This work was supported by startup funds from the University of Nevada, Reno and by NIGMS of the National Institute of Health under grant number P20 GM103650.

1. Building a Behavior Arena and Preparing Hardware to Enable Optogenetic Stimulation in the Behavior Arena
<ol>
	<li>To build a light-deprived behavior arena, construct a box with a dimension of 89 x 61 x 66 cm3 (35&#34; L x 24&#34; W x 26&#34; H) made of black colored plexiglass acrylic sheets (3 mm thick) (see Table of Materials). Materials to build such a box should be available at local hardware stores. Place this box on a table-top in the behavior room (Figure 1A).</li>
	<li>Mount a monochrome USB 3.0 CCD camera fitted with an IR long-pass 830 nm filter and an 8 mm F1.4 C-mount lens (see Table of Materials) to the center of the ceiling of the black box. Place two infra-red LED strips (see Table of Materials) on the table-top in order to illuminate the larvae in the dark arena (Figure 1A).</li>
	<li>To build the LED platform, obtain a 22 cm &#215; 22 cm square Aluminum plate (preferably spray painted with a matte black finish to eliminate any reflections). In the center of the plate, using a metal cutter, cut a hole that is large enough to fit around the CCD camera.</li>
	<li>Cover the metal plate with red LED strip lights (see Table of Materials). Solder LED light strip wires in series and feed the strip wires into an optocoupler relay controlled by a Raspberry Pi 2B microprocessor (see Table of Materials) (Figure 1B and 2).</li>
	<li>Install and configure Ubuntu Mate/Raspian Jesse/Linux based operating system on the Raspberry Pi processor before connecting the optocoupler relay to the LED strips. Attach a power supply to power the LED strips and the optocoupler (Figure 2) (see Table of Materials). Mount the LED platform around the CCD camera (Figure 1B). 
	NOTE: Ubuntu Mate v16.04 operating system is freely available. A set of simple Python based commands can be easily adapted to program patterns of LED light stimuli (see file of syntax).</li>
	<li>Ensure homogenous irradiance at various points in the behavior arena. Measure the absolute irradiance at the surface of the arena with the help of a Spectrometer and determine it to be ~1.3 W/m2 throughout the surface of the arena. 
	NOTE: At this intensity, no significant changes were observed in temperature over the course of the experiments. Another study10 used a higher irradiance of ~1.9 W/m2 and observed no changes in temperature over the course of the experiments.</li>
</ol>
2. Preparation of Drosophila Larvae for Behavior Analyses
<ol>
	<li>Maintain the flies on standard fly food (see Table of Materials) at 25 &#176;C, 50-60% RH, and a 12 h/12 h light/dark cycle.</li>
	<li>In order to express CsChrimson in a single pair of ORNs, cross virgin females from a UAS-IVS-CsChrimson line to males from an OrX-Gal4 line (&#39;X&#39; corresponds to one of 21 larval odor receptor (Or) genes that are uniquely expressed in each of 21 pairs of ORNs)9,11.
	<ol>
		<li>Alternately, in order to express CsChrimson in all 21 larval ORNs, cross virgin females from a UAS-IVS-CsChrimson line to males from an Orco-Gal4 line (&#39;Orco&#39; is the co-receptor that is expressed in all 21 ORNs).</li>
		<li>Use UAS-IVS-CsChrimson line by itself as a control in these experiments. 
		NOTE: The fly stocks listed here are all available at the Bloomington Drosophila Stock Center (see Table of Materials).</li>
	</ol>
	</li>
	<li>Once male and female flies in a cross are allowed to mate and lay eggs for 48 h, transfer adults to a fresh vial.
	<ol>
		<li>To the surface of the food vial containing eggs, add 400 &#181;L of a mixture containing 400 &#181;M all-trans retinal (ATR) dissolved in Dimethyl sulfoxide (DMSO) and 89 mM sucrose dissolved in distilled water. 
		NOTE: The small amount of sucrose promotes larval feeding of the ATR solution. ATR is a cofactor required for upregulating of CsChrimson expression9,10. ATR is light sensitive.</li>
		<li>Once ATR is added to the food vials containing eggs, incubate the vials in the dark for an additional 72 h. 
		NOTE: While this study and the above two studies have not observed effects on larval behavior due to ATR feeding, it is recommended controlling for the effects of ATR feeding by subjecting test lines to the same amount of the above mixture that does not contain ATR.</li>
	</ol>
	</li>
	<li>Extract third-instar larvae (~120 h after egg laying) from the surface of fly food by floating them using a high density (15%) sucrose solution. Using a P1000 micropipette separate the larvae floating on the surface of the sucrose solution into a 1000 mL glass beaker.</li>
	<li>Wash larvae 3&#8211;4 times by exchanging 800 mL fresh distilled water in the glass beaker each time. Allow larvae to rest for 10 min before subjecting them to the behavior assay.</li>
</ol>
3. Behavior Assay
<ol>
	<li>Maintain consistent temperature (between 22&#8211;25 &#176;C) and humidity (between 50 and 60% RH) in the behavior room.</li>
	<li>Prepare larval crawling medium by pouring 150 mL of melted agarose (1.5%) into a 22 cm x 22 cm square Petri dish. One Petri dish is poured for each trial of the assay, 8&#8211;10 trials per experiment. Allow agarose to solidify and cool for 1&#8211;2 h in the Petri dishes before using them in the behavior assay.
	<ol>
		<li>Transfer no more than 20 of the prepped third-instar larvae to the center of Petri dish (Figure 1C). Cover the Petri dish with its lid. Place the Petri dish in the behavior arena under the CCD camera. 
		NOTE: Depending on the experiment, behavior assays are typically carried out for 3&#8211;5 min. If an odorant is used in the assay to provide guidance cues, it has been observed that the resulting odor gradient remains stable for approximately 5 min1. Longer assay times are not recommended. Deleterious effects on larvae due to dehydration or from prolonged 630 nm red-light exposure have not been observed within these time points.</li>
	</ol>
	</li>
	<li>Turn ON the 850 nm infra-red LED light source to visualize larvae in the video. Start the CCD camera to record larval movement.</li>
	<li>Using the software associated with the Raspberry Pi processor, program the procedure to administer appropriate patterns of red light stimulation. 
	NOTE: A set of simple Python based commands can be easily adapted to program patterns of LED light stimuli (see file of syntax).</li>
</ol>
4. Data Processing and Analysis
<ol>
	<li>Import the recorded video of each trial into any available programming software like Matlab.</li>
	<li>Obtain XY coordinates of every larva in a movie as a function of time. Based on limits of the tracking software, 15&#8211;20 third-instar larvae can be tracked in a single movie1,8. 
	NOTE: A set of simple Matlab codes (&#39;Tracklarva&#39;) that can be easily adapted to suit appropriate conditions (see file of syntax) is provided. This program combines all trials in an experiment and outputs the XY coordinates of every larva for the entire duration of the assay (see code syntax below). Alternately, one could use several open-source based programs that are freely available for researchers. E.g. JAABA (http://jaaba.sourceforge.net/)12.</li>
	<li>Use the generated XY coordinates to plot larval trajectories and to further analyze larval locomotion.
	<ol>
		<li>For behavior analyses, use navigational statistics such as speed, path curvature, heading angle, to segment individual larval trajectories into alternating sequences of runs and turns.</li>
		<li>Runs are defined as continuous periods of forward movement. Turns separate successive runs. Turns are flagged when changes to trajectory orientation angles were &#62;45&#176; (see file of syntax).</li>
	</ol>
	</li>
	<li>Calculate average values for run speed, run length, run direction, and other parameters as required. 
	NOTE: A set of simple syntax to extract &#39;runs&#39; and &#39;stops&#39; from larval trajectories is provided (see file of syntax). Simple Matlab or Excel based functions can be applied to the extracted data to calculate values for &#39;run speed&#39;, &#39;run length&#39; etc.</li>
	<li>Represent data for each behavioral metric as mean &#177; SEM.</li>
</ol>

