These studies were supported by grants from the National Institutes of Health, National Institute for Alcoholism and Alcohol Abuse: R01AA016837 (JCB) and P20AA017828 (AGD and JCB).

1. Steps to Perform on the Day before the Assay
<ol>
	<li>Pick L4 stage worms to fresh nematode growth medium (NGM) plates seeded with a lawn of OP50 E. coli, and culture them at 20 &#730;C O/N. Each assay condition requires 10 worms; pick an excess of worms to allow for O/N loss of worms.
	<ol>
		<li>Only assay animals that are first-day adults; many mutants grow at a slower speed than wild type. Adjust the timing of picking for strains that have developmental delays so that all animals tested are first-day adults.</li>
	</ol>
	</li>
</ol>
2. Steps to Perform on the Day of the Assay
<ol>
	<li>Preparation for the assay:
	<ol>
		<li>Perform assays on standard unseeded 60 x 15 mm NGM plates. Dry all of the plates to be used at 37 &#730;C for 2 hr, with lids off. For each experimental trial, use four dried NGM plates; these will be the 0 mM and 400 mM ethanol assay plates and their accompanying acclimation plates. 
		NOTE: The use of NGM plates is important here; differences in the composition of the plates, in particular their osmolarity, can strongly affect the dose response of ethanol on behavior, this is due, at least in part, to changes in the amount of ethanol accumulated by the animals20. NGM agar is 160 mOsm.</li>
		<li>After drying, weigh each of the unseeded NGM plates to be used in the assay and note the weight. Determine the volume of media in the plates based on the weight of an empty plate. To approximate the conversion of weight of the agar to volume, assume that the media weighs the same as an equal volume of water. 
		NOTE: Our most consistent results have been found with NGM media that has dehydrated sufficiently that an original 10 ml volume has a post-drying volume between 8.3&#8211;8.9 ml. An alternative to the 2 hr dry time is to dry until the plates reach this volume range to account for differences in incubators.</li>
		<li>Melt 4 copper rings (inner diameter of 1.6 cm) into the surface of each of the plates to act as corrals for the different genotypes or treatment groups of worms.Grasp each ring with forceps, and heat in a flame (a strong flame from a Bunsen burner works well) for approximately 3 sec. Immediately place the ring on a plate while still holding the ring with the forceps to prevent it from &#8216;skipping&#8217;.
		<ol>
			<li>Ensure that the ring rests flat against the surface of the agar by pressing down gently with the forceps at several points on the ring. When placing the ring, be careful to leave room for an additional three rings.</li>
			<li>Melt the three additional copper rings into the surface of the plate, taking care to place them as close together as possible so that all four will be in the field of view of the camera. 
			NOTE: Making a good seal with the agar is essential to keep the worms in the rings during the assay. Rings that skip around on the plate are unlikely to make the correct seal with the agar and may scar the agar, which allows worms to burrow and interferes with visualizing the worms.</li>
			<li>For each assay plate prepare an accompanying &#8220;acclimation&#8221; plate, which should be dried and unseeded and will receive no ethanol. Place four copper rings on these plates.</li>
		</ol>
		</li>
		<li>Label the bottom of the plates next to each ring with the worm strain to be used in the ring, taking care to not write in the field of view of the ring itself. Match the labels on the assay plate with its accompanying acclimation plate.</li>
		<li>Calculate the amount of 100% ethanol to add to each assay plate so that the final concentration is 0 mM or 400 mM ethanol (weight to volume). For each experiment (n = 1), there will be one 0 mM and one 400 mM ethanol plate; acclimation plates do not receive ethanol. Add the ethanol, pipetting it around the surface of the plate. Use laboratory film to seal the plate and allow it to equilibrate on the bench top for 2 hr.</li>
		<li>When 1.5 hr has elapsed, begin the acclimation step of the assay, step 2.2.</li>
	</ol>
	</li>
	<li>Perform locomotion assay: Assay plates
	<ol>
		<li>Carefully transfer 10 worms of each experimental group to the center of a copper ring on the acclimation plate. Remove any food that is visible on the agar at this point by scraping gently with the worm pick. Vary the order in which the experimental groups are put on the plates across experimental trials so that the same strains are not put on the plates in the same order across trials. 
