We would like to thank Chris Wreden and Michelle Bland for comments on the manuscript and for technical help.

1. Preparation of Larvae
<ol>
	<li>One week before recording behavior, set up a cross (minimum of 25 virgins and 5 males). Maintain all crosses and progeny at 25 &#176;C. 
	NOTE: The temperature of culture conditions can be altered, but the time line described below would need to be adjusted to account for changes in developmental speed.</li>
	<li>5 days before recording, first thing in the morning, place the cross into a collection cage with an agar/juice cap and a dab (0.5-1 ml) of yeast paste in the center of the agar/juice cap.
	<ol>
		<li>To make a collection cage, poke holes into a 6 oz. square polyethylene Drosophila bottle.</li>
		<li>To make agar/juice caps, mix 18 g of agar with 600 ml water in a conical flask. In a separate flask mix 20 g sucrose with 200 ml apple juice. Microwave until solids are dissolved. Combine and stir, allowing mixture to cool to 60 &#176;C. Add 20 ml 10% methyl-p-hydroxybenzoate in 95% ethanol, and stir. Add ~7 ml to the bottom half of a 35 x 10 mm round petri dish. Allow agar/juice to solidify for 1 hr at RT. Store at 4 &#176;C.</li>
		<li>To make yeast paste, add equal parts by volume dry yeast and water. Store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Check the health of larval collections.
	<ol>
		<li>In the morning, 4 and 3 days before recording, remove the cap and place a fresh cap on the cage. Change the caps at the same time each day to get a 24 hr collection of eggs and newly hatched larva.</li>
		<li>After each cap is removed, examine it to see how many eggs the cross is laying. Expect between 500-2,000 eggs. Adjust the number of adults, if needed. 
		NOTE: A few eggs should have hatched into larvae, and the newly-hatched larvae will be found away from the yeast paste.</li>
		<li>Age each cap for 24 hr. Re-examine the cap to determine whether larvae are healthy. 
		NOTE: Most eggs should have hatched into larvae, and the larvae should have crawled into the yeast paste. Signs of poor health include a large number of unhatched eggs, dead larval bodies away from the yeast paste, and larvae with significant dark patches in the abdomen, indicating an immune reaction. If larvae are unhealthy, change the cage, make fresh caps/yeast paste, and check the genotype.</li>
		<li>Discard the old caps.</li>
	</ol>
	</li>
	<li>Collect larvae for behavioral recordings.
	<ol>
		<li>At both 2 and 1 day before recording, remove the cap and replace with a fresh cap.</li>
		<li>Label each removed cap and keep at 25 &#176;C. This will produce a staged series of larvae for behavioral recordings. Larvae from newly-hatched (0-4 hr posthatch) through second instar (24-48 hr posthatch) can be used in the channels. Save caps for recordings on consecutive days.</li>
		<li>Discard all caps after 48 hr.</li>
		<li>If needed, repeat steps 1.4.1-1.4.3 for up to 5 additional d. After that fewer fertilized eggs are produced.</li>
	</ol>
	</li>
</ol>
2. Preparation of Channels
<ol>
	<li>Prepare the linear channel PDMS (Polydimethylsiloxane, a silicone elastomer) casting mold (Figure 1A). PDMS casting molds are available from the Heckscher lab upon request.
	<ol>
		<li>Clean the casting mold with isopropyl alcohol. Air dry, dab dry with laboratory wipes, or use canned air.</li>
		<li>Place the casting mold into a 10 cm Petri dish lid or other container.</li>
	</ol>
	</li>
	<li>Prepare and pour agarose solution.
	<ol>
		<li>Make a 3% agarose solution in water (3 g in 100 ml) by microwaving until fully dissolved.</li>
		<li>Cool to 55 &#176;C, allowing bubbles to rise to surface.</li>
		<li>Pour the agarose mixture into the petri dish containing the casting mold so that the mold is just covered. Let agarose set until solidified.</li>
	</ol>
	</li>
	<li>Remove the agarose channels from the casting mold. Trim the edges of the agarose disc with a razor blade.
	<ol>
		<li>Put the channels in a 10 cm Petri dish immersed in water. They can be kept at 4 &#176;C for 7 days.</li>
	</ol>
	</li>
</ol>
3. Loading a Larva into a Channel to Record Behavior
NOTE: If storing channels at 4 &#176;C, allow channels to come to RT before using for behavioral recording.
<ol>
	<li>Use a clean razor blade to cut a single channel from those prepared in Step 2 (Figure 2A).</li>
	<li>Use a pair of forceps to place the channel grooved-side-up on a glass coverslip (Figure 2B). The size and thickness of the coverslip depends on the specific scope used for imaging.</li>
	<li>Use fine forceps to select a larva from the agar/juice cap prepared in Step 1 and place into the cut channel (Figure 2C).
	<ol>
		<li>Do not squeeze the larva. Pick it up gently with the tip of the forceps tine. It is also possible to manipulate larva with a paintbrush with a fine point. 
		NOTE: This is important as larvae are very sensitive to touch. If they are not handled with care, nociceptive responses might dominate their behavior during the first few seconds of the experiment.</li>
		<li>Wash the larvae by briefly submerging them in water.</li>
		<li>After placing the larva onto the cut channel,&#160;use the tip of the forceps tine to gently tease the larva into the channel grove. 
		NOTE: For most applications the width of the larva and the width of the channel should match. The average width of larvae at different stages of development follows: 0 hr posthatch -&#160;140 &#181;m, 24 hr posthatch -&#160;180 &#181;m, 48 hr posthatch -&#160;320 &#181;m, 72 hr posthatch -&#160;500 &#181;m, 96 hr posthatch -&#160;750 &#181;m8. Channels come in the following widths: 100, 150, 200, 250, and 300 &#181;m, and all channels are 150 &#181;m&#160;deep.</li>
