Name: in vivo imaging of dauer-specific neuronal remodeling in c. elegans
Uploaded: 2014-09-04T15:00:00.000+00:00
Duration: 9 min 39 s
Description: Watch this Scientific Journal Video about In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans at JoVE.com

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 d...

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><tbody><tr><td>N2 &lt;em&gt;C. elegans&lt;/em&gt; Bristol strain</td><td>Caenorhabditis Genetics Center</td><td></td><td></td></tr><tr><td>PT2660 myIs13[P&lt;sub&gt;klp-6&lt;/sub&gt;::gfp] III</td><td>Upon request from Schroeder lab</td><td></td><td></td></tr><tr><td>PT2762 myIs14[P&lt;sub&gt;klp-6&lt;/sub&gt;::gfp] V</td><td>Upon request from Schroeder lab</td><td></td><td></td></tr><tr><td>JK2868 qIs56[P&lt;sub&gt;lag-2&lt;/sub&gt;::gfp] V</td><td>Caenorhabditis Genetics Center</td><td></td><td></td></tr><tr><td>OP50 E. coli</td><td>Caenorhabditis Genetics Center</td><td></td><td></td></tr><tr><td>Petri dishes</td><td>Tritech Research</td><td>60 mm T3308, 35 mm T3501</td><td></td></tr><tr><td>NGM agar</td><td>Various</td><td></td><td>3 g NaCl, 17 g agar, 2.5 g peptone in 975 ml H&lt;sub&gt;2&lt;/sub&gt;O autoclaved cooled to 55&amp;deg;C followed by 1 ml of 5 mg/ml cholesterol in ethanol, 25 ml of 1 M KPO&lt;sub&gt;4&lt;/sub&gt; buffer (pH 6.0), 1 ml of 1 M CaCl&lt;sub&gt;2&lt;/sub&gt;, 1 ml of 1 M MgSO&lt;sub&gt;4&lt;/sub&gt;</td></tr><tr><td>S Media</td><td>Various</td><td></td><td>S media consists of S basal solution (1.46 g NaCl, 0.25g K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 1.5 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 0.25 ml of 5 mg/ml cholesterol in ethanol and H&lt;sub&gt;2&lt;/sub&gt;O 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&lt;sub&gt;, &lt;/sub&gt;H&lt;sub&gt;2&lt;/sub&gt;O to 100 ml, autoclaved), 2.5 ml of Trace Metals Solution (1.86 g disodium EDTA, 0.69 g FeSO&lt;sub&gt;4&lt;/sub&gt;&amp;nbsp;&amp;bull;7 H&lt;sub&gt;2&lt;/sub&gt;O, 0.2 g MnCl2&amp;bull;4 H&lt;sub&gt;2&lt;/sub&gt;O, 0.29 g ZnSO&lt;sub&gt;4&lt;/sub&gt;&amp;nbsp;&amp;bull;7 H&lt;sub&gt;2&lt;/sub&gt;O, 0.025 g CuSO&lt;sub&gt;4&lt;/sub&gt;&amp;nbsp;&amp;bull;5 H&lt;sub&gt;2&lt;/sub&gt;O, H&lt;sub&gt;2&lt;/sub&gt;O to 1 L. Autoclaved and stored in the dark), 0.75 ml of 1 M CaCl&lt;sub&gt;2&lt;/sub&gt;, and 0.75 ml of 1 M MgSO&lt;sub&gt;4&lt;/sub&gt;.</td></tr><tr><td>Dauer pheromone plates for dose-response assay</td><td>Various</td><td></td><td>Mix 0.085 g Noble agar, 0.5 ml of 0.513 M NaCl and 5 &amp;micro;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 H&lt;sub&gt;2&lt;/sub&gt;O to bring the volume to 4.725 ml.&amp;nbsp; Boil over a Bunsen burner until the agar is dissolved and then cool to 55 &amp;deg;C. Add 125 &amp;micro;l 1 M KPO&lt;sub&gt;4&lt;/sub&gt; buffer (pH 6.0), 50 &amp;micro;l 0.1 M CaCl&lt;sub&gt;2&lt;/sub&gt;, 50 &amp;micro;l 0.1 M MgSO&lt;sub&gt;4&lt;/sub&gt; and 50 &amp;micro;l of 5 mg/ml streptomycin sulfate to each tube.&amp;nbsp; 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><td>Microsphere beads 0.1 micron</td><td>Polysciences Inc.</td><td>00876-15</td><td></td></tr><tr><td>M9 buffer</td><td>Various</td><td></td><td>Mix 3 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 6 g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 5 g NaCl, 1 ml 1 M MgSO&lt;sub&gt;4&lt;/sub&gt;, H&lt;sub&gt;2&lt;/sub&gt;O 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 bioassa...