<ol>
	<li>Mathew, D., 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=Functional+diversity+among+sensory+receptors+in+a+Drosophila+olfactory+circuit.">Functional diversity among sensory receptors in a Drosophila olfactory circuit.</a> Proceedings of the National Academy of Sciences of the United States of America. 110, 2134-2143 (2013).</li><li>Ramaekers, A., 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=Glomerular+maps+without+cellular+redundancy+at+successive+levels+of+the+Drosophila+larval+olfactory+circuit.">Glomerular maps without cellular redundancy at successive levels of the Drosophila larval olfactory circuit.</a> Current biology : CB. 15, 982-992 (2005).</li><li>Couto, A., Alenius, M., Dickson, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular,+anatomical,+and+functional+organization+of+the+Drosophila+olfactory+system.">Molecular, anatomical, and functional organization of the Drosophila olfactory system.</a> Current biology : CB. 15, 1535-1547 (2005).</li><li>Kreher, S. A., Kwon, J. Y., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+basis+of+odor+coding+in+the+Drosophila+larva.">The molecular basis of odor coding in the Drosophila larva.</a> Neuron. 46, 445-456 (2005).</li><li>Kreher, S. A., Mathew, D., Kim, J., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translation+of+sensory+input+into+behavioral+output+via+an+olfactory+system.">Translation of sensory input into behavioral output via an olfactory system.</a> Neuron. 59, 110-124 (2008).</li><li>Hallem, E. A., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coding+of+odors+by+a+receptor+repertoire.">Coding of odors by a receptor repertoire.</a> Cell. 125, 143-160 (2006).</li><li>Monte, P., 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=Characterization+of+the+larval+olfactory+response+in+Drosophila+and+its+genetic+basis.">Characterization of the larval olfactory response in Drosophila and its genetic basis.</a> Behav Genet. 19, 267-283 (1989).</li><li>Gershow, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controlling+airborne+cues+to+study+small+animal+navigation.">Controlling airborne cues to study small animal navigation.</a> Nature methods. 9, 290-296 (2012).</li><li>Klapoetke, N. C., 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=Independent+optical+excitation+of+distinct+neural+populations.">Independent optical excitation of distinct neural populations.</a> Nature methods. 11, 338-346 (2014).</li><li>Hernandez-Nunez, L., 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=Reverse-correlation+analysis+of+navigation+dynamics+in+Drosophila+larva+using+optogenetics.">Reverse-correlation analysis of navigation dynamics in Drosophila larva using optogenetics.</a> eLife. 4, (2015).</li><li>Brand, A. H., Perrimon, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+gene+expression+as+a+means+of+altering+cell+fates+and+generating+dominant+phenotypes.">Targeted gene expression as a means of altering cell fates and generating dominant phenotypes.</a> Development. 118, 401-415 (1993).</li><li>Kabra, M., Robie, A. A., Rivera-Alba, M., Branson, S., Branson, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=JAABA:+interactive+machine+learning+for+automatic+annotation+of+animal+behavior.">JAABA: interactive machine learning for automatic annotation of animal behavior.</a> Nature methods. 10, 64-67 (2013).</li><li>Newquist, G., Novenschi, A., Kohler, D., Mathew, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+contributions+of+Olfactory+Receptor+Neurons+in+a+Drosophila+olfactory+circuit.">Differential contributions of Olfactory Receptor Neurons in a Drosophila olfactory circuit.</a> eNeuro. 3, (2016).</li><li>Schulze, A., 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=Dynamical+feature+extraction+at+the+sensory+periphery+guides+chemotaxis.">Dynamical feature extraction at the sensory periphery guides chemotaxis.</a> eLife. 4, (2015).</li><li>Tastekin, I., 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=Role+of+the+Subesophageal+Zone+in+Sensorimotor+Control+of+Orientation+in+Drosophila+Larva.">Role of the Subesophageal Zone in Sensorimotor Control of Orientation in Drosophila Larva.</a> Current Biology. 25, 1448-1460 (2015).</li><li>Famiglietti, E. V., Kolb, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+basis+for+ON-and+OFF-center+responses+in+retinal+ganglion+cells.">Structural basis for ON-and OFF-center responses in retinal ganglion cells.</a> Science. 194, 193-195 (1976).</li><li>Luo, L., 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=Bidirectional+thermotaxis+in+Caenorhabditis+elegans+is+mediated+by+distinct+sensorimotor+strategies+driven+by+the+AFD+thermosensory+neurons.">Bidirectional thermotaxis in Caenorhabditis elegans is mediated by distinct sensorimotor strategies driven by the AFD thermosensory neurons.</a> Proceedings of the National Academy of Sciences of the United States of America. 111, 2776-2781 (2014).</li><li>Berck, M. 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The authors have nothing to disclose.