		<li>Use the tip of a forceps tine or paintbrush to adjust the position of the larva within the channel. It can be oriented in any direction: ventral up, ventral down, ventral to one side, depending on what structure is to be imaged.</li>
	</ol>
	</li>
	<li>Using forceps, gently flip the channel over such that the larva is now on the cover glass. Place one or two drops of water at the ends of the channel so that it fills with water (Figure 2D).
	<ol>
		<li>Gently lift the sides of the channel to remove any air bubbles.</li>
		<li>Adjust the orientation of the larva. To do so, gently nudge the channel but not the cover glass to roll the larva. If this does not change the orientation, remove the cover glass and adjust larval position. The larva is ready to be imaged.</li>
		<li>Image the channel/coverslip on an inverted microscope. If imaging on an upright microscope, simply invert the channel/coverslip.</li>
	</ol>
	</li>
</ol>
4. Measure Feature of Interest in Behavioral Recording
<ol>
	<li>For a structure of interest (e.g., tail, CNS), measure a feature of interest (e.g., fluorescence intensity, location) over time. 
	NOTE: Structures such as the head and tail can be manually annotated. For example, the head can be identified as containing dark H-shaped mouth hooks, and the tail can be identified as containing posterior tracheal spiricles. This paper focuses on the nerve cord. Use the neuronal GAL4 line elav-GAL4 to drive a UAS-myr-GFP transgene (elav&#62;GFP)9,10 to fluorescently label the central nervous system (CNS). The CNS contains two anterior brain lobes attached to the nerve cord, which extends to the posterior (Figure 3).</li>
	<li>Measure the pixel intensity of the nerve cord (f(NC)) at each time point (t). Below describes a manual annotation approach, but this step could be automated.
	<ol>
		<li>In Fiji (or similar software) in the &#39;Analyze&#39; menu use the &#34;Set Measurements ...&#34; dialogue box to report the mean gray value and slice position.</li>
		<li>Manually draw a box in the center of the nerve cord in the first frame of the movie (Figure 3). This step could also be automated with image segmentation and registration algorithms.</li>
		<li>Use the &#34;Measure&#34; command in the &#39;Analyze&#39; menu to report the average pixel intensity in the boxed region of interest. This generates a &#34;Results&#34; window.</li>
		<li>In the next frame, manually reposition the box without resizing by using the arrow keys. Use the &#34;Measure&#34; command again. The updated results will be shown in the &#34;Results&#34; window. Repeat this step for each frame of interest.</li>
		<li>Save the &#34;Results&#34; using the &#39;Save As...&#39; command from the File menu. The results will be exported as an .xls file.</li>
	</ol>
	</li>
</ol>
5. Analyze the Measurements
<ol>
	<li>Normalize each f(NC) value to a value between 0 and 1 (z(t)).
	<ol>
		<li>In spreadsheet (or other program), open the Results.xls file. This file will have two columns representing frame number and average fluorescence intensity.</li>
		<li>Make a new column and use the formula z(t) = f(NC[t])-min(f[NC])/max(f[NC])-min(f[NC]) to generate a normalized fluorescence value corresponding to each time point.</li>
	</ol>
	</li>
	<li>Determine the stride period for each stride in the recording. 
	NOTE: A stride cycle is the unit of repetitive whole body motion for forward (and reverse) crawling. By convention a forward stride starts (and ends) with the forward movement of the tail, head, and other internal organs such as the gut or CNS1.
	<ol>
		<li>Manually inspect the behavioral recording and record the initiation times (i) of each stride (i(Stride Cycle[n])). 
		NOTE: In the example below, forward movement of the CNS as the initiation of a stride cycle was used (Figure 3).</li>
		<li>For each stride calculate the period: &#916;t(Stride cycle) = i(Stride Cycle [n+1])-i(Stride Cycle [n]).</li>
	</ol>
	</li>
	<li>Calculate percentage of stride cycle elapsed for each time point of the recording.
	<ol>
		<li>Make an additional column in the spreadsheet file representing time elapsed since the start of a stride cycle (c). Set time since stride initiation to zero at the initiation of each stride (c(Stride Cycle[n]) = 0). Adjust times after initiation accordingly.</li>
		<li>Make an additional column in the spreadsheet file representing percent stride cycle elapsed (% stride cycle). Divide adjusted times by stride period and multiply by 100 to convert to percentage of stride cycle elapsed: % stride cycle = (c(Stride Cycle[n]))/ (&#916;t(Stride cycle)*100).</li>
	</ol>
	</li>
</ol>
6. Generate Polar Coordinate Plots to Represent Dynamics of Structures of Interest over the Crawl Cycle
<ol>
	<li>In MATLAB, use the &#39;Import Data&#39; dialogue in the Home tab to import the updated Results.xls.</li>
	<li>Make a new array &#34;Theta&#34; containing percent stride cycle values (% stride cycle). (&#34;Theta = Var1&#34; where Var1 represents % stride cycle).</li>
	<li>Make a new array &#34;Rho&#34; contain normalized fluorescence values (z(t)). (&#34;Rho = Var2&#34; where Var2 represents z(t)).</li>
	<li>Use the &#39;polarplot&#39; command to make polar coordinate plots with the percent stride cycle as the angle, and the fluorescence as the radius (&#34;polarplot(Theta, Rho)&#34;).</li>
</ol>