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

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

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Bioengineering

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for the targeted expression of genetically encoded calcium sensors. This protocol outlines the use of these sensors for the in vivo imaging of calcium dynamics in targeted cells, specifically the body wall muscles of the worms. By utilizing the co-expression of presynaptic channelrhodopsin, stimulation of acetylcholine release from excitatory motor neurons can be induced using blue light pulses, resulting in muscle depolarization and reproducible changes in cytoplasmic calcium levels. Two worm immobilization techniques are discussed with varying levels of difficulty. Comparison of these techniques demonstrates that both approaches preserve the physiology of the neuromuscular junction and allow for the reproducible quantification of calcium transients. By pairing optogenetics and functional calcium imaging, changes in postsynaptic calcium handling and homeostasis can be evaluated in a variety of mutant backgrounds. Data presented validates both immobilization techniques and specifically examines the roles of the C. elegans sarco(endo)plasmic reticular calcium ATPase and the calcium-activated BK potassium channel in the body wall muscle calcium regulation.

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

This method provides a way to couple optogenetics and genetically encoded calcium sensors to image baseline cytosolic calcium levels and changes in evoked calcium transients in the body wall muscles of the model organism C. elegans.

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for ...

Biochemistry

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, ...

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure the activity of neurons during various behaviors. To correlate neural activity with behavior, the animal should not be immobilized but should be able to move. Many behavioral changes occur during long time scales and require recording over many hours of behavior. This also makes it necessary to culture the worms in the presence of food. How can worms be cultured and their neural activity imaged over long time scales? Agarose Microchamber Imaging (AMI) was previously developed to culture and observe small larvae and has now been adapted to study all life stages from early L1 until the adult stage of C. elegans. AMI can be performed on various life stages of C. elegans. Long-term calcium imaging is achieved without immobilizing the animals by using short externally triggered exposures combined with an electron multiplying charge-coupled device (EMCCD) camera recording. Zooming out or scanning can scale up this method to image up to 40 worms in parallel. Thus, a method is described to image behavior and neural activity over long time scales in all life stages of C. elegans.

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Imaging behavior and neural activity over long time scales without immobilization of the animal is a prerequisite to understand behavior. Agarose microfluidic chambers imaging (AMI) can be used to image neural activity and behavior for all life stages of Caenorhabditis elegans.

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure ...

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using fluorescent-labeled reporter proteins or differential interference contrast (DIC) microscopy, allows for the examination of the temporal progression of these dynamic events which is otherwise inferred from analysis of fixed samples1,2. Moreover, the study of the developmental regulations of cellular processes necessitates conducting time-lapse experiments on an intact organism during development. The Caenorhabiditis elegans embryo is light-transparent and has a rapid, invariant developmental program with a known cell lineage3, thus providing an ideal experiment model for studying questions in cell biology4,5and development6-9. C. elegans is amendable to genetic manipulation by forward genetics (based on random mutagenesis10,11) and reverse genetics to target specific genes (based on RNAi-mediated interference and targeted mutagenesis12-15). In addition, transgenic animals can be readily created to express fluorescently tagged proteins or reporters16,17. These traits combine to make it easy to identify the genetic pathways regulating fundamental cellular and developmental processes in vivo18-21. In this protocol we present methods for live imaging of C. elegans embryos using DIC optics or GFP fluorescence on a compound epifluorescent microscope. We demonstrate the ease with which readily available microscopes, typically used for fixed sample imaging, can also be applied for time-lapse analysis using open-source software to automate the imaging process.

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

The C. elegans embryo is a powerful system for studying cell biology and development. We present a protocol for live imaging of C. elegans embryos utilizing DIC optics or fluorescence using readily available epifluorescent microscopes and open-source software.