		NOTE: The goal is to transfer the animals to the plate with minimal quantities of food, because if food on the acclimation plate is transferred to the assay plate the worms will aggregate around the food and the effect of the drug on locomotion will be obscured.</li>
		<li>Be careful not to break the surface of the agar with the pick, as this will allow the worms to burrow and will disrupt the assay. Incubate the worms at RT for 30 min.</li>
		<li>Ensure that there is an appropriate interval between the initiations of each plate. An experienced experimenter can move 40 animals, 10/ring for 4 rings in &#60; 1.5 min, but any interval up to 2 min&#160;is acceptable. The standard assay has movies recording at 10-12 min&#160;and 30-32 min&#160;of exposure for both 0 mM and 400 mM ethanol.</li>
		<li>Initiate the acclimation plate for the 0 mM exposure and the acclimation plate for the 400 mM exposure approximately 2 min&#160;and 30 sec&#160;apart to allow for saving of the first movie file before the second movie must begin.</li>
		<li>After the 30-min&#160;acclimation period, transfer the worms from the acclimation plates to the assay plates. Transfer the worms to the assay plates (0 mM or 400 mM ethanol) in the same order that they were added to the acclimation plate, keeping track of the timing between the completions of each plate. Seal the plate with laboratory film to minimize loss of ethanol to vaporization.</li>
		<li>Use a scooping motion with a thin-edged flattened worm pick to collect worms on top of the flattened pick. Perform the transfer of worms from the unseeded acclimation plate to the assay plate without the use of bacteria, which is commonly used in transferring worms as it helps to glue the worm to the pick. 
		NOTE: The speed at which animals are transferred at this step is very important because the first worms on the plate will be exposed to ethanol longer than the last worms added to the plate, and earlier time points may show some time-dependent effects. This is the major reason for rotating which strains are placed on a plate first across experimental replicates.</li>
	</ol>
	</li>
	<li>Perform Locomotion Assay: Filming
	<ol>
		<li>Use a microscope/camera combination that allows for simultaneous imaging of all four rings in the field of view (approximately 42x42 mm2 square), such as a 0.5x microscope objective, 0.8x magnification and a camera with a 2048x2048 7.4 &#181;m pixel CCD.</li>
		<li>Use even illumination across the field of view, which aids in object recognition at the intensity threshold step (see below). A 3&#8221;x3&#8221; backlight works well.</li>
		<li>Image the worms on a plate positioned media-side up (lid down), which generates contrast that is lost when using the backlight compared with a traditional microscope transmitted light source.</li>
		<li>Prepare the image analysis software to capture time-lapse movies of the moving worms. Set the software to capture 12-bit gray-scale images every 1 sec, as a 2-min&#160;(120 frame) movie. To reduce the size of the output file, while still retaining sufficient image resolution, use a 2x2 binning mode to capture 1024x1024 pixel images.</li>
		<li>Record the movies. Begin recording a 120-frame movie of the first plate (0 mM ethanol) 10 min&#160;after the last worms were placed on that plate. Save the movie file.</li>
		<li>Record the second plate (400 mM ethanol). Repeat this process for both plates beginning 30 min&#160;after the last worms were placed on the first plate (0 mM ethanol) to capture the 30&#8211;32 min&#160;time points for each exposure. 
		NOTE: Allow sufficient time to save the image files after each capture session before beginning recording of the next plate to be analyzed.</li>
		<li>For future-proofing of archived movies and to allow these movies to be opened and analyzed by other public domain open source imaging software programs, such as ImageJ28, convert a copy of each movie to 8-bit 256 gray scale TIFF files. 
		NOTE: The image analysis software described here uses a proprietary file format.</li>
	</ol>
	</li>
	<li>Analysis of movies using ImagePro software.
	<ol>
		<li>Once captured, analyze the movies using Image-Pro Plus software or its equivalent. 
		NOTE: A variety of other object tracking software is available that has been used to successfully track multiple C. elegans at one time29 and such software could reasonably substitute for that described here as the object identification steps are based on similar principles. The method described relates to Image-Pro Plus software versions 6.0-6.3 and 7.0, newer versions of Image-Pro Plus software may have minor differences.</li>