Here, we described a method that allows for the measurement of Drosophila larval behavior in response to simultaneous optogenetic activation of olfactory neurons. Previously described larval tracking methods1,8 use different odor delivery technology to activate ORNs. However, these methods cannot control for either the specificity or temporal patterns of ORN activation. Our method overcomes these deficits by using light stimuli instead of odor stimuli for more precise control of ORN activation.
The materials needed to build the behavior arena can be easily obtained at the local hardware store and requires minimal effort at assembly. The electronics needed to prepare the optogenetics module are also easily available and constructed. The method described here uses red light to activate a red-shifted channel rhodopsin (CsChrimson) expressed in specific neurons. The resulting larval behavior in response to the corresponding ORN activation is recorded using a CCD camera and measured using custom written software that is provided here. Our method allows researchers to ask answers to several questions that were not possible before: 1) what is the impact of different olfactory stimulus patterns on the animal's ability to navigate toward an odor? 2) similar to ON-center and OFF-center ganglion cells in mammalian retina16, are there ORNs that respond specifically to a decrease in olfactory stimuli in addition to ORNs that respond to an increase in olfactory stimuli? Finally, our method permits a variety of future applications, including measuring the impact of downstream neurons in the olfactory circuit (PNs and LNs) to larval navigation.
While there are several advantages to our method, we acknowledge certain limitations. It is unclear whether concentration effects observed with odorants can be easily replicated using this system. While our present setup does not allow this, the optogenetics module could be easily modified to accommodate increases or decreases in the intensity of the light stimulus. In the future, we will check to see whether changing intensity of light stimulus mimics concentration effects of odorants. Simple fly genetic techniques can be used to express CsChrimson in either all 21 pairs of ORNs (using Orco-Gal4) or in a single pair of ORNs (using individual Or-Gal4s). However, complicated genetics would be required to express CsChrimson in '1 &lt; n &lt; 21' neurons. Due to this, it would be difficult to replicate the effects observed with odor mixtures where individual components of the mixture elicit responses from more than one ORN. Even though larval navigational behavior is considered to be a low dimensional behavior, we acknowledge that our larval tracking program could be further improved in the future by considering additional behavioral descriptors based on animal posture (e.g. probability of head turns, body bends etc.)8,17. Our study was restricted to first order sensory neurons in the larva. Further investigation is required before our method can be applied to second order projection neurons and local neurons that are embedded in the brain lobe region of the brain18.
In summary, our method offers the ability to dissect the function of every ORN in the simple, tractable olfactory circuit of the Drosophila larva. By doing so, our method will enable development of more precise computational models that describe how odor signals are translated into different behavioral outputs.

