The work described in this manuscript was done for an intermediate-level class in neuroscience. Funds for the reagents and supplies were provided by the Biology Department at Vassar College. The microscopes and digital imaging system were also provided by the Biology Department at Vassar College. The author thanks all the many students who took this course.

All use of invertebrate animals was in compliance with the animal care and use guidelines of the institution.
1. Preparation of pesticide-coated Petri plates
<ol>
	<li>Prepare agar Petri dishes (6 cm diameter work best) using standard procedures6. 
	NOTE: These can be made months in advance and stored in the refrigerator until use. Bring plates to room temperature (RT) on the benchtop. In most cases, the student groups should be provided with a plate for each dilution of the pesticide mixture, including a water control.</li>
	<li>Prepare 1 mL of each dilution of the chosen pesticide in labeled 1.5 mL microfuge tubes. For safe handling, be sure to read the instructions and provide gloves, or work in a fume hood, if suggested. 
	NOTE: Suggested dilutions to test: 1 mL of full-strength pesticide, 1:10 dilution in water (100 &#181;L of pesticide, 900 &#181;L of distilled water), 1:100 dilution (10 &#181;L of pesticide, 990 &#181;L of water), 1:1000 dilution (10 &#181;L of the 1:10 dilution, 990 &#181;L of water), etc.</li>
	<li>Make a solution spreader by carefully bending a glass Pasteur pipette (9 ml size) in a Bunsen burner flame. Use the flame to close the opening of the pipette. Then ethanol-sterilize the spreader before each spreading on the Petri plate.</li>
	<li>Using a micropipetter, place 100 &#181;L of the dilution (or water control) on the center of the agar and spread gently across the surface with the sterilized spreader. Re-sterilize the spreader prior to each use.</li>
	<li>Set aside the Petri dishes covered, and let the solution soak into the surface until &#34;dry.&#34; 
	&#8203;NOTE: Depending on the pesticide and humidity levels in the laboratory, this may take some time. Students might need to prepare their plates the day before the exposure period.</li>
	<li>For exposure periods longer than 12 h, add 50 &#181;L of an overnight culture of Escherichia coli to the plates prior to adding the nematodes. For acute experiments (12 h or less), there is no need to have E. coli on the plates.</li>
</ol>
2. Adding nematodes to pesticide-coated plates
NOTE: Students will need to have a plate of adult nematodes (5 day cultures are best) of the fluorescent strain LX929, which is available from the Caenorhabditis elegans Genetics Center. There are two possible procedures for this. If students have had prior experience working with nematodes (for example, if this procedure is part of a multi-week exercise and they have already learned how to pick nematodes with a worm pick), use Step 2.1. If they have not, use Step 2.2.
<ol>
	<li>Pick nematodes with a worm pick. Fashion a worm pick from a 1 in piece of platinum wire that is melted to a Pasteur pipette using a Bunsen burner flame. Pick up a small blob of sticky E. coli onto the pick and then pick up adult nematodes using the sticky blob. For a decent sample size, pick 10 nematodes for each treatment plate. 
	NOTE: Students without much experience can follow Steps 2.2-2.3.</li>
	<li>Use a micropipette to place 1 mL of sterile water onto the plate of nematodes. Swish the water around to lift up the nematodes, then remove the liquid with the nematodes to a 1.5 mL microfuge tube.</li>
	<li>Let the nematodes settle by gravity for about 10 min, then carefully remove at least 500 &#181;L of the water and discard. Resuspend the nematodes and, using a cut off yellow pipette tip, remove 50-100 &#181;L of suspended worms to each treatment plate. Be sure that each plate has at least 10 adult worms.</li>
	<li>Set a timer, or note the time, and expose the nematodes at RT for the chosen time period. During the chosen exposure time, prepare microscope slides for wet mounts. The slides should be made the same day as the microscopy. 
	NOTE: Student groups will have chosen their time of exposure during the planning for their independent study.</li>
</ol>
3. Preparing microscope slides for wet mounts
NOTE: Steps 3.1-3.3 can be performed by each student group. Prepare three slides each.
<ol>
	<li>Prepare two slides by placing a piece of label tape on only one side of the slide (Figure 1, left panel). The tape is used to create the correct thickness for the agarose to form a pad of uniform thickness. Then position a clean, unused slide in between them on the benchtop, as illustrated in the figure.</li>
	<li>Use a micropipetter to place a 10 &#181;L drop of melted agarose (3% in ddH2O, heated using a 65 &#176;C dry block heater) onto the center of the slide that is in the middle.</li>
	<li>Flatten the drop by placing another clean slide on top, perpendicular to the slide with the drop, as shown in the figure (Figure 1, right panel). Flatten by applying pressure, so the agarose drop forms the thickness of the labeling tape.</li>
