer. During this initial period there is frequently the formation of an additional neurite process extending from the IL2Q cell bodies. However, this initial additional process usually retracts within 2 hr following the onset of the molt into dauer12.
During hours 2 - 5 following the onset of the molt into dauer the pharynx begins to remodel showing a gradual reduction in size (Figure 4B). Concomitantly, short puncta form and retract along the IL2Q primary dendrite (Figure 5). During hours 4 - 8 following the onset of the molt, the body undergoes a gradual radial shrinkage resulting in the pulling away of the nematode from the L2d cuticle (Figure 4C). This period is correlated with outgrowth of 2&#176; dendrites from the primary dendrites and the establishment of additional dendrites from the IL2Q cell bodies (dauer-specific primary dendrites&#8212;1d&#176;). The dendritic growth is not constant, but rather characterized by both the dynamic formation and retraction of processes as illustrated in Figure 5. The 2&#176; and 1d&#176; dendrites generally extend to the ventral (for IL2Vs) or dorsal (for IL2Ds) midlines. Once the dauer cuticle is completely separated from the L2d cuticle, the application of fluorescent light causes the animal to rapidly spin on its longitudinal axis, making subsequent imaging challenging. Attempts to remove the developing animal from the slide, mechanically remove the L2D cuticle, and mount the animal on a new slide while still maintaining dauer development were not successful.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51834/51834fig1highres.jpg" width="500" /> 
Figure 1.&#160;Crude dauer pheromone is a yellow liquid that can be stored indefinitely at -20 &#176;C. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51834/51834fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51834/51834fig2highres.jpg" width="500" /> 
Figure 2.&#160;DIC micrographs of C. elegans dauers with typical dauer morphological features.&#160;(A) Dorsal mid-body view of a dauer demonstrating an occluded stoma (arrow) and a shrunken pharynx with rectangular terminal bulb (outlined in red, compare with L2 larva shown in Figure 4A). Newly formed dauers will frequently retain part of the L2d cuticle (arrowhead). (B) Dorsal view of hypodermis showing highly refractile bodies (arrow) typical of dauers. Again the L2d cuticle is evident (arrowhead). (C) Lateral cuticle view showing lateral alae, a set of four prominent longitudinal ridges that run from the anus to anterior of the nerve ring. Scale bars, 10 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51834/51834fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51834/51834fig3highres.jpg" width="500" /> 
Figure 3.&#160;Dose response assay to crude dauer pheromone. L1 animals exposed to increasing levels of dauer pheromone incorporated into modified NGM media are more likely to form dauers. Data were collected and pooled from two separate experiments except for the following concentrations, which were only tested once: 0.05%, 2%, and 4%. Data were fit to a sigmoidal dose response curve. The R2 and EC values were calculated using logistic regression. The number of animals examined for each concentration is as follows: 0% n=299, 0.05% n=81, 0.1% n=352, 0.5% n=368, 1% n=241, 2% n=125, 4% n=155. Error bars represent 95% confidence intervals.
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51834/51834fig4highres.jpg" width="500" /> 
Figure 4.&#160;DIC lateral right view micrographs of C. elegans during different stages of the molt into dauer.&#160;(A) L2 animal with typical rounded terminal pharyngeal bulb (outlined in red). (B) Approximately 3 hr following the molt into dauer, the terminal bulb is shrinking and the L2d cuticle is beginning to detach from the newly formed dauer cuticle (arrowhead). (C) Approximately 10 hr following the onset of the dauer molt, the animal has undergone extensive radial shrinkage and detachment from the L2d cuticle (arrowhead). Occasionally, the cuticle lined excretory duct of the L2d stage (arrow) can be seen still attached to the developing dauer. Scale bars, 10 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51834/51834fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51834/51834fig5highres.jpg" width="500" /> 
Figure 5.&#160;Lateral view epifluorescent micrographs of a single animal expressing the IL2 reporter Pklp-6::tdTomato. Top, full-length image of IL2 neurons approximately 3 hr following the onset of the molt into dauer. Inset shows portion of IL2Q dendrite at various time-points following the onset of the molt into dauer. A rapid extension of the IL2Q branches occurs starting approximately 5 hr following the onset of the molt into dauer. Scale bars, 10 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51834/51834fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans at JoVE.com

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

in-vivo-imaging-of-dauer-specific-neuronal-remodeling-in-c-elegans

Nanyang Technological University

Research

JoVE Journal

Neuroscience

11.2K Views.  University of Illinois Urbana-Champaign. Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant ‘dauer’ juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer. 

