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 border="0" cellpadding="0" cellspacing="0" width="643">
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		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">6 oz square Drosophila bottle</td>
			<td style="width:101px;">Scimart</td>
			<td style="width:119px;">DR-103</td>
			<td style="width:212px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">agar</td>
			<td style="width:101px;">sigma</td>
			<td style="width:119px;">A1296</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">sucrose</td>
			<td style="width:101px;">sigma</td>
			<td style="width:119px;">S9378</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">apple juice</td>
			<td></td>
			<td></td>
			<td>not from concentrate</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">Tegosept</td>
			<td>Fisher</td>
			<td>T2300</td>
			<td>methyl-p-hydroxybenzoate</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">35 x 10 mm round petri dish</td>
			<td style="width:101px;">Fisher</td>
			<td align="right">351008</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">baker&#39;s yeast</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">PDMS casting mold</td>
			<td style="width:101px;">FlowJem</td>
			<td style="width:119px;"></td>
			<td>can be requested from authors</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">isopropyl alcohol</td>
			<td style="width:101px;">Fisher</td>
			<td style="width:119px;">A417-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">laboratory wipes</td>
			<td style="width:101px;">Fisher</td>
			<td style="width:119px;">06-666-11</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">canned air</td>
			<td style="width:101px;">Fisher</td>
			<td style="width:119px;">18-431</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">10 cm petri dish</td>
			<td style="width:101px;">BioPioneer</td>
			<td style="width:119px;">GS82-1473-001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">agarose</td>
			<td style="width:101px;">Fisher</td>
			<td style="width:119px;">50-444-176</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">razor blade</td>
			<td style="width:101px;">Fisher</td>
			<td style="width:119px;">12-640</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">forceps</td>
			<td style="width:101px;">FST</td>
			<td style="width:119px;">11241-40</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">22 x 40 cover glass, #1.5</td>
			<td style="width:101px;">Fisher</td>
			<td style="width:119px;">50-365-605</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">Fiji (version 1.51d)</td>
			<td style="width:101px;">NIH</td>
			<td style="width:119px;"></td>
			<td>fiji.sc</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">Excel 2016</td>
			<td style="width:101px;">Microsoft</td>
			<td></td>
			<td>www.microsoftstore.com</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:211px;">MATLAB R2016</td>
			<td style="width:101px;">Mathworks</td>
			<td style="width:119px;"></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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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

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

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 ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

A Single-fly Assay for Foraging Behavior in Drosophila

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster. In a light-tight box illuminated by red light that is invisible to fruit flies, a camera linked to custom data acquisition software monitors the position of six flies simultaneously. Each fly is confined to walk in individual arenas containing a food odor at the center. The testing arenas rest on a porous floor that functions to prevent odor accumulation. Latency to locate the odor source, a metric that reflects olfactory sensitivity under different physiological states, is determined by software analysis. Here, we discuss the critical mechanics of running this behavioral paradigm and cover specific issues regarding fly loading, odor contamination, assay temperature, data quality, and statistical analysis.

A Single-fly Assay for Foraging Behavior in Drosophila

In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster.

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Assays to Detect UV-reflecting Structures and Determine their Importance in Mate Preference using the Sailfin Molly Poecilia latipinna

Many organisms use cues and signals beyond human sensitivity during social interactions. It is important to take into account how organisms perceive their worlds when trying to understand their behavior and ecology. Sensitivity to the ultraviolet spectrum (UV; 300 - 400 nm) is found across multiple genera of birds, fish, reptiles, amphibians, and even mammals. This protocol describes a technique for examining organisms for the presence of UV-reflecting structures and a method for testing whether these cues are used as social signals in the context of mate choice. A spectrophotometer is used to detect the presence of UV reflectance and variation in reflective intensity between individuals and sexes. An example of this technique is presented in which a dichotomous mate choice test exposes sexually receptive individuals to opposite sex individuals whose visual appearance can be manipulated by filters that either transmit full spectrum or block UV wavelengths. This system allowed for the determination that female, but not male, sailfin mollies (Poecilia latipinna) were using UV markings as part of their mating decisions. These types of studies serve to expand our knowledge of the range of organisms that utilize UV and provide insight into how UV plays a role in their lives.

Assays to Detect UV-reflecting Structures and Determine their Importance in Mate Preference using the Sailfin Molly Poecilia latipinna

This protocol outlines the use of spectrophotometry to detect ultraviolet-reflecting structures on organisms (in this example, the sailfin molly Poecilia latipinna) and describes dichotomous choice tests for fish that allow inferences to be made on the role of ultraviolet cues during mate selection.

Many organisms use cues and signals beyond human sensitivity during social interactions. It is important to take into account how organisms perceive their ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...