Olfactory information in a Drosophila larva&#39;s environment is sensed by only 21 functionally distinct ORNs, the activities of which ultimately determine larval behavior1,2,3,4. Yet, relatively little is known about the logic by which sensory information is encoded in the activities of these 21 ORNs. There is thus a need to experimentally measure the functional contributions of each larval ORN to behavior.
Although the sensory response profile of the entire repertoire of Drosophila larval ORNs has been studied in detail1,4,5, the contributions of individual ORNs to the olfactory circuit and thereby to navigational behavior remain largely unknown. Difficulties in larval behavior studies, so far, arise due to the inability to spatially and temporally activate single ORNs. A panel of odorants that specifically activate 19 of the 21 Drosophila larval ORNs was recently described1. Each odorant in the panel, at low concentrations, elicits a physiological response only from its cognate ORN. However, at higher concentrations that are normally used for conventional behavior assays, each odorant elicits physiological responses from multiple ORNs1,5,6. Further, odorants in this panel have varied volatilities that complicate interpretation of behavior studies that depend on formation of stable odor gradients7,8. Finally, naturally occurring odor stimuli have a temporal component that is difficult to replicate under laboratory conditions. It is therefore important to develop a method that can measure larval behavior while simultaneously activating individual ORNs in a spatial and temporal manner.
Here, we demonstrate a method that has advantages over previously described larval tracking assays1,8. The tracking assay described in Gershow et al. uses electronically controlled valves to maintain a stable gradient of odor in the behavior arena8. However, due to the level of complex engineering involved to build the odor stimulus setup, this method is difficult to replicate in other laboratories. Further, the issues related to using odorants to specifically activate single ORNs remain unresolved. The tracking assay described in Mathew et al. uses a simpler odor delivery system, but the resulting odor gradient is dependent on the volatility of test odorant and is unstable for long durations of the assay1. Thus, by replacing odor stimuli with light stimuli, our method has the advantages of specificity and precise temporal control of ORN activation and is not dependent on formation of odor gradients of different strengths.
Our method is easy to set up and is appropriate for researchers interested in measuring aspects of Drosophila larval navigation. This technique could be adapted to other model systems provided that the researcher is able to drive the expression of CsChrimson in their favorite system&#39;s neuron(s) of choice. CsChrimson is a red-shifted version of channel rhodopsin. It is activated at wavelengths that are invisible to the larva&#39;s phototaxis system. We are therefore able to manipulate the activity of neurons with specificity, reliability, and reproducibility9. By modifying the custom written software to account for size changes of the subjects, this method could easily be adapted for crawling larvae of other insect species.