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using ...

Biology

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding these biological processes often requires long-term and repeated imaging of the same animal. Long recovery times associated with conventional immobilization methods done on agar pads have detrimental effects on animal health making it inappropriate to repeatedly image the same animal over long periods of time. This paper describes a microfluidic chip design, fabrication method, on-chip C. elegans culturing protocol, and three examples of long-term imaging to study developmental processes in individual animals. The chip, fabricated with polydimethylsiloxane and bonded on a cover glass, immobilizes animals on a glass substrate using an elastomeric membrane that is deflected using nitrogen gas. Complete immobilization of C. elegans enables robust time-lapse imaging of cellular and sub-cellular events in an anesthetic-free manner. A channel geometry with a large cross-section allows the animal to move freely within two partially sealed isolation membranes permitting growth in the channel with a continuous food supply. Using this simple chip, imaging of developmental phenomena such as neuronal process growth, vulval development, and dendritic arborization in the PVD sensory neurons, as the animal grows inside the channel, can be performed. The long-term growth and imaging chip operates with a single pressure line, no external valves, inexpensive fluidic consumables, and utilizes standard worm handling protocols that can easily be adapted by other laboratories using C. elegans.

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

The protocol describes a simple microfluidic chip design and microfabrication methodology used to grow C. elegans in presence of a continuous food supply for up to 36 h. The growth and imaging device also enables intermittent long-term high-resolution imaging of cellular and sub-cellular processes during development for several days.

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding ...

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the neurotransmitter glutamate, and the excessive stimulation of its postsynaptic receptors. However, information on the subsequent molecular events that culminate in the distinct neuronal swelling morphology of this type of neurodegeneration is lacking. Other aspects, such as changes in specific subcellular compartments, or the basis for the differential cellular vulnerability of distinct neuronal subtypes, remain under-explored. Furthermore, a range of factors that come into play in studies that use in vitro or ex vivo preparations might modify and distort the natural progression of this form of neurodegeneration. It is therefore important to study excitotoxic necrosis in live animals by monitoring the effects of interventions that regulate the extent of neuronal necrosis in the genetically amenable and transparent&#160;model system of the nematode Caenorhabditis elegans. This protocol describes methods of studying excitotoxic necrosis in C. elegans neurons, combining optical, genetic, and molecular analysis. To induce excitotoxic conditions in C. elegans, a&#160;knockout of a glutamate transporter gene (glt-3) is combined with a neuronal sensitizing genetic background (nuls5 [Pglr-1::G&#945;S(Q227L)]) to produce glutamate receptor hyperstimulation and neurodegeneration. Nomarski differential interference contrast (DIC), fluorescent, and confocal microscopy in live animals are methods used to quantify neurodegeneration, follow subcellular localization of fluorescently labeled proteins, and quantify mitochondrial morphology in the degenerating neurons. Neuronal Fluorescence Activated Cell Sorting (FACS) is used to distinctly sort at-risk neurons for cell-type specific transcriptomic analysis of neurodegeneration. A combination of live imaging and FACS methods as well as the benefits of the C. elegans model organism allow researchers to leverage this system to obtain reproducible data with a large sample size. Insights from these assays could translate to novel targets for therapeutic intervention in neurodegenerative diseases.

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

In a C. elegans excitotoxicity model, this protocol employs in vivo imaging to analyze the regulation of necrotic neurodegeneration, the effect of genes encoding candidate mediators, and involvement of mitochondria. Cell dissociation and sorting is used to specifically obtain at-risk neurons for cell-specific transcriptomic analysis of neurodegeneration and neuroprotection mechanisms.

Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the ...

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

In neurons, microtubule orientation has been a key assessor to identify axons that have plus-end out microtubules and dendrites that generally have mixed orientation. Here we describe methods to label, image, and analyze the microtubule dynamics and growth during the development and regeneration of touch neurons in C. elegans. Using genetically encoded fluorescent reporters of microtubule tips, we imaged the axonal microtubules. The local changes in microtubule behavior that initiates axon regeneration following axotomy can be quantified using this protocol. This assay is adaptable to other neurons and genetic backgrounds to investigate the regulation of microtubule dynamics in various cellular processes.