		<li>Analyze movies in 2-min&#160;(120 frame) segments. First, apply a filter to the images to flatten the background and enhance the contrast of the worm objects. Select the filter as follows: Menu&#62;Process&#62;Filters&#62;Enhancement&#62;Flatten: Parameters: Background = Dark; Feature Width = 20 Pix.</li>
		<li>Re-save the movies after the filtering step but always retain the original movie in its unfiltered form.
		<ol>
			<li>Analyze the locomotion of animals in each ring separately with a circular region of interest (ROI) that is placed and sized to overlap with the copper ring. Identify and track the worms with the Menu&#62;Measure&#62;Track Objects&#8230; command; this brings up the Tracking Data Table window. The Tracking Options button allows specific tracks to be excluded and to limit any experimental artifacts.</li>
		</ol>
		</li>
		<li>Under the Auto Tracking tab, use the following parameters: Track Parameters: Velocity limit (search radius) = 400 &#181;m/frame, Acceleration limit = Auto, Minimum total track length = 400 &#181;m, Predominant motion type = chaotic; Objects in tracks parameters: Allow partial tracks = yes, Min length = 21 frames, Tracking prediction depth = 1 frame.</li>
		<li>To initiate the tracking process, click the Find All Tracks Automatically function button to bring up the Count/Size options dialog box and the Tracking dialog box. Select the Manual option for the Intensity Range Selection in the Count/Size dialog box, which provides the important threshold step that uses the gray-scale intensity of the worm pixels to highlight a worm object for analysis and to ignore the lighter background pixels.
		<ol>
			<li>Adjust the intensity threshold sliders (one to set the upper limit, one to set the lower limit) to create an inclusive range that highlights all dark objects. A range of 0&#8211;1,500 on a scale of 0&#8211;4,095 is a good starting point for finer tuning.</li>
			<li>Apply a size filter to exclude objects that are larger and smaller than a single worm object, such as the copper ring and any debris on the plates. Set two size parameters to do this that cover sizes for individual worms in a variety of postures with the Measure&#62;Select Measurements menu item on the Count/Size options dialog box. (Area range: 28,000&#8211;120,000 &#181;m2 and Perimeter range: 600&#8211;2,500 &#181;m).</li>
			<li>If a mutant strain is significantly smaller or larger than other animals on the plate to be analyzed, broaden the range of the filter settings for object recognition to accommodate the different sizes of the strains first before tracking animals so that settings do not need to be changed mid-analysis.</li>
		</ol>
		</li>
		<li>Complete the tracking process by clicking Continue in the Tracking dialog box. Visually compare the output tracks with the progress of each worm in the movie to ensure that every worm is represented unless there is a valid reason to exclude it based on the automatic filter settings. Manually delete tracks that were produced by the presence of confirmed non-worm objects that met the size filter criteria.</li>
		<li>Use the software to calculate the velocity of each worm between each frame (distance traveled for the centroid of an object per 1 sec&#160;between frames) and display the average velocity for each track and the average velocity across all tracks for the population of worms in the copper ring. Consider this final average to be n = 1 in terms of experimental trials. Export the data to a spreadsheet program for statistical analyses and data archiving.</li>
		<li>Once the data has been recorded for the first ring on the plate, move the ROI to the next ring and repeat the tracking process, without changing any of the parameters.</li>
		<li>Calculate relative speed of locomotion through:</li>
		<li>Relative speed (%) = treated (400 mM) speed/ untreated (0 mM) speed x 100. 
		NOTE: Different genetic manipulations often alter the basal locomotion rate of animals. In order to determine the effect of ethanol on the locomotion speed of animals and to be able to compare these effects across different genotypes or conditions, calculate a relative speed.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The simple neurobiology and genetic tools available in C. elegans make the worm an excellent model in which to study the molecular bases of the effects of ethanol on behavior. Here, we describe an assay that has been used to identify several molecular and environmental mediators of the acute behavioral response to ethanol8,10,20,24,25. This method allows the differentiation and simultaneous examination of two different ethanol response behavior phenotypes, initial sensitivity and the development of ac...