<table border="0" cellpadding="0" cellspacing="0" style="width:925px;" width="924">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
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		<tr height="40">
			<td height="40" style="height:40px;width:359px;">Video camera to capture larval movement</td>
			<td style="width:296px;"></td>
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			<td style="width:63px;"></td>
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			<td height="33" style="height:33px;">CCD Camera&#160;</td>
			<td style="width:296px;">Edmund Optics</td>
			<td style="width:208px;">106215</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">M52 to M55 Filter Thread Adapter</td>
			<td style="width:296px;">Edmund Optics</td>
			<td>59-446</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">2&#34; Square Threaded Filter Holder for Imaging Lenses&#160;</td>
			<td style="width:296px;">Edmund Optics</td>
			<td>59-445</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">RG-715, 2&#34; Sq. Longpass Filter</td>
			<td style="width:296px;">Edmund Optics</td>
			<td>46-066</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Electronics for optogenetic setup</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr height="33">
			<td height="33" style="height:33px;">Raspberry Pi 2B</td>
			<td>RASPBERRY-PI.org</td>
			<td>RPI2-MODB-V1.2</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">3 Channel programmable power supply</td>
			<td>newegg.com</td>
			<td>9SIA3C62037092</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">8 Channel optocoupler relay</td>
			<td>amazon.com</td>
			<td>6454319</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">630nm Quad-row LED strip lights</td>
			<td>environmentallights.com</td>
			<td>red3528-450-reel</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:359px;">850nm LED strips</td>
			<td>environmentallights.com</td>
			<td>wp-4000K-CC5050-60x2-kit</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:359px;">Software&#160;</td>
			<td style="width:296px;"></td>
			<td style="width:208px;"></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:359px;">Matlab</td>
			<td>Mathworks Inc.</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:359px;">Ubuntu MATE v16.04</td>
			<td>Nubuntu</td>
			<td>https://github.com/yslo/nubuntu</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Other items</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Plexiglass black acrylic</td>
			<td>Home Depot</td>
			<td style="width:208px;">MC1184848bl</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Fly food and other reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Nutrifly fly food</td>
			<td style="width:296px;">Genesee Scientific</td>
			<td>66-112</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Agarose powder</td>
			<td style="width:296px;">Genesee Scientific</td>
			<td>20-102</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">22cm X 22cm square petri-dish</td>
			<td style="width:296px;">VWR Inc.</td>
			<td>25382-327</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">DMSO</td>
			<td>Sigma-Aldrich</td>
			<td>D2650</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>84097</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">All trans-retinal</td>
			<td>Sigma-Aldrich</td>
			<td>R2500</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Flies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">UAS-IVS-CsChrimson&#160;</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>55134</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:359px;">Orco-Gal4</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>26818</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Or42a-Gal4</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>9970</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Or7a-Gal4</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>23907</td>
			<td></td>
		</tr>
	</tbody>
</table>

tracking drosophila larval behavior in response to optogenetic stimulation of olfactory neurons