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

A protocol for imaging the dynamic microtubules in vivo using fluorescently labeled End binding protein has been presented. We described the methods to label, image, and analyze the dynamic microtubules in the Posterior lateral microtubule (PLM) neuron of C. elegans.

In neurons, microtubule orientation has been a key assessor to identify axons that have plus-end out microtubules and dendrites that generally have mixed ...

Quantifying Levels of Dopaminergic Neuron Morphological Alteration and Degeneration in Caenorhabditis elegans

Dopamine neuron loss is involved in the pathology of Parkinson&#39;s Disease (PD), a highly prevalent neurodegenerative disorder affecting over 10 million people worldwide. Since many details about PD etiology remain unknown, studies investigating genetic and environmental contributors to PD are needed to discover methods of prevention, management, and treatment. Proper characterization of dopaminergic neuronal loss may be relevant not only to PD research, but to other increasingly prevalent neurodegenerative disorders.
There are established genetic and chemical models of dopaminergic neurodegeneration in the Caenorhabditis elegans model system, with easy visualization of neurobiology supported by the nematodes&#39; transparency and invariant neuronal architecture. In particular, hermaphroditic C. elegans&#39; dopaminergic neuron morphological changes can be visualized using strains with fluorescent reporters driven by cell-specific promotors such as the dat-1 dopamine transporter gene, which is expressed exclusively in their eight dopaminergic neurons.
With the capabilities of this model system and the appropriate technology, many laboratories have studied dopaminergic neurodegeneration. However, there is little consistency in the way the data is analyzed and much of the present literature uses binary scoring analyses that capture the presence of degeneration but not the full details of the progression of neuron loss. Here, we introduce a universal scoring system to assess morphological changes and degeneration in C. elegans&#39; cephalic neuron dendrites. This seven-point scale allows for analysis across a full range of dendrite morphology, ranging from healthy neurons to complete dendrite loss, and considering morphological details including kinks, branching, blebs, and breaks. With this scoring system, researchers can quantify subtle age-related changes as well as more dramatic chemical-induced changes. Finally, we provide a practice set of images with commentary that can be used to train, calibrate, and assess the scoring consistency of researchers new to this method. This should improve within- and between- laboratory consistency, increasing rigor and reproducibility.

Quantifying Levels of Dopaminergic Neuron Morphological Alteration and Degeneration in Caenorhabditis elegans

In this article, we showcase how to use a seven-point scoring system to consistently quantify changes to dopaminergic neuron dendrite morphology in C. elegans. This system is intended for analyses of dopaminergic neurodegeneration assays utilizing genetic, chemical, and age-based models of neurodegenerative disorders.

Dopamine neuron loss is involved in the pathology of Parkinson&#39;s Disease (PD), a highly prevalent neurodegenerative disorder affecting over 10 million ...

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a crucial component of intracellular signaling. The visualization of subcellular Ca2+ dynamics in vivo,&#160;where neurons can be studied in their native, intact circuitry, has proven technically challenging in complex nervous systems. The transparency and relatively simple nervous system of the nematode Caenorhabditis elegans enable the cell-specific expression and in vivo visualization of fluorescent tags and indicators. Among these are fluorescent indicators that have been modified for use in the cytoplasm as well as various subcellular compartments, such as the mitochondria. This protocol enables non-ratiometric Ca2+ imaging in vivo with a subcellular resolution that permits the analysis of Ca2+ dynamics down to the level of individual dendritic spines and mitochondria. Here, two available genetically encoded indicators with different Ca2+ affinities are used to demonstrate the use of this protocol for measuring relative Ca2+ levels within the cytoplasm or mitochondrial matrix in a single pair of excitatory interneurons (AVA). Together with the genetic manipulations and longitudinal observations possible in C. elegans, this imaging protocol may be useful for answering questions regarding how Ca2+ handling regulates neuronal function and plasticity.

Subcellular Imaging of Neuronal Calcium Handling In Vivo

The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a ...

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

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Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

In vivo Neuronal Calcium Imaging in C. elegans

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

Quantifying Levels of Dopaminergic Neuron Morphological Alteration and Degeneration in Caenorhabditis elegans

Subcellular Imaging of Neuronal Calcium Handling In Vivo