Alcohol use disorders (AUD) are widespread and produce serious health, social, and economic problems. In humans, the susceptibility to developing an AUD is heavily influenced by both genetics and the environment1,2. A strong physiological predictor of abuse liability is the initial level of response (LR) to alcohol (ethanol) that is exhibited by na&#239;ve drinkers3-5. This LR phenotype is influenced by genetics and non-genetic components6. Determining the molecular mechanisms that influence the LR to ethanol is an important goal of the study of ethanol response behaviors.
The nematode, Caenorhabditis elegans, has been increasingly used as a model for studying the effects of ethanol on behavior7-9. There is strong molecular conservation in the machinery of nervous system function between worms and mammals, and several genes that have been shown to influence the LR to ethanol in worms have been shown to influence LR to ethanol in mammals10-16, and have been implicated in abuse liability in humans17-19. 
Ethanol intoxicates worms, which is reflected in a decrease in their locomotion speed. Several different laboratories have developed behavioral assays that differ in several ways, for example, the locomotion behavior that they study (crawling versus swimming11,12,14,20,21) or in the composition of the solutions in which the assays are performed (nematode growth medium versus Dent&#8217;s saline20,22). Interestingly, these diverse assays have yielded somewhat different dose response profiles for the effects of ethanol. These results have pointed to important differences in the underlying behaviors of crawling and swimming9,23, as well as a role for the environmental variable osmolarity in ethanol responses20, and have highlighted the importance of describing experimental detail of the various assays.
An assay to measure the acute effects of ethanol on crawling behavior is presented here. This assay has been used extensively to study the genetic and environmental influences on the LR to ethanol8,10,20,24,25. The mammalian LR phenotype is a composite of at least two components, initial sensitivity to ethanol and acute functional tolerance to ethanol26,27. In worms, the LR phenotype has been shown to be separable into these two components through the use of this behavioral assay. The influences of genetic and environmental manipulations on both phenotypes can be examined using this single assay. Importantly, these two phenotypes are genetically separable.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>C. elegans strains</td>
			<td>Caenorhabditis Genetics Center</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>60 x15 mm Petri plates, triple vented</td>
			<td>Greiner Bio-One</td>
			<td>628161</td>
			<td>Other plate brands will suffice.</td>
		</tr>
		<tr>
			<td>NGM agar</td>
			<td>Various</td>
			<td></td>
			<td>NaCl (3g/L), agar (17g/L), peptone (2.5g/L), 1 mL cholesterol (5mg/mL in ethanol), 1 mL (1M) MgSO4, 1 mL (1M) CaCl2, 25 mL (1M) KPO4, pH=6, 975 mL H2O</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Various</td>
			<td></td>
			<td>e.g. Fisher Scientific #10300</td>
		</tr>
		<tr>
			<td>37&#176;C Incubator</td>
			<td>Various</td>
			<td></td>
			<td>For drying agar</td>
		</tr>
		<tr>
			<td>Digital balance</td>
			<td>Various</td>
			<td></td>
			<td>For determining plate weights and agar volume</td>
		</tr>
		<tr>
			<td>Copper rings</td>
			<td>Plumbmaster</td>
			<td>STK#35583 (48 cap thread gasket)</td>
			<td>1.6 cm inner diameter, 1.8 cm outer diameter copper rings</td>
		</tr>
		<tr>
			<td>100% ethanol</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm M</td>
			<td>Bemis</td>
			<td>PM996</td>
			<td></td>
		</tr>
		<tr>
			<td>CCD camera</td>
			<td>QImaging</td>
			<td>RET-4000R-F-M-12</td>
			<td>This camera has a large field of view.</td>
		</tr>
		<tr>
			<td>Stereomicroscope with C-mount and 0.5X objective</td>
			<td>Leica</td>
			<td>MZ6</td>
			<td>Discontinued model, M60 is current equivalent.</td>
		</tr>
		<tr>
			<td>Light source</td>
			<td>Schott</td>
			<td>A08923</td>
			<td>3&#8221;x3&#8221;&#160; backlight for even illumination across the field of view</td>
		</tr>
		<tr>
			<td>Imaging and tracking software</td>
			<td>Media Cybernetics</td>
			<td>ImagePro-Plus v6.0-6.3</td>
			<td>Newer versions of the software have tracking functions.</td>
		</tr>
	</tbody>
</table>