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

To demonstrate the specificity of ORN activation, our method was successfully applied to determine the impact of two different ORN (ORN::7a &#38; ORN::42a) (ORNs expressing either Or7a or Or42a) activation on larval behavior (Figure 3). Consistent with recent studies that individual larval ORNs are functionally distinct1,10,13, our representative data demonstrates that when ORN::7a expressing CsChrimson was stimulated by light, there was a significant decrease in run length compared to control animals. Conversely, when ORN::42a expressing CsChrimson was stimulated by light, there was a significant increase in run length compared to control animals (Figure 3). The collective data analyzed from ~100-120 larval tracks were obtained from (n = 8) trials performed for each genotype. The error bars represent SEM. While we describe only a single behavior parameter (run length) here, we note that each larval track can be further analyzed to calculate parameters for speed, path curvature, and body bends1,8,13,14,15. More parameters related to directionality such as heading angle, run length and run speed toward and away from odors can be obtained if an odor source is provided on one side of the arena1,8,13.
To demonstrate our method&#39;s ability to alter the temporal patterns of ORN activation, we varied our stimulus to alternate between lights OFF and ON. We subjected larvae expressing UAS-CsChrimson in ORN::42a to three different temporal patterns of light stimuli during the lights ON period (0.04 Hz, 1 Hz, and Constant). We then measured changes in behavioral parameters that happen during lights OFF &#8594; ON phase and during lights On &#8594; OFF phase. We found that for ORN::42a, different temporal patterns of light stimulation elicited different behavioral responses (Figure 4). Such changes were not observed in control larvae that do not express UAS-CsChrimson in any ORNs. These results highlight the importance of understanding how temporal patterns of ORN activation contribute to animal behavior.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57353/57353fig1.jpg" /> 
Figure 1: Behavior arena and larval crawling medium. (A) Front view of the black-box behavior arena. The open door of the arena reveals a CCD camera suspended from the ceiling of the box. (B) Bottom view of a metal platform containing red LED light strips used for optogenetic stimulations is mounted around the CCD camera. (C) Top view of the larval crawling medium used in the assay. Prior to start of recording larval movement, ~20 washed larvae are laid along the center of a 22 cm x 22 cm Petri dish layered with 1.5% agarose. The Petri dish containing larvae is placed in the center of the arena under the CCD camera and in between two infra-red LED light strips that are used as a light source for the camera. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57353/57353fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57353/57353fig2.jpg" /> 
Figure 2: Optogenetics setup. An infographic showing the electronic arrangement for the optogenetics setup. Briefly, LED light (630 nm) strips are connected in series and wires from the strips are fed into an optocoupler connected to a raspberry PI 2B microprocessor. Both the LED light strips and the optocoupler are powered by a power supply. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57353/57353fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57353/57353fig3.jpg" /> 
Figure 3: Impact of light activation of individual ORNs on larval behavior. Run length of larvae expressing CsChrimson in ORN::7a and ORN::42a were differently affected compared to control larvae upon light activation. Each bar represents average RI &#177; SEM (n = 8). Run length of larvae when ORN::7a was activated was significantly lower than control. Run length of larvae when ORN::42a was activated was significantly higher than control. Bars represent mean &#177; SEM (n = 8, Student t-test; &#34;*&#34; is p &#60;0.05, &#34;**&#34; is p &#60;0.001). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57353/57353fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57353/57353fig4.jpg" /> 
Figure 4: Impact of different temporal patterns of ORN activation on larval behavior. (A) Three temporal patterns of stimuli used for activating ORNs. Stimulus a: 1 min constant light during, Stimulus b: 0.04 Hz light stimulation, Stimulus c: 1 Hz light stimulation during LED ON period in minute 2. (B) Larvae were subjected to the three different patterns of light stimuli described in A. Each dot represents the change in larval behavior, under each pattern of light stimuli, when light activation is switched from ON to OFF. Change in &#39;run length&#39; (Av. run length (OFF) - Av. run length (ON)) is plotted on the X-axis. Change in &#39;run speed&#39; (Av. run speed (OFF) - Av. run speed (ON)) is plotted on the Y-axis. The left graph (grey dots) represents measurements from control larvae and the right graph (red dots) represents measurements from larvae expressing CsChrimson in ORN::42a. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57353/57353fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Final Code PDF" src="/files/ftp_upload/57353/57353code.jpg" /> 
Supplemental file of syntax: A set of simple Matlab codes (&#39;Tracklarva&#39;) that can be easily adapted to suit appropriate conditions. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57353/Final_Code.pdf" target="_blank">Please click here to download this file.</a>

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Śledzenie zachowania larw Drosophila w odpowiedzi na optogenetyczną stymulację neuronów węchowych | Neuronauka | Wideo

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

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

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

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

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7.2K Views.  University of Nevada. This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.