	<li>The agarose will solidify within about 1 min. Then, apply steady pressure to separate the two slides. The agar circle will adhere to one of the slides. Rest this slide, agar side up, on the benchtop, and allow it to dry for 1-2 min before using.</li>
</ol>
4. Mounting live nematodes to microscope slides
<ol>
	<li>To add nematodes to the prepared slide with the agarose pad, add a 5 &#181;L drop of water containing 1 M sodium azide (NaN3) to the agar pad. This solution anesthetizes the worms and immobilizes them. 
	CAUTION: Sodium azide stock solution can cause illness if ingested.</li>
	<li>Then transfer nematodes (at least 10 per treatment slide) to the droplet using a worm pick. Sterilize the pick before and after using it to transfer nematodes.</li>
	<li>Use forceps to add a coverslip by placing it at an angle and slowly lowering it. Prevent the coverslip from slipping or lowering too quickly by using a pencil or metal probe.</li>
	<li>Place the prepared wet mount on a fluorescent microscope to observe the worms, first under 10x magnification and phase contrast, then under 40x magnification with phase contrast and fluorescent illumination.</li>
</ol>
5. Fluorescent microscopy
NOTE: Student groups should make a wet mount of several worms of each type on separate microscope slides, which they should label appropriately. The instructions that follow are general tips about the use of a standard fluorescence compound microscope that can have an attached trinocular setup, digital camera, and computer system. The setup used for this manuscript is seen in Figure 2. Students should be familiar with the settings of the microscope, including the objectives, types of light filters, ability to switch between bright field and fluorescence light, how to send the light signal to the digital camera, etc.
<ol>
	<li>Place one prepared wet mount at a time on the microscope stage and secure it in place using the microscope stage clips.</li>
	<li>Begin viewing the wet mounts by first viewing with the lowest magnification objective, which is typically 10x, and use bright field or phase contrast to visualize and focus on the nematodes.</li>
	<li>Once a nematode is located in the field of view, focus on it using the fine focus knob on the microscope. Switch the illumination to the fluorescence by turning the dial. Then direct the light to the camera to capture the digital images. Adjust the illumination using the imaging software so that the neurons are brightly illuminated but not oversaturated.</li>
	<li>At some point, obtain a separate image of a ruler at the same magnification as the cell images to provide a magnification scale for their figure. Place a short section of a transparent ruler, or a micrometer slide, onto the microscope stage, focus on it using the fine focus knob, and obtain an image using the imaging software.</li>
	<li>Obtain additional images (from at least 10 separate nematodes) at higher magnifications, if possible. The intense light will likely bleach the area from which it is being imaged, so carefully monitor the imaging and do not allow the light to remain on the same area of the slide for long periods of time. Be sure to name the acquired images to keep track of the nematode and region, as well as the treatment and magnification.</li>
	<li>Make a folder on the computer and move the images into the folder for use in analysis and figure preparation.</li>
</ol>
6. Morphological integrity assessment scale
NOTE: A key cytological characteristic of neurodegenerative damage in neurons is a change to the soma morphology. There are three morphologies that can easily be viewed and counted.
<ol>
	<li>Count the neurons with smooth, uniformly fluorescent processes and non-round cell bodies as normal, as shown in Figure 3.</li>
	<li>Often the soma becomes rounded and swollen-looking compared with healthy, young neurons. Count the neurons that look like this as &#34;rounded.&#34;</li>
	<li>Neurons with long neural processes can exhibit blebbing, where there are rounded puncta or even gaps in the processes. Count the neurons that show these features as &#34;blebbed.&#34;</li>
</ol>
7. Data analysis and figure preparation
<ol>
	<li>If the percentage healthy (no rounding, no blebbing) and the percentage impaired (rounding and/or blebbing) for at least ten nematodes for each treatment were counted (Figure 3), perform statistical analysis using available statistical software.</li>
	<li>Count all of the neurons that are fluorescently tagged, keeping a tally of those exhibiting normal morphology, blebbing morphology, or rounding morphology, and then calculate the percentage of each morphology out of the total count for each nematode analyzed.</li>
	<li>To prepare figures, import the images into an appropriate software like Powerpoint and then add labels and a scale bar based on the ruler or micrometer image acquired.</li>
</ol>