Video: In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Super-resolution Imaging of Neuronal Dense-core Vesicles

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and medical applications. Strengths of fluorescence include its sensitivity, specificity, and compatibility with live imaging. Unfortunately, conventional forms of fluorescence microscopy suffer from one major weakness, diffraction-limited resolution in the imaging plane, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, this limitation has been overcome with the introduction of super-resolution fluorescence microscopy techniques, such as photoactivated localization microscopy (PALM). Unlike its conventional counterparts, PALM can produce images with a lateral resolution of tens of nanometers. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of cellular structure and organization.
Unfortunately, PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. In this article, we show how to implement single-color PALM studies of vesicular structures in fixed, cultured neurons. PALM is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm. Key steps in our approach include labeling neurons with photoconvertible (green to red) chimeras of vesicle cargo, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a high-resolution PALM image. We also demonstrate the efficacy of our approach by presenting exceptionally well-resolved images of dense-core vesicles (DCVs) in cultured hippocampal neurons, which refute the hypothesis that extrasynaptic trafficking of DCVs is mediated largely by DCV clusters.

We describe how to implement photoactivated localization microscopy (PALM)-based studies of vesicles in fixed, cultured neurons. Key components of our protocol include labeling vesicles with photoconvertible chimeras, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a super-resolution image.

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its completely mapped nervous system and transparent anatomy, presents an ideal model for understanding real-time neural dynamics using calcium indicators. In combination with microfluidic technologies and experimental designs, calcium-imaging studies using these indicators are performed in both free-moving and trapped animals. However, most previous studies utilizing trapping devices, such as the olfactory chip described in Chronis et al., have devices designed for use in the more common hermaphrodite, as the less common male is both morphologically and structurally dissimilar. An adapted olfactory chip was designed and fabricated for increased efficiency in male neuronal imaging with using young adult animals. A turn was incorporated into the worm loading port to rotate the animals and to allow for the separation of the individual neurons within a bilateral pair in 2D imaging. Worms are exposed to a controlled flow of odorant within the microfluidic device, as described in previous hermaphrodite studies. Calcium transients are then analyzed using the open-source software ImageJ. The procedure described herein should allow for an increased amount of male-based C. elegans calcium imaging studies, deepening our understanding of the mechanisms of sex-specific neuronal signaling.

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of an adapted &#34;olfactory chip&#34; for the efficient calcium imaging of C. elegans males is described here. Studies of male exposure to glycerol and a pheromone are also shown.

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its ...

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

Toxicity of misfolded proteins and mitochondrial dysfunction are pivotal factors that promote age-associated functional neuronal decline and neurodegenerative disease across species. Although these neurotoxic challenges have long been considered to be cell-intrinsic, considerable evidence now supports that misfolded human disease proteins originating in one neuron can appear in neighboring cells, a phenomenon proposed to promote pathology spread in human neurodegenerative disease.
C. elegans adult neurons that express aggregating proteins can extrude large (~4 &#181;m) membrane-surrounded vesicles that can include the aggregated protein, mitochondria, and lysosomes. These large vesicles are called &#8220;exophers&#8221; and are distinct from exosomes (which are about 100x smaller and have different biogenesis). Throwing out cellular debris in exophers may occur by a conserved mechanism that constitutes a fundamental, but formerly unrecognized, branch of neuronal proteostasis and mitochondrial quality control, relevant to processes by which aggregates spread in human neurodegenerative diseases.
While exophers have been mostly studied in animals that express high copy transgenic mCherry within touch neurons, these protocols are equally useful in the study of exophergenesis using fluorescently tagged organelles or other proteins of interest in various classes of neurons.
Described here are the physical features of C. elegans exophers, strategies for their detection, identification criteria, optimal timing for quantitation, and animal growth protocols that control for stresses that can modulate exopher production levels. Together, details of protocols outlined here should serve to establish a standard for quantitative analysis of exophers across laboratories. This document seeks to serve as a resource in the field for laboratories seeking to elaborate molecular mechanisms by which exophers are produced and by which exophers are reacted to by neighboring and distant cells.

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

This protocol describes approaches for detection and quantitation of large aggregate and/or organelle extrusions (~4 &#181;m) produced by C. elegans cells in the form of membrane-bound exophers. We describe strains, growth conditions, scoring criteria, timing, and microscopy considerations needed to facilitate dissection of this debris expulsion mechanism.