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

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 ...

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 ...

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

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 ...

Appetitive Associative Olfactory Learning in Drosophila Larvae

In the following we describe the methodological details of appetitive associative olfactory learning in Drosophila larvae. The setup, in combination with genetic interference, provides a handle to analyze the neuronal and molecular fundamentals of specifically associative learning in a simple larval brain.
Organisms can use past experience to adjust present behavior. Such acquisition of behavioral potential can be defined as learning, and the physical bases of these potentials as memory traces1-4. Neuroscientists try to understand how these processes are organized in terms of molecular and neuronal changes in the brain by using a variety of methods in model organisms ranging from insects to vertebrates5,6. For such endeavors it is helpful to use model systems that are simple and experimentally accessible. The Drosophila larva has turned out to satisfy these demands based on the availability of robust behavioral assays, the existence of a variety of transgenic techniques and the elementary organization of the nervous system comprising only about 10,000 neurons (albeit with some concessions: cognitive limitations, few behavioral options, and richness of experience questionable)7-10.
Drosophila larvae can form associations between odors and appetitive gustatory reinforcement like sugar11-14. In a standard assay, established in the lab of B. Gerber, animals receive a two-odor reciprocal training: A first group of larvae is exposed to an odor A together with a gustatory reinforcer (sugar reward) and is subsequently exposed to an odor B without reinforcement 9. Meanwhile a second group of larvae receives reciprocal training while experiencing odor A without reinforcement and subsequently being exposed to odor B with reinforcement (sugar reward). In the following both groups are tested for their preference between the two odors. Relatively higher preferences for the rewarded odor reflect associative learning - presented as a performance index (PI). The conclusion regarding the associative nature of the performance index is compelling, because apart from the contingency between odors and tastants, other parameters, such as odor and reward exposure, passage of time and handling do not differ between the two groups9.

Appetitive Associative Olfactory Learning in Drosophila Larvae

Drosophila larvae are able to associate odor stimuli with gustatory reward. Here we describe a simple behavioral paradigm that allows the analysis of appetitive associative olfactory learning.

In the following we describe the methodological  details of appetitive associative olfactory learning in Drosophila larvae. The setup, in combination with ...

Drosophila Adult Olfactory Shock Learning

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the elucidation of the molecular mechanisms involved in cognitive diseases1-7. Learning and memory can be assayed in larvae to study the effect of neurodevelopmental genes8-10 and in flies to measure the contribution of adult plasticity genes1-7. Furthermore, the short lifespan of Drosophila facilitates the analysis of genes mediating age-related memory impairment5,11-13. The availability of many inducible promoters that subdivide the Drosophila nervous system makes it possible to determine when and where a gene of interest is required for normal memory as well as relay of different aspects of the reinforcement signal3,4,14,16.
Studying memory in adult Drosophila allows for a detailed analysis of the behavior and circuitry involved and a measurement of long-term memory15-17. The length of the adult stage accommodates longer-term genetic, behavioral, dietary and pharmacological manipulations of memory, in addition to determining the effect of aging and neurodegenerative disease on memory3-6,11-13,15-21.
Classical conditioning is induced by the simultaneous presentation of a neutral odor cue (conditioned stimulus, CS+) and a reinforcement stimulus, e.g., an electric shock or sucrose, (unconditioned stimulus, US), that become associated with one another by the animal1,16. A second conditioned stimulus (CS-) is subsequently presented without the US. During the testing phase, Drosophila are simultaneously presented with CS+ and CS- odors. After the Drosophila are provided time to choose between the odors, the distribution of the animals is recorded. This procedure allows associative aversive or appetitive conditioning to be reliably measured without a bias introduced by the innate preference for either of the conditioned stimuli. Various control experiments are also performed to test whether all genotypes respond normally to odor and reinforcement alone.

Drosophila Adult Olfactory Shock Learning

The method to measure adult Drosophila associative memory is described. The assay is based on the ability of the fly to associate an odor presented with a negative reinforcer (electric shock) and then recall this information at a later time, allowing memory to be measured.

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...