<ol>
	<li>Heckscher, E. S., Lockery, S. R., Doe, C. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+Drosophila+larval+crawling+at+the+level+of+organism,+segment,+and+somatic+body+wall+musculature.">Characterization of Drosophila larval crawling at the level of organism, segment, and somatic body wall musculature.</a> J Neurosci. 32 (36), 12460-12471 (2012).</li><li>Marder, E., Calabrese, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+of+rhythmic+motor+pattern+generation.">Principles of rhythmic motor pattern generation.</a> Physiol rev. 76 (3), 687 (1996).</li><li>Mullins, O. J., Hackett, J. T., Buchanan, J. T., Friesen, W. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+control+of+swimming+behavior:+Comparison+of+vertebrate+and+invertebrate+model+systems.">Neuronal control of swimming behavior: Comparison of vertebrate and invertebrate model systems.</a> Prog Neurobiol. 93 (2), 244-269 (2011).</li><li>Ohyama, T., 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+multilevel+multimodal+circuit+enhances+action+selection+in+Drosophila.">A multilevel multimodal circuit enhances action selection in Drosophila.</a> Nature. 520 (7549), 633-639 (2015).</li><li>Landgraf, M., Thor, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+Drosophila+motoneurons:+specification+and+morphology.">Development of Drosophila motoneurons: specification and morphology.</a> Semin cell devl bio. 17 (1), 3-11 (2006).</li><li>Heckscher, E. S., 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=Even-Skipped(+)+Interneurons+Are+Core+Components+of+a+Sensorimotor+Circuit+that+Maintains+Left-Right+Symmetric+Muscle+Contraction+Amplitude.">Even-Skipped(+) Interneurons Are Core Components of a Sensorimotor Circuit that Maintains Left-Right Symmetric Muscle Contraction Amplitude.</a> Neuron. 88 (2), 1-16 (2015).</li><li>Green, C. H., Burnet, B., Connolly, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+and+patterns+of+inter-and+intraspecific+variation+in+the+behaviour+of+Drosophila+larvae.">Organization and patterns of inter-and intraspecific variation in the behaviour of Drosophila larvae.</a> Anim Behav. 31 (1), 282-291 (1983).</li><li>Graf, S. A., Sokolowski, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rover/Sitter+Drosophila+melanogaster+Larval+Foraging+Polymorphism+as+a+Function+of+Larval+Development,+Food-Patch+Quality,+and+Starvation.">Rover/Sitter Drosophila melanogaster Larval Foraging Polymorphism as a Function of Larval Development, Food-Patch Quality, and Starvation.</a> J Insect Behav. 2 (3), 301-313 (1989).</li><li>Lee, T., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosaic+analysis+with+a+repressible+cell+marker+for+studies+of+gene+function+in+neuronal+morphogenesis.">Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis.</a> Neuron. 22 (3), 451-461 (1999).</li><li>Rebay, I., Rubin, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Yan+Functions+as+a+General+Inhibitor+of+Differentiation+and+Is+Negatively+Regulated+by+Activation+of+the+Rasl+/+MAPK+Pathway.">Yan Functions as a General Inhibitor of Differentiation and Is Negatively Regulated by Activation of the Rasl / MAPK Pathway.</a> Cell. 81 (6), 857-866 (1995).</li><li>Chen, T. -. W., 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=Ultrasensitive+fluorescent+proteins+for+imaging+neuronal+activity.">Ultrasensitive fluorescent proteins for imaging neuronal activity.</a> Nature. 499 (7458), 295-300 (2013).</li><li>Tufte, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The visual display of quantitative information. , (2004).</li></ol>

The authors have nothing to disclose.

A microfluidic device was built to make linear agarose channels that can accommodate Drosophila larvae (Figure 1). When Drosophila larvae are placed in these linear agarose channels their behavioral repertoire is limited to crawling, which allows for detailed observation of the dynamics of larval structures over the crawl cycle.
A successful recording occurs when a larva perform a series of rhythmic strides (Figure 3). If this does not occur,...

The overall goal of this method is to study Drosophila larval crawling in detail. Experiments on locomotion have played an important role in developing and testing theories on motor control2. Traditionally locomotion has been studied in aquatic animals (e.g., leech, lamprey, tadpole)3. The repetitive nature of locomotion in these animals has allowed for the study of rhythmogenesis, for analysis of the biophysical events driving locomotion, and for monitoring the neural firing patterns that accompany locomotion.
The use of Drosophila larvae for studies of locomotion presents a unique combination of advantages over other model systems: facile genetics, well-characterized development, a body that is optically clear at first and second instars, and an ongoing transmission electron microscopic reconstruction of the entire nervous system4-6. However, Drosophila larval locomotion on flat open surfaces is somewhat complex including pauses, turns, and meandering crawls7. This publication presents a method to use linear agarose channels to guide Drosophila larval locomotor behavior such that larvae perform sustained, straight, rhythmic crawling behavior.
Studying Drosophila larval behavior in agarose channels, instead of behavior on flat open surfaces, has several advantages. First, it allows researchers to specifically select crawling behavior from the many movements that are part of the larval behavioral repertoire. Second, by adjusting the width of the channel versus the larval body size, crawling speed can be adjusted. Third, channels allow for the larva to be viewed from dorsal, ventral, or lateral side depending on how the larva is loaded and oriented within the channel. This versatility in larval orientation allows for any structure of interest to be continually visible during crawling. Fourth, channels are amenable for use with a wide variety of microscopes and objectives. For example, linear channels can be used for low-resolution imaging on bright-field stereoscopes and/or for high-resolution imaging on spinning-disc confocal microscopes1. Fifth, this method can be used in combination with optogenetic/thermogenetic neuronal manipulations in any genetic background. Finally, because both the larval body (at first and second instars) and agarose channels are optically clear, channels can be used when studying the dynamic movements, or changes in fluorescent intensity of larval structures labeled by genetically-encoded fluorescent probes.
The method described is appropriate for detailed kinematic studies of first and second instar Drosophila larval behavior. This publication analyzes the dynamic changes in fluorescent intensity of the CNS during forward larval crawling to demonstrate the use of channels and as a precursor to neuronal calcium imaging.

<table><tbody><tr><td>6 oz square Drosophila bottle</td><td>Scimart</td><td>DR-103</td><td></td></tr><tr><td>agar</td><td>sigma</td><td>A1296</td><td></td></tr><tr><td>sucrose</td><td>sigma</td><td>S9378</td><td></td></tr><tr><td>apple juice</td><td></td><td></td><td>not from concentrate</td></tr><tr><td>Tegosept</td><td>Fisher</td><td>T2300</td><td>methyl-p-hydroxybenzoate</td></tr><tr><td>35 x 10 mm round petri dish</td><td>Fisher</td><td>351008</td><td></td></tr><tr><td>baker&#039;s yeast</td><td></td><td></td><td></td></tr><tr><td>PDMS casting mold</td><td>FlowJem</td><td></td><td>can be requested from authors</td></tr><tr><td>Isopropyl alcohol</td><td>Fisher</td><td>A417-1</td><td></td></tr><tr><td>laboratory wipes</td><td>Fisher</td><td>06-666-11</td><td></td></tr><tr><td>canned air</td><td>Fisher</td><td>18-431</td><td></td></tr><tr><td>10 cm petri dish</td><td>BioPioneer</td><td>GS82-1473-001</td><td></td></tr><tr><td>agarose</td><td>Fisher</td><td>50-444-176</td><td></td></tr><tr><td>razor blade</td><td>Fisher</td><td>12-640</td><td></td></tr><tr><td>forceps</td><td>FST</td><td>11241-40</td><td></td></tr><tr><td>22 x 40 cover glass, #1.5</td><td>Fisher</td><td>50-365-605</td><td></td></tr><tr><td>Fiji (version 1.51d)</td><td>NIH</td><td></td><td>fiji.sc</td></tr><tr><td>Excel 2016</td><td>Microsoft</td><td></td><td>www.microsoftstore.com</td></tr><tr><td>MATLAB R2016</td><td>Mathworks</td><td></td><td>www.mathworks.com</td></tr></tbody></table>

using linear agarose channels to study drosophila larval crawling behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

This article describes a method for guiding Drosophila larval behavior using agarose channels and for measuring the dynamics of larval structures over a crawl cycle. Larvae in linear channels perform sustained bouts of rhythmic crawling (Figure 3). Because both larvae and channels are optically clear, channels can be used with larvae expressing fluorescent probes expressed in any structure of interest. We recorded larvae expressing GFP in all neurons (elav-Ga...

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Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

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Research

JoVE Journal

Neuroscience

7.3K Views.  University of Chicago. The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Video: Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Biology

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Bioengineering

Methods to Assay Drosophila Behavior

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular disease and various neurological diseases1. We have optimized simple and robust behavioral assays for determining larval locomotion, adult climbing ability (RING assay), and courtship behaviors of Drosophila. These behavioral assays are widely applicable for studying the role of genetic and environmental factors on fly behavior. Larval crawling ability can be reliably used for determining early stage changes in the crawling abilities of Drosophila larvae and also for examining effect of drugs or human disease genes (in transgenic flies) on their locomotion. The larval crawling assay becomes more applicable if expression or abolition of a gene causes lethality in pupal or adult stages, as these flies do not survive to adulthood where they otherwise could be assessed. This basic assay can also be used in conjunction with bright light or stress to examine additional behavioral responses in Drosophila larvae. Courtship behavior has been widely used to investigate genetic basis of sexual behavior, and can also be used to examine activity and coordination, as well as learning and memory. Drosophila courtship behavior involves the exchange of various sensory stimuli including visual, auditory, and chemosensory signals between males and females that lead to a complex series of well characterized motor behaviors culminating in successful copulation. Traditional adult climbing assays (negative geotaxis) are tedious, labor intensive, and time consuming, with significant variation between different trials2-4. The rapid iterative negative geotaxis (RING) assay5 has many advantages over more widely employed protocols, providing a reproducible, sensitive, and high throughput approach to quantify adult locomotor and negative geotaxis behaviors. In the RING assay, several genotypes or drug treatments can be tested simultaneously using large number of animals, with the high-throughput approach making it more amenable for screening experiments.

Methods to Assay Drosophila Behavior

Drosophila melanogaster is a genetically and behaviorally tractable model system that has been used to understand the molecular and cellular basis of many important biological processes for over a century 1. Drosophila has been well exploited to gain insights into the genetic basis of fly behavior.