<ol>
	<li>Duzguner, V., Erdogan, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+exposure+to+imidacloprid+induces+inflammation+and+oxidative+stress+in+the+liver+&amp;+central+nervous+system+of+rats.">Chronic exposure to imidacloprid induces inflammation and oxidative stress in the liver &amp; central nervous system of rats.</a> Pesticide Biochemistry and Physiology. 104 (1), 58-64 (2012).</li><li>Catae, A. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exposure+to+a+sublethal+concentration+of+imidacloprid+and+the+side+effects+on+target+and+nontarget+organs+of+Apis+mellifera+(Hymenoptera,+Apidae).">Exposure to a sublethal concentration of imidacloprid and the side effects on target and nontarget organs of Apis mellifera (Hymenoptera, Apidae).</a> Ecotoxicology. 27 (2), 109-121 (2018).</li><li>Bradford, B. R., Whidden, E., Gervasio, E. D., Checchi, P. M., Raley-Susman, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neonicotinoid-containing+insecticide+disruption+of+growth,+locomotion,+and+fertility+in+Caenorhabditis+elegans.">Neonicotinoid-containing insecticide disruption of growth, locomotion, and fertility in Caenorhabditis elegans.</a> PLOS One. 15 (9), 0238637 (2020).</li><li>Jospin, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+neuronal+acetylcholine+receptor+regulates+the+balance+of+muscle+excitation+and+inhibition+in+Caenorhabditis+elegans.">A neuronal acetylcholine receptor regulates the balance of muscle excitation and inhibition in Caenorhabditis elegans.</a> PLoS Biology. 7 (12), 1000265 (2009).</li><li>Susman, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Discovery-Based Learning in the Life Sciences. , (2015).</li><li>Stiernagle, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maintenance+of+C.+elegans.">Maintenance of C. elegans.</a> WormBook. , (2006).</li><li>Brody, H. A., Chou, E., Gray, J. M., Pokyrwka, N. J., Raley-Susman, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mancozeb-induced+behavioral+deficits+precede+structural+neural+degeneration.">Mancozeb-induced behavioral deficits precede structural neural degeneration.</a> NeuroToxicology. 34, 74-81 (2013).</li><li>Chen, P., Martinez-Finley, E. J., Bornhorst, J., Chakraborty, S., Aschner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metal-induced+neurodegeneration+in+C.+elegans.">Metal-induced neurodegeneration in C. elegans.</a> Frontiers in Aging Neuroscience. 5, (2013).</li><li>Hart, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavior.">Behavior.</a> WormBook. , (2006).</li><li>Harlow, P. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+nematode+Caenorhabditis+elegans+as+a+tool+to+predict+chemical+activity+on+mammalian+development+and+identify+mechanisms+influencing+toxicological+outcome.">The nematode Caenorhabditis elegans as a tool to predict chemical activity on mammalian development and identify mechanisms influencing toxicological outcome.</a> Scientific Reports. 6 (1), 22965 (2016).</li></ol>

There are no conflicts of interest to disclose.

The protocols described in this manuscript work successfully alone or as part of a multi-week independent student group project. The protocols are also amenable to stand-alone, one-week exploratory experiences. The nematode strains are cheap and easy to maintain in the research laboratory. Students can readily learn how to pick worms with a worm pick or how to move them by flushing plates with water and allowing them to settle by gravity. The experiments can be performed over the course of a single lab period or over mul...

Caenorhabditis elegans is an excellent model organism for laboratory course training in biological science courses for introductory- and intermediate-level students. This laboratory procedure can be used as part of a multi-week module that explores various effects of commonly used pesticides on C. elegans behavior and cell biology. Students can learn how to design and carry out independent projects that teach them data analysis and presentation skills. This paper focuses on the protocols for exposing C. elegans to pesticide mixtures and then observing and analyzing the effects on neuron morphology.
Lawn chemical pesticide mixtures are widely used for residential and agricultural use and can be purchased at any local garden store. There is increasing concern about the safety of these chemicals for humans and wildlife1,2,3. Students can read the scientific literature and select a pesticide for experimental evaluation and, in so doing, can learn about basic biology and neurobiology, as well as important laboratory skills like experimental design and analysis, and general lab skills like pipetting and serial dilutions, dissecting microscopy, fluorescent microscopy, digital photography, and figure production.
The protocols described in this paper can stand alone in an intermediate-level course in biology or neuroscience or be part of a multi-week module that could also include measurements of behaviors governed by particular groups of neurons. For example, described in this protocol is an assessment of the morphology of cholinergic neurons that govern locomotion using a strain of nematode that expresses GFP (LX 929) in cholinergic neurons4. These strains can be obtained for very low prices from the Caenorhabditis elegans Genetics Center (https://cgc.umn.edu/). Strains expressing GFP in dopaminergic neurons (OH 7457), cholinergic neurons (LX 929), or mCherry expressed in all neurons (PVX4) are all good choices. Students could also measure locomotion and obtain data to accompany the assessment of morphology. A full description of a multi-week student group project can be found in Susman5.
This student group project is quite inexpensive and easy to set up for groups of four students. The materials needed include a dissecting microscope, access to a fluorescence compound microscope that can have an attached digital camera, Petri plates and access to nematode growth agar, growth-limited bacteria (strain OP50, from the CGC), a gas flame Bunsen burner or an alcohol lamp, an autoclave, platinum wire, and general laboratory supplies like micropipetters, microscope slides, coverslips, and glass Pasteur pipettes. Depending on the chemical toxicant being examined by the student groups, the steps in the protocol might need to occur under a fume hood or with gloves. This protocol uses chemical mixtures that are water-soluble (not volatile), and all safe-handling procedures recommended by the manufacturer are followed.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agar</td>
			<td>Fisher Scientific</td>
			<td>BP97445</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Fisher Scientific</td>
			<td>MP1AGAH0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol lamp</td>
			<td>Fisher Scientific</td>
			<td>&#160;17012826</td>
			<td></td>
		</tr>
		<tr>
			<td>Bunsen burner</td>
			<td>Fisher Scientific</td>
			<td>17-012-820</td>
			<td></td>
		</tr>
		<tr>
			<td>C. elegans strains</td>
			<td>C. elegans Genetics Center</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl</td>
			<td>Fisher Scientific</td>
			<td>10035-04-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Cholesterol</td>
			<td>Fisher Scientific</td>
			<td>AAA1147030</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Fisher Scientific</td>
			<td>12-545-AP</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital camera</td>
			<td>Nikon</td>
			<td></td>
			<td>These can vary depending on the requirement</td>
		</tr>
		<tr>
			<td>Dissecting scope</td>
			<td>Nikon</td>
			<td>SMZ745</td>
			<td></td>
		</tr>
		<tr>
			<td>E. coli strain (OP50)</td>
			<td>C. elegans Genetics Center</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Fisher Scientific</td>
			<td>BP2818100</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent scope</td>
			<td>Nikon</td>
			<td></td>
			<td>These can vary depending on the requirement</td>
		</tr>
		<tr>
			<td>Imaging software</td>
			<td>Nikon</td>
			<td></td>
			<td>These can vary depending on the requirement</td>
		</tr>
		<tr>
			<td>Inoculation loop</td>
			<td>Fisher Scientific</td>
			<td>&#160;131045</td>
			<td></td>
		</tr>
		<tr>
			<td>LB Broth Base</td>
			<td>Fisher Scientific</td>
			<td>BP9723-500</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>Fisher Scientific</td>
			<td>10034-99-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfuge tubes</td>
			<td>Fisher Scientific</td>
			<td>&#160;05408129</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>Fisher Scientific</td>
			<td>22-265446</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipets</td>
			<td>Fisher Scientific</td>
			<td>13-678-20A</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Fisher Scientific</td>
			<td>AS4050</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips</td>
			<td>Fisher Scientific</td>
			<td>94060316</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetters</td>
			<td>Fisher Scientific</td>
			<td>14-386-319</td>
			<td></td>
		</tr>
		<tr>
			<td>Platinum wire</td>
			<td>Genesee Scientific</td>
			<td>59-1M30P</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Phosphate buffer</td>
			<td>Fisher Scientific</td>
			<td>AAJ61413AP</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Fisher Scientific</td>
			<td>AC447810250</td>
			<td></td>
		</tr>
	</tbody>
</table>

examining the effect of pesticides on caenorhabditis elegans neurons

Caenorhabditis elegans is a powerful model organism used in many research laboratories to understand the consequences of exposure to chemical pollutants, pesticides, and a wide variety of toxic substances. These nematodes are easy to work with and can be used to generate novel research findings, even in the undergraduate biology laboratory. A multi-week laboratory series of authentic, student-driven research projects trains students in a toolkit of techniques and approaches in behavioral measurements, cell biology, and microscopy that they then apply to their projects. One technique in that toolkit is quantifying the percentage of neurons exhibiting neurodegenerative damage following exposure to a chemical toxicant like a pesticide. Young adult C. elegans nematodes can be exposed to different concentrations of commercially available pesticides or other types of toxicants for 2-24 h. Then, undergraduate students can visualize different neuron subtypes using fluorescent-expressing strains of C. elegans. These techniques do not require sophisticated image processing software and are effective at even low magnifications, making the need for expensive confocal microscopy unnecessary. This paper demonstrates how to treat the nematodes with pesticides and how to image and score the neurons. It also provides a straightforward protocol for the microscopy and analysis of neuron morphology. The materials used for this technique are inexpensive and readily available in most undergraduate biology departments. This technique can be combined with behavioral measures like locomotion, basal slowing, or egg-laying to conduct a potentially publishable series of experiments and give undergraduate students an authentic research experience at a very low cost.

The methods and protocols described in this paper provide important laboratory skills for intermediate-level undergraduate students in biology or neuroscience. Students can gain important experience in developing an independent project and conducting an experiment of their own design that could provide novel results. Figure 3 shows an optimal result of a student project that eventually became part of a published paper3. While the majority of student projects do not re...

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Examining the Effect of Pesticides on Caenorhabditis elegans Neurons

Young adult Caenorhabditis elegans nematodes are exposed to different concentrations of commercial pesticides or other toxicants for 2-24 h. Then, different neurons can be visualized using fluorescent-expressing strains. This paper demonstrates how to expose nematodes to pesticides and assess neuron damage.

Examining the Effect of Pesticides on Caenorhabditis elegans Neurons

Caenorhabditis elegans is a powerful model organism used in many research laboratories to understand the consequences of exposure to chemical pollutants, ...

examining-the-effect-of-pesticides-on-caenorhabditis-elegans-neurons

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2.2K Views.  Vassar College. Young adult Caenorhabditis elegans nematodes are exposed to different concentrations of commercial pesticides or other toxicants for 2-24 h. Then, different neurons can be visualized using fluorescent-expressing strains. This paper demonstrates how to expose nematodes to pesticides and assess neuron damage. 

