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, check for obstacles like an air bubble in the channel, and check the health of the larva. Another important element of a successful recording is that the larva is oriented optimally to visualize structures of interest. If the larva is not oriented correctly, or if the larva crawls out of the channel, simply remove the coverslip and remount the larva. In our experience, ~20% of larvae initially mounted yield excellent behavioral recordings without adjustment.
In the past, measurements were taken of the position of larval structures such as the mouth hook, gut, and abdominal segments during crawling behavior. To visualize the movement of these structures over the crawling stride cycle, polar coordinate plots were generated. In this paper, the fluorescence intensity of the nerve cord was measured and polar coordinate plots used to visualize the dynamics of fluorescence over the crawling stride cycle (Figure 4). There are several advantages to representing the data on polar coordinate plots: it eliminates crawl speed as a variable, it can summarize data from many animals and many strides, and it allows visualization of both overall trends and variation in data11. Notably, it is possible to measure the dynamics of any fluorescently-labeled structure of interest. In principle, this analysis is applicable to tracking any type of dynamic changes that occurs over a crawl cycle.
There is a wide array of applications for the methods described in this paper. In the past, linear agarose channels have been used to record larval behavior at the whole organism, segment and individual muscle levels1. These data showed that larvae use a &#34;visceral-pistoning&#34; mechanism for both forward and reverse crawling, and they allowed the neuromuscular mechanism driving both forward and reverse crawling to be determined1. In the future, researchers can use channels to study crawling in different genetic backgrounds. In addition, it should be possible to use channels to analyze the activity of larval neurons using calcium imaging during crawling. This should lead to the understanding of which neurons fire in phase with particular movements of the crawl cycle. Finally, there is no reason that channels must follow the linear design presented in this paper; using channels with different dimensions will no doubt help answer a variety of question about Drosophila larval locomotion and motor control as a whole.

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-Gal4/+; UAS-myr-GFP/+) and monitored the dynamic changes in fluorescence intensity in the nerve cord over the crawl cycle. We show that the CNS moves forward at nearly the same time as the larval head and tail (Figure 4A-B). As a wave of muscle contraction passes along the body axis, the CNS moves in and out of the plane of focus causing the fluorescence of the nerve cord to change (Figure 4). To quantify changes in nerve cord fluorescence intensity for several strides in several animals we represented the data on a polar coordinate plot (Figure 4C). Plotting the data on polar coordinate plots shows that the dynamics of the nerve cord fluorescence over the stride cycle follows a reproducible pattern.
<img alt="Figure 1" src="/files/ftp_upload/54892/54892fig1.jpg" /> 
Figure 1: Design of Linear Channels to Guide Drosophila Larval Crawling Behavior. (A) The design of the microfluidic device used to make linear agarose channels is shown. The widths of channels in this device vary from 100-300 &#181;m by increments of 50 &#181;m. The depth is 150 &#181;m. (B) A Drosophila larva is loaded into an agarose channel. A dorsal view is shown with anterior (head) to the right. Scale bar = 200 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54892/54892fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54892/54892fig2.jpg" /> 
Figure 2: Diagram Illustrating How to Load a Larva into a Channel. See protocol section 3 for details <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54892/54892fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54892/54892fig3.jpg" /> 
Figure 3: A Fluorescently-labeled Drosophila&#160;Larva Performs Sustained, Rhythmic, Linear Crawling when Placed into an Agarose Channel. At left, a schematic of a larva expressing GFP in all neurons (elav&#62;GFP) is shown. A box shows the region where fluorescence intensity of the nerve cord (distinguished by its elongated morphology) can be measured. At right is an example of a larva crawling through a channel at one second intervals. A dorsal view is shown with anterior up. Arrows indicate the initiation of a stride. Scale bar = 200 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54892/54892fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54892/54892fig4.jpg" /> 
Figure 4: Dynamics of Fluorescence Changes in Nerve Cord Over the Crawl Cycle are Presented on Polar Coordinate Plots. (A) A diagram of a single stride. Percentages refer to percent of stride cycle completed. By convention forward movement of the tail, head, and internal organs such as the CNS marks the initiation of a stride (or 0% of stride cycle). Note that the CNS (white) moves forward and backward, as well as up and down. A side view is shown with anterior to the right. (B) A kymograph shows the movement of the head, tail, and CNS. Note that the fluorescence intensity of the nerve cord during the stride cycle is dynamic. (C) A polar coordinate plot shows the dynamic changes in normalized fluorescence intensity of the nerve cord over the stride cycle. Each dot represents a normalized fluorescence intensity of a single larva at a single time point (n = 3 larvae, 3 strides each). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54892/54892fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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 ...

using-linear-agarose-channels-to-study-drosophila-larval-crawling

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JoVE Journal

Neuroscience

7.4K Views.  University of Chicago.  De Drosophila larven is een krachtig modelsysteem om neuronale controle van gedrag te bestuderen. Deze publicatie beschrijft het gebruik van lineaire agarose kanalen om aanhoudende aanvallen van lineaire kruipen en methoden te lokken om de dynamiek van larvale structuren te kwantificeren tijdens repetitieve kruipen gedrag. 

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

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen distribution. It is also possible that aberrant behavior of oxygen-sensing neurons could induce hypoxia-like behavior in the presence of normal oxygen levels. In D. melanogaster, development at low oxygen levels results in inhibition of growth and sluggish behavior during the larval phases. However, these established manifestations of oxygen deficit overlap considerably with the phenotypes of many mutations that regulate growth, stress responses or locomotion. As result, there is currently no assay available to identify i) cellular hypoxia induced by a mutation or ii) hypoxia-like behavior when induced by abnormal neuronal behavior.
We have recently identified two distinctive behaviors in D. melanogaster larvae that occur at normal oxygen levels in response to internal detection of hypoxia. First, at all stages, such larvae avoid burrowing into food, often straying far away from a food source. Second, tunneling into a soft substratum, which normally occurs during the wandering third instar stage is completely abolished if larvae are hypoxic. The assay described here is designed to detect and quantitate these behaviors and thus to provide a way to detect hypoxia induced by internal damage rather than low external oxygen. Assay plates with an agar substratum and a central plug of yeast paste are used to support animals through larval life. The positions and state of the larvae are tracked daily as they proceed from first to third instar. The extent of tunneling into the agar substratum during wandering phase is quantitated after pupation using NIH ImageJ. The assay will be of value in determining when hypoxia is a component of a mutant phenotype and thus provide insight into possible sites of action of the gene in question.

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

The protocol describes a simple assay to identify Drosophila melanogaster larvae that are experiencing hypoxia under normal atmospheric oxygen levels. This protocol allows hypoxic larvae to be distinguished from other mutants that show overlapping phenotypes such as sluggishness or slow growth.

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen ...

Behavior

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, ...

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 ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...

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.remotexs.ntu.edu.sg/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.

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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