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, Drosophila research lacks popularity in developing countries, possibly due to the misinformed idea that establishing a lab and performing relevant experiments with such tiny insects is difficult and requires expensive, specialized apparatuses. Here, we describe how to build an affordable flylab to quantitatively analyze a myriad of behavioral parameters in D. melanogaster, by 3D-printing many of the necessary pieces of equipment. We provide protocols to build in-house vial racks, courtship arenas, apparatuses for locomotor assays, etc., to be used for general fly maintenance and to perform behavioral experiments using adult flies and larvae. We also provide protocols on how to use more sophisticated systems, such as a high resolution oxygraph, to measure mitochondrial oxygen consumption in larval samples, and show its association with behavioral changes in the larvae upon the xenotopic expression of the mitochondrial alternative oxidase (AOX). AOX increases larval activity and mitochondrial leak respiration, and accelerates development at low temperatures, which is consistent with a thermogenic role for the enzyme. We hope these protocols will inspire researchers, especially from developing countries, to use Drosophila to easily combine behavior and mitochondrial metabolism data, which may lead to information on genes and/or environmental conditions that may also regulate human physiology and disease states.

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

We provide protocols for anyone with a "maker culture" mind to start building a flylab for quantitative analysis of a myriad of behavioral parameters in Drosophila melanogaster, by 3D-printing many of the necessary pieces of equipment. We also describe a high resolution respirometry protocol using larvae to combine behavioral and mitochondrial metabolism data.

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, ...

Behavior

A Simple Technique to Assay Locomotor Activity in Drosophila

Drosophila melanogaster is an ideal model organism for studying various diseases due to its abundance of advanced genetic manipulation techniques and diverse behavioral features. Identifying behavioral deficiency in animal models is a crucial measure of disease severity, for example, in neurodegenerative diseases where patients often experience impairments in motor function. However, with the availability of various systems to track and assess motor deficits in fly models, such as drug-treated or transgenic individuals, an economical and user-friendly system for precise evaluation from multiple angles is still lacking. A method based on the AnimalTracker application programming interface (API) is developed here, which is compatible with the Fiji image processing program, to systematically evaluate the movement activities of both adult and larval individuals from recorded video, thus allowing for the analysis of their tracking behavior. This method requires only a high-definition camera and a computer peripheral hardware integration to record and analyze behavior, making it an affordable and effective approach for screening fly models with transgenic or environmental behavioral deficiencies. Examples of behavioral tests using pharmacologically treated flies are given to show how the techniques can detect behavioral changes in both adult flies and larvae in a highly repeatable manner.

A Simple Technique to Assay Locomotor Activity in Drosophila

The present protocol assesses the locomotor activity of Drosophila by tracking and analyzing the movement of flies in a hand-made arena using open-source software Fiji, compatible with plugins to segment pixels of each frame based on high-definition video recording to calculate parameters of speed, distance, etc.

Drosophila melanogaster is an ideal model organism for studying various diseases due to its abundance of advanced genetic manipulation techniques and ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a ...

Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis

The larvae of Drosophila melanogaster show obvious light-avoiding behavior during the foraging stage. Drosophila larval phototaxis can be used as a model to study animal avoidance behavior. This protocol introduces a light-spot assay to investigate larval phototactic behavior. The experimental set-up includes two main parts: a visual stimulation system that generates the light spot, and an infrared light-based imaging system that records the process of larval light avoidance. This assay allows tracking of the behavior of larva before entering, during encountering, and after leaving the light spot. Details of larval movement including deceleration, pause, head casting, and turning can be captured and analyzed using this method.

Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis

This protocol introduces a light-spot assay to investigate Drosophila larval phototactic behavior. In this assay, a light spot is generated as light stimulation, and the process of larval light avoidance is recorded by an infrared light-based imaging system.

The larvae of Drosophila melanogaster show obvious light-avoiding behavior during the foraging stage. Drosophila larval phototaxis can be used as a model ...

Tracking Freely Walking Flies: A Method to Assess Locomotion in Drosophila

Source: Sun, M. and Chen, L. L. <a href="https://www.jove.com/video/55602/" target="_blank">A Novel Method to Model Chronic Traumatic Encephalopathy in Drosophila</a>. J. Vis. Exp. (2017).
This video describes how to track freely walking flies to assess their general locomotion ability. The featured protocol demonstrates the assay&#39;s setup and data analysis using video tracking software.

Śledzenie swobodnie chodzących much: metoda oceny lokomocji u Drosophila

Source: Sun, M. and Chen, L. L. A Novel Method to Model Chronic Traumatic Encephalopathy in Drosophila. J. Vis. Exp. (2017).
This video describes how to ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

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High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

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Methods to Assay Drosophila Behavior

Functional Analysis of the Larval Feeding Circuit in Drosophila

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

A Simple Technique to Assay Locomotor Activity in Drosophila

Foraging Path-length Protocol for Drosophila melanogaster Larvae

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis

Śledzenie swobodnie chodzących much: metoda oceny lokomocji u Drosophila