Video: Examining the Effect of Pesticides on Caenorhabditis elegans Neurons

Cut-loading: A Useful Tool for Examining the Extent of Gap Junction Tracer Coupling Between Retinal Neurons

In addition to chemical synaptic transmission, neurons that are connected by gap junctions can also communicate rapidly via electrical synaptic transmission. Increasing evidence indicates that gap junctions not only permit electrical current flow and synchronous activity between interconnected or coupled cells, but that the strength or effectiveness of electrical communication between coupled cells can be modulated to a great extent1,2. In addition, the large internal diameter (~1.2 nm) of many gap junction channels permits not only electric current flow, but also the diffusion of intracellular signaling molecules and small metabolites between interconnected cells, so that gap junctions may also mediate metabolic and chemical communication. The strength of gap junctional communication between neurons and its modulation by neurotransmitters and other factors can be studied by simultaneously electrically recording from coupled cells and by determining the extent of diffusion of tracer molecules, which are gap junction permeable, but not membrane permeable, following iontophoretic injection into single cells. However, these procedures can be extremely difficult to perform on neurons with small somata in intact neural tissue.
Numerous studies on electrical synapses and the modulation of electrical communication have been conducted in the vertebrate retina, since each of the five retinal neuron types is electrically connected by gap junctions3,4. Increasing evidence has shown that the circadian (24-hour) clock in the retina and changes in light stimulation regulate gap junction coupling3-8. For example, recent work has demonstrated that the retinal circadian clock decreases gap junction coupling between rod and cone photoreceptor cells during the day by increasing dopamine D2 receptor activation, and dramatically increases rod-cone coupling at night by reducing D2 receptor activation7,8. However, not only are these studies extremely difficult to perform on neurons with small somata in intact neural retinal tissue, but it can be difficult to adequately control the illumination conditions during the electrophysiological study of single retinal neurons to avoid light-induced changes in gap junction conductance.
Here, we present a straightforward method of determining the extent of gap junction tracer coupling between retinal neurons under different illumination conditions and at different times of the day and night. This cut-loading technique is a modification of scrape loading9-12, which is based on dye loading and diffusion through open gap junction channels. Scrape loading works well in cultured cells, but not in thick slices such as intact retinas. The cut-loading technique has been used to study photoreceptor coupling in intact fish and mammalian retinas7, 8,13, and can be used to study coupling between other retinal neurons, as described here.

An easy and convenient method to determine the extent of gap junction tracer coupling between retinal neurons is described. This technique enables one to investigate the function of the electrical synapses between neurons in the intact retina under different illumination conditions and at different times of the day and night.

In addition to chemical synaptic transmission, neurons that are connected by gap junctions can also communicate rapidly via electrical synaptic ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

A Caenorhabditis elegans Model System for Amylopathy Study

Amylopathy is a term that describes abnormal synthesis and accumulation of amyloid beta (A&beta;) in tissues with time. A&beta; is a hallmark of Alzheimer's disease (AD) and is found in Lewy body dementia, inclusion body myositis and cerebral amyloid angiopathy 1-4. Amylopathies progressively develop with time. For this reason simple organisms with short lifespans may help to elucidate molecular aspects of these conditions. Here, we describe experimental protocols to study A&beta;-mediated neurodegeneration using the worm Caenorhabditis elegans. Thus, we construct transgenic worms by injecting DNA encoding human A&beta;42 into the syncytial gonads of adult hermaphrodites. Transformant lines are stabilized by a mutagenesis-induced integration. Nematodes are age synchronized by collecting and seeding their eggs. The function of neurons expressing A&beta;42 is tested in opportune behavioral assays (chemotaxis assays). Primary neuronal cultures obtained from embryos are used to complement behavioral data and to test the neuroprotective effects of anti-apoptotic compounds.

A Caenorhabditis elegans Model System for Amylopathy Study

We describe methods to study aspects of amylopathies in the worm C. elegans. We show how to construct worms expressing human A&beta;42 in neurons and how to test their function in behavioral assays. We further show how to obtain primary neuronal cultures that can be used for pharmacological testing.

Amylopathy is a term that describes abnormal synthesis and accumulation of amyloid beta (A&beta;) in tissues with time. A&beta; is a hallmark of ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Methods for Studying the Mechanisms of Action of Antipsychotic Drugs in Caenorhabditis elegans

Caenorhabditis elegans is a simple genetic organism amenable to large-scale forward and reverse genetic screens and chemical genetic screens. The C. elegans genome includes potential antipsychotic drug (APD) targets conserved in humans, including genes encoding proteins required for neurotransmitter synthesis and for synaptic structure and function. APD exposure produces developmental delay and/or lethality in nematodes in a concentration-dependent manner. These phenotypes are caused, in part, by APD-induced inhibition of pharyngeal pumping1,2. Thus, the developmental phenotype has a neuromuscular basis, making it useful for pharmacogenetic studies of neuroleptics. Here we demonstrate detailed procedures for testing APD effects on nematode development and pharyngeal pumping. For the developmental assay, synchronized embryos are placed on nematode growth medium (NGM) plates containing APDs, and the stages of developing animals are then scored daily. For the pharyngeal pumping rate assay, staged young adult animals are tested on NGM plates containing APDs. The number of pharyngeal pumps per unit time is recorded, and the pumping rate is calculated. These assays can be used for studying many other types of small molecules or even large molecules.

Methods for Studying the Mechanisms of Action of Antipsychotic Drugs in Caenorhabditis elegans

Approaches for testing the effects of antipsychotic drugs (APDs) in Caenorhabditis elegans are demonstrated. Assays are described for testing drug effects on development and viability and on pharyngeal pumping rate. These methods are also applicable for pharmacogenetic experiments with drug classes other than APDs.

Caenorhabditis elegans is a simple genetic organism amenable to  large-scale forward and reverse genetic screens and chemical genetic screens. The C.  ...

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the development of more effective treatments is the relative lack of understanding of the molecular underpinnings of the effects of ethanol on behavior. The nematode, Caenorhabditis elegans (C.&#160;elegans), provides a useful model in which to generate and test hypotheses about the molecular effects of ethanol. Here, we describe an assay that has been developed and used to examine the roles of particular genes and environmental factors in behavioral responses to ethanol, in which locomotion is the behavioral output. Ethanol dose-dependently causes an acute depression of crawling on an agar surface. The effects are dynamic; animals exposed to a high concentration demonstrate an initial strong depression of crawling, referred to here as initial sensitivity, and then partially recover locomotion speed despite the continued presence of the drug. This ethanol-induced behavioral plasticity is referred to here as the development of acute functional tolerance. This assay has been used to demonstrate that these two phenotypes are distinct and genetically separable. The straightforward locomotion assay described here is suitable for examining the effects of both genetic and environmental manipulations on these acute behavioral responses to ethanol in C.&#160;elegans.

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

C. elegans is a useful model for studying the effects of ethanol on behavior. We present a behavioral assay that quantifies the effects of ethanol on the locomotion speed of crawling worms; both initial sensitivity and the development of acute functional tolerance to ethanol can be measured with this assay.

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the ...

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 ...

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Dissecting the signaling pathways that control the alteration of morphogenic processes during embryonic development requires robust and sensitive metrics. Embryonic elongation of the nematode Caenorhabditis elegans is a late developmental stage consisting of the elongation of the embryo along its longitudinal axis. This developmental stage is controlled by intercellular communication between hypodermal cells and underlying body-wall muscles. These signaling mechanisms control the morphology of hypodermal cells by remodeling the cytoskeleton and the cell-cell junctions. Measurement of embryonic lethality and developmental arrest at larval stages as well as alteration of cytoskeleton and cell-cell adhesion structures in hypodermal and muscle cells are classical phenotypes that have been used for more than 25 years to dissect these signaling pathways. Recent studies required the development of novel metrics specifically targeting either early or late elongation and characterizing morphogenic defects along the antero-posterior axis of the embryo. Here, we provide detailed protocols enabling the accurate measurement of the length and the width of the elongating embryos as well as the length of synchronized larvae. These methods constitute useful tools to identify genes controlling elongation, to assess whether these genes control both early and late phases of this stage and are required evenly along the antero-posterior axis of the embryo.

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Here we detail protocols specifically designed to monitor morphogenic defects that occur during early and late phases of embryonic elongation of the nematode Caenorhabditis elegans. Ultimately, these protocols are designed to identify genes that regulate these phases and to characterize their differential requirements along the antero-posterior axis of the embryo.

Dissecting the signaling pathways that control the alteration of morphogenic processes during embryonic development requires robust and sensitive metrics. ...

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their locomotory patterns upon detection of diverse environmental stimuli and can modulate their suite of behavioral responses in response to starvation conditions. Nematodes typically exhibit a decreased aversive response when removed from a food source for over 30 min. Observation of behavioral changes in response to a changing nutritional status can provide insight into the mechanisms that regulate the transition from a well-fed to starved state.
We have developed an assay that measures a nematode&#39;s ability to cross an aversive barrier (i.e. copper) then reach a food source over a prolonged period of time. This protocol builds upon previous work by integrating multiple variables in a manner that allows for continued data collection as the organisms shift towards an increasingly starved condition. Moreover, this assay permits an increased sample size so that larger populations of nematodes can be simultaneously evaluated.
Organisms defective for the ability to detect or respond to copper immediately cross the chemical barrier, while wild type nematodes are initially repelled. As wild type worms are increasingly starved, they begin to cross the barrier and reach the food source. We designed this assay to evaluate a mutant that is incapable of responding to diverse environmental cues, including food sensation or detection of aversive chemicals. When evaluated via this protocol, the defective organisms immediately crossed the barrier, but were also incapable of detecting a food source. Hence, these mutants repeatedly cross the chemical barrier despite temporarily reaching a food source. This assay can straightforwardly test populations of worms to evaluate potential pathway defects related to aversion and starvation.

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

Here, we present a Caenorhabditis elegans-specific assay designed to evaluate changes in copper aversion behavior and the ability to locate a common food source, as the organism progresses from a well-fed to starved nutritional state.

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their ...

Monitoring the Effect of Osmotic Stress on Secretory Vesicles and Exocytosis

Amperometry recording of cells subjected to osmotic shock show that secretory cells respond to this physical stress by reducing the exocytosis activity and the amount of neurotransmitter released from vesicles in single exocytosis events. It has been suggested that the reduction in neurotransmitters expelled is due to alterations in membrane biophysical properties when cells shrink in response to osmotic stress and with assumptions made that secretory vesicles in the cell cytoplasm are not affected by extracellular osmotic stress. Amperometry recording of exocytosis monitors what is released from cells the moment a vesicle fuses with the plasma membrane, but does not provide information on the vesicle content before the vesicle fusion is triggered. Therefore, by combining amperometry recording with other complementary analytical methods that are capable of characterizing the secretory vesicles before exocytosis at cells is triggered offers a broader overview for examining how secretory vesicles and the exocytosis process are affected by osmotic shock. We here describe how complementing amperometry recording with intracellular electrochemical cytometry and transmission electron microscopy (TEM) imaging can be used to characterize alterations in secretory vesicles size and neurotransmitter content at chromaffin cells in relation to exocytosis activity before and after exposure to osmotic stress. By linking the quantitative information gained from experiments using all three analytical methods, conclusions were previously made that secretory vesicles respond to extracellular osmotic stress by shrinking in size and reducing the vesicle quantal size to maintain a constant vesicle neurotransmitter concentration. Hence, this gives some clarification regarding why vesicles, in response to osmotic stress, reduce the amount neurotransmitters released during exocytosis release. The amperometric recordings here indicate this is a reversible process and that vesicle after osmotic shock are refilled with neurotransmitters when placed cells are reverted into an isotonic environment.

Osmotic stress affects exocytosis and the amount of neurotransmitter released during this process. We demonstrate how combining electrochemical methods together with transmission electron microscopy can be used to study the effect of extracellular osmotic pressure on exocytosis activity, vesicle quantal size, and the amount of neurotransmitter released during exocytosis.

Amperometry recording of cells subjected to osmotic shock show that secretory cells respond to this physical stress by reducing the exocytosis activity ...