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.

Representative data (Figure 1) from several different genotypes and their paired controls are presented8,24; data were specifically chosen that highlight differences in the assayed animals. The degree of effect at 10 min of exposure is considered the initial sensitivity of a strain, which is shown on the left axes in Figure 1B-G. Mutant strains with a relative speed larger than the control at 10 min&#160;are considered to be ethanol resistant (Figure 1F, G), w...

Watch this Scientific Journal Video about An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans at JoVE.com

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.

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 ...

an-assay-for-measuring-effects-ethanol-on-locomotion-speed

Research

JoVE Journal

Neuroscience

8.6K Views.  Virginia Commonwealth University. 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. 

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

Microdialysis of Ethanol During Operant Ethanol Self-administration and Ethanol Determination by Gas Chromatography

Operant self-administration methods are commonly used to study the behavioral and pharmacological effects of many drugs of abuse, including ethanol. However, ethanol is typically self-administered orally, rather than intravenously like many other drugs of abuse. The pharmacokinetics of orally administered drugs are more complex than intravenously administered drugs. Because understanding the relationship between the pharmacological and behavioral effects of ethanol requires knowledge of the time course of ethanol reaching the brain during and after drinking, we use in vivo microdialysis and gas chromatography with flame ionization detection to monitor brain dialysate ethanol concentrations over time. 
Combined microdialysis-behavioral experiments involve the use of several techniques. In this article, stereotaxic surgery, behavioral training and microdialysis, which can be adapted to test a multitude of self-administration and neurochemical centered hypotheses, are included only to illustrate how they relate to the subsequent phases of sample collection and dialysate ethanol analysis. Dialysate ethanol concentration analysis via gas chromatography with flame-ionization detection, which is specific to ethanol studies, is described in detail. Data produced by these methods reveal the pattern of ethanol reaching the brain during the self-administration procedure, and when paired with neurochemical analysis of the same dialysate samples, allows conclusions to be made regarding the pharmacological and behavioral effects of ethanol.

A method to determine the time course of ethanol concentration in the brains of rats during operant ethanol self-administration is described. Gas chromatography with flame ionization detection is used to quantify ethanol in the dialysate samples, because it has the sensitivity required for the small volumes that are generated.

Operant self-administration methods are commonly used to study the behavioral and pharmacological effects of many drugs of abuse, including ethanol.  ...

An Assay for Permeability of the Zebrafish Embryonic Neuroepithelium

The brain ventricular system is conserved among vertebrates and is composed of a series of interconnected cavities called brain ventricles, which form during the earliest stages of brain development and are maintained throughout the animal's life. The brain ventricular system is found in vertebrates, and the ventricles develop after neural tube formation, when the central lumen fills with cerebrospinal fluid (CSF) 1,2. CSF is a protein rich fluid that is essential for normal brain development and function3-6.
In zebrafish, brain ventricle inflation begins at approximately 18 hr post fertilization (hpf), after the neural tube is closed. Multiple processes are associated with brain ventricle formation, including formation of a neuroepithelium, tight junction formation that regulates permeability and CSF production. We showed that the Na,K-ATPase is required for brain ventricle inflation, impacting all these processes 7,8, while claudin 5a is necessary for tight junction formation 9. Additionally, we showed that "relaxation" of the embryonic neuroepithelium, via inhibition of myosin, is associated with brain ventricle inflation.
To investigate the regulation of permeability during zebrafish brain ventricle inflation, we developed a ventricular dye retention assay. This method uses brain ventricle injection in a living zebrafish embryo, a technique previously developed in our lab10, to fluorescently label the cerebrospinal fluid. Embryos are then imaged over time as the fluorescent dye moves through the brain ventricles and neuroepithelium. The distance the dye front moves away from the basal (non-luminal) side of the neuroepithelium over time is quantified and is a measure of neuroepithelial permeability (Figure 1). We observe that dyes 70 kDa and smaller will move through the neuroepithelium and can be detected outside the embryonic zebrafish brain at 24 hpf (Figure 2).
This dye retention assay can be used to analyze neuroepithelial permeability in a variety of different genetic backgrounds, at different times during development, and after environmental perturbations. It may also be useful in examining pathological accumulation of CSF. Overall, this technique allows investigators to analyze the role and regulation of permeability during development and disease.

We describe a live whole animal quantitative measurement for permeability of the embryonic zebrafish brain. The technique analyzes the ability to retain cerebrospinal fluid and molecules of different molecular weights within the neural tube lumen and quantifies their movement out of the ventricles. This method is useful for determining differences in epithelial permeability and maturation during development and disease.

The brain ventricular system is conserved among vertebrates and is composed of a series of interconnected cavities called brain ventricles, which form ...

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 ...

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 Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in their etiology. The fruit fly, Drosophila melanogaster, is a powerful model for exploring molecular-genetic mechanisms underlying alcohol-related behaviors and therefore holds great promise for identifying and understanding the function of genes that influence AUD. The use of the Drosophila model for these types of studies depends on the availability of assays that reliably measure behavioral responses to ethanol. This report describes an assay suitable for assessing ethanol sensitivity and rapid tolerance in flies. Ethanol sensitivity measured in this assay is influenced by the volume and concentration of ethanol used, a variety of previously reported genetic manipulations, and also the length of time the flies are housed without food immediately prior to testing. In contrast, ethanol sensitivity measured in this assay is not affected by the vigor of fly handling, sex of the flies, and supplementation of growth medium with antibiotics or live yeast. Three different methods for quantitating ethanol sensitivity are described, all leading to essentially indistinguishable ethanol sensitivity results. The scalable nature of this assay, combined with its overall simplicity to set-up and relatively low expense, make it suitable for small and large scale genetic analysis of ethanol sensitivity and rapid tolerance in Drosophila.

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Straightforward assays for measuring ethanol sensitivity and rapid tolerance in Drosophila facilitate the use of this model organism for investigating these important ethanol-related behaviors. Here, a relatively simple, scalable assay for measuring ethanol sensitivity and rapid tolerance in flies is described.

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in ...

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 ...

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the over-production of amyloid-beta and the deposition of amyloid-beta plaques in brain regions of the afflicted individuals. Throughout the years scientists have generated numerous Alzheimer&#8217;s disease mouse models that attempt to replicate the amyloid-beta pathology. Unfortunately, the mouse models only selectively mimic the disease features. Neuronal death, a prominent effect in the brains of Alzheimer&#8217;s disease patients, is noticeably lacking in these mice. Hence, we and others have employed a method of directly infusing soluble oligomeric species of amyloid-beta - forms of amyloid-beta that have been proven to be most toxic to neurons - stereotaxically into the brain. In this report we utilize male C57BL/6J mice to document this surgical technique of increasing amyloid-beta levels in a select brain region. The infusion target is the dentate gyrus of the hippocampus because this brain structure, along with the basal forebrain that is connected by the cholinergic circuit, represents one of the areas of degeneration in the disease. The results of elevating amyloid-beta in the dentate gyrus via stereotaxic infusion reveal increases in neuron loss in the dentate gyrus within 1 week, while there is a concomitant increase in cell death and cholinergic neuron loss in the vertical limb of the diagonal band of Broca of the basal forebrain. These effects are observed up to 2 weeks. Our data suggests that the current amyloid-beta infusion model provides an alternative mouse model to address region specific neuron death in a short-term basis. The advantage of this model is that amyloid-beta can be elevated in a spatial and temporal manner.

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the ...

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. ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

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 ...

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

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Microdialysis of Ethanol During Operant Ethanol Self-administration and Ethanol Determination by Gas Chromatography

An Assay for Permeability of the Zebrafish Embryonic Neuroepithelium

A Caenorhabditis elegans Model System for Amylopathy Study

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

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay