We thank Ye Zhang for the help with the sample traces and Tin Ki Tsang for the help with the pictures. This work was supported by a Ray Thomas Edwards Foundation Early Career Award and an NIH grant (R01DC015519) to C.-Y.S. and NIH grants (R01DC009597 and R01DK092640) to J.W.W.

1. Preparation of the Hardware for at4 Recording
<ol>
	<li>Use a pipette puller instrument to prepare electrodes with aluminosilicate glass capillaries (O.D 1.0 mm, I.D. 0.64 mm). Blunt the tip of the reference electrode slightly with a pair of fine forceps to facilitate insertion into the clypeus of the fly (i.e.&#160;a rounded plate at the front of the fly head, above the mouthparts). 
	NOTE: 7-day-old WT males (Berlin) were used in this study. Use AHL saline solution14 as the electrolyte for both electrodes.</li>
	<li>Prepare 1 L of AHL by mixing 900 mL of distilled water with 6.312 g of NaCl, 0.373 g of KCl, 0.337 g of NaHCO3, 0.1120 g of NaH2PO4, 1.892 g of Trehalose &#1468;2H2O, 3.423 g of sucrose, 1.192 g of HEPES, and 8.2 mL of 1M MgCl2. Using distilled water, bring the total volume up to 1 L. Bring the pH to 7.4 using 1 N NaOH and sterilize the solution with a vacuum-driven filter system. For long-term storage, keep the AHL aliquots at 4 &#176;C. 
	NOTE: Consistent deliveries of palmitoleic acid are contingent upon uniformity between cartridges. It is critical that each cartridge is assembled in a reproducible manner.</li>
	<li>Using a razor blade, remove 0.9 cm from the tip of a 200 &#181;L pipette tip to create the first cartridge section, measuring 4.1 cm; refer to the dimensions detailed in Figure 1A. Use another 200 &#181;L pipette tip and remove 1.7 cm and 1.5 cm from the tip and the base, respectively, to create the second cartridge section, measuring 1.8 cm (Figure 1A). Use a ruler to ensure reproducibility.</li>
	<li>Use a &#8539;&#34; hole puncher to cut discs from filter paper.</li>
	<li>Use forceps to place a filter paper disc at the tip of the second cartridge section. Visually confirm that there is an opening in the cartridge tip through which air can pass.</li>
	<li>Attach the first and second cartridge sections together, as shown in Figure 1A. Angle the second cartridge section downward to facilitate square aiming at the prep (Figure 1B).</li>
	<li>Connect the cartridge with the odor-delivery tube, which is mounted on a micromanipulator. 
	NOTE: This design allows the cartridge to be swiveled outward to facilitate exchange (Figure 1C).</li>
	<li>Set the constant humidified airflow to 2 L/min in one mass controller and the odorant flow to 500 mL/min in another mass controller.</li>
	<li>Using the software (see the Table of Materials), program the procedure to administer a 500 ms odor puff.</li>
</ol>
2. Preparation of Palmitoleic Acid Odorant Solutions for Delivery
NOTE: Or47b ORNs respond to both cis- and trans-palmitoleic acid. As palmitoleic acid is unstable at RT, stocks are stored at -20 &#176;C and used within a month upon opening. Ethanol is the solvent of choice for palmitoleic acid.
<ol>
	<li>Use a vortex mixer to thoroughly mix 10 &#181;L of cis- or trans-palmitoleic acid stocks or dilutions with 90 &#181;L of 100% ethanol for ten-fold serial dilutions in 1.7 mL microtubes. Prepare fresh palmitoleic acid dilutions daily prior to experiments and use within a day. 
	NOTE: For odorants that are not soluble in ethanol, a glass vial is recommended for preparing odor dilutions with other types of organic solvents.</li>
	<li>Using a P10 micropipette, apply 5 &#181;L of cis-palmitoleic acid solutions of the desired dilutions to the filter paper in each corresponding cartridge. 
	NOTE: The highest dosage (10-1) contains 450 &#181;g of the compound. For trans-palmitoleic acid solutions, apply 4.5 &#181;L instead so that the highest dosage (10-1) also contains 450 &#181;g of the compound.</li>
	<li>To completely evaporate the solvent, place the palmitoleic acid cartridges in a vacuum desiccator for 1 h at RT and 7.59 mmHg of pressure. 
	NOTE: The cartridges can be used for up to 4 h at RT.</li>
</ol>
3. Preparation of Drosophila for Ready Access to the at4 Sensilla for In Vivo Electrophysiological Recordings
NOTE: WT flies (Berlin) are reared in standard cornmeal medium at 25&#176;C in a 12:12 light-dark cycle. Upon eclosion, flies are separated by sex into groups of ten, whereby they are group-housed until 7 d&#160;of age. Or47b ORNs in both male and female flies respond to palmitoleic acid. For simplicity, only male flies are examined in the current study.
<ol>
	<li>Assemble a fly-prep slide: on a glass slide, place a glass coverslip (18 x 18 mm2) on a small amount of modeling clay, forming an ~3&#176; angle with the glass slide. Place double-sided tape on the inner edge of the coverslip and on the area of the slide immediately below. Replace with fresh tape for every day of recording (Figure 2A).</li>
	<li>Use a fly aspirator15 to collect the fly of interest in the tubing and then fit a 200-&#181;L pipette tip over the end of the tubing. Simultaneously flick the tube forward while blowing air into the tube to push the fly to the end of the pipette tip. Use a razor blade to cut just below the body of the fly and 2 heads&#39; lengths above the fly.</li>
	<li>Tamp the bottom of the pipette tip with modeling clay, pushing the fly upward until both the antennae and the clypeus are exposed (Figure 2B). To avoid killing the fly, add only enough clay to expose the antennae and aristae, as this prevents the fly&#39;s abdomen from being crushed. Furthermore, add clay slowly and gently to prevent any sudden constriction. Confirm that the fly is alive by checking for antennal or proboscis movement.</li>
	<li>Use forceps to maneuver the pipette tip that houses the fly. Orient the head so that the clypeus is facing to the right of the observer. Adjust the prep along the coverslip using fine forceps until the lateral side of the antenna lies against the taped coverslip surface (Figure 2B).</li>
	<li>Place a holding rod on the arista to secure the antenna to the double-sided tape to prevent movement (Figure 2B). 
	NOTE: The holding rod is pulled from a borosilicate glass capillary with a pipette puller and held in position with modeling clay (Figure 2A).</li>
	<li>Place the prep on the stage of the rig (Figure 2C). Using the microscope, confirm that the trichoids are visible along the distal-lateral edge of the third segment of the antenna. 
	NOTE: Ideally, the sensilla should be clearly silhouetted against the background, which simplifies their identification and facilitates recording (Figure 3). In this preparation, the majority of the accessible trichoid sensilla are of the at4 type.</li>
	<li>Keep the prep under constant humidified airflow (2 L/min) delivered via a separate air delivery tube from a distance of about 2 cm from the prep (Figure 4), as described previously2,15.</li>
</ol>
4. Recording of at4 Sensillum Activity from Or47b ORNs in the at4 Trichoids in Response to Palmitoleic Acid
<ol>
	<li>Insert the reference electrode into the clypeus (Figure 3A). To avoid tissue damage, ensure that the electrode is inserted just below the surface, where it can contact the hemolymph under the cuticle, with a swift and smooth motion.</li>
	<li>Lower the recording electrode slowlyuntil it enters the same plane of view as the target sensillum (Figure 3B). Record under a 50X objective lens. 
	NOTE: The tough trichoidal cuticle necessitates inserting the recording electrode into the sensillar base, whose wider area provides a larger target that reduces the likelihood that the electrode is deflected away (Figure 3B, inset).</li>
	<li>Before applying odor stimuli to a sensillum, observe the following selection criteria; any trichoid failing to meet these standards should be rejected and another sensillum chosen instead.
	<ol>
		<li>Observe a high signal-to-noise ratio (see Figure 3C for an example).</li>
		<li>Observe identifiable spikes from at4A and at4C neurons (Figure 3C). 
		NOTE: Of note, the at4B spike amplitude appears very similar to at4A10 and cannot be identified easily without odor stimulation.</li>
		<li>Observe that the basal firing rate of the at4A neurons is around or under 20 Hz. 
		NOTE: This criterion is specific for at4A because the basal firing rate for the neuron is higher than that of the basiconic ORNs2. A much higher basal firing indicates that the neurons may have been damaged during electrode insertion.</li>
	</ol>
	</li>
	<li>Connect the cartridge to the odorant-delivery tube. Start with the solvent control and then the odorants, from low to high concentrations. Use the micromanipulator to maneuver the cartridge towards the prep while aiming the cartridge squarely at the head of the prep. Visually confirm that the cartridge is pointed directly at the antenna (Figure 4) from a few millimeters away. 
	NOTE: The goal is to orient the opening of the cartridge directly at the antenna and position it in close proximity to the target tissue.</li>
	<li>Ensure that the odorant cartridge is separated from the recording electrode on its right by 1 - 2 mm and from the fly prep slide below by approximately 1 mm. 
	NOTE: In the setup described here, the odorant cartridge is closely bordered by the recording electrode, the reference electrode, and the fly-prep slide (Figure 4). 
	NOTE: Pay attention to the distance between the cartridge and the recording/reference electrodes. A distance of around 4 mm is recommended1. Unintentional contact may terminate the signal and break the tip of the recording electrode, damaging the current neuron and complicating further recordings. 
	NOTE: Consider the distance separating the cartridge and the fly-prep slide. Touching the coverslip may also dislodge the recording electrode to disrupt the recording.</li>
	<li>Press &#34;Record&#34; in the data acquisition software to begin the recording. 
	NOTE: For each 10 s recording, a single 500 ms odor pulse is delivered directly to the antenna, as described in step 1.9.</li>
	<li>After odorant application, carefully retract the cartridge before replacing it with a cartridge of the next-highest concentration. Continue until the entire dosage range is obtained. 
	NOTE: It is recommended that only one Or47b ORN is recorded from each fly to avoid any possible effects of adaptation.</li>
	<li>Thoroughly rinse the recording electrode with distilled water after finishing recording for the day.</li>
	<li>Analyze and plot the data using commercially available offline analysis software.</li>
</ol>

<ol>
	<li>Lin, H. -. H., 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=Hormonal+modulation+of+pheromone+detection+enhances+male+courtship+success.">Hormonal modulation of pheromone detection enhances male courtship success.</a> Neuron. 90 (6), 1272-1285 (2016).</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 (1), 143-160 (2006).</li><li>Silbering, A. F., 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=Complementary+function+and+integrated+wiring+of+the+evolutionarily+distinct+Drosophila+olfactory+subsystems.">Complementary function and integrated wiring of the evolutionarily distinct Drosophila olfactory subsystems.</a> J Neurosci. 31 (38), 13357-13375 (2011).</li><li>Min, S., Ai, M., Shin, S. A., Suh, G. S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dedicated+olfactory+neurons+mediating+attraction+behavior+to+ammonia+and+amines+in+Drosophila.">Dedicated olfactory neurons mediating attraction behavior to ammonia and amines in Drosophila.</a> Proc Nat Acad Sci USA. 110, 1321-1329 (2013).</li><li>Kurtovic, A., Widmer, A., Dickson, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+single+class+of+olfactory+neurons+mediates+behavioural+responses+to+a+Drosophila+sex+pheromone.">A single class of olfactory neurons mediates behavioural responses to a Drosophila sex pheromone.</a> Nature. 446 (7135), 542-546 (2007).</li><li>Van der Goes van Naters, W., 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=Receptors+and+neurons+for+fly+odors+in+Drosophila.">Receptors and neurons for fly odors in Drosophila.</a> Curr Biol. 17, 606-612 (2007).</li><li>Schlief, M. L., Wilson, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Olfactory+processing+and+behavior+downstream+from+highly+selective+receptor+neurons.">Olfactory processing and behavior downstream from highly selective receptor neurons.</a> Nat Neurosci. 10 (5), 623-630 (2007).</li><li>Laughlin, J. D., Ha, T. S., Jones, D. N. M., Smith, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+Pheromone-sensitive+neurons+is+mediated+by+conformational+activation+of+pheromone-binding+protein.">Activation of Pheromone-sensitive neurons is mediated by conformational activation of pheromone-binding protein.</a> Cell. 133 (7), 1255-1265 (2008).</li><li>Gomez-Diaz, C., Reina, J. H., Cambillau, C., Benton, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ligands+for+pheromone-sensing+neurons+are+not+conformationally+activated+odorant+binding+proteins.">Ligands for pheromone-sensing neurons are not conformationally activated odorant binding proteins.</a> PLoS Biol. 11 (4), e1001546 (2013).</li><li>Dweck, H. K. 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=Pheromones+mediating+copulation+and+attraction+in+Drosophila.">Pheromones mediating copulation and attraction in Drosophila.</a> Proc Nat Acad USA. 112, 2829-2835 (2015).</li><li>Cappa, C. D., Lovejoy, E. R., Ravishankara, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaporation+rates+and+vapor+pressures+of+the+even-numbered+C8-C18monocarboxylic+acids.">Evaporation rates and vapor pressures of the even-numbered C8-C18monocarboxylic acids.</a> J Phys Chem A. 112 (17), 3959-3964 (2008).</li><li>Kimura, K. -. I., Sato, C., Yamamoto, K., Yamamoto, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+the+back+or+front:+the+courtship+position+is+a+matter+of+smell+and+sight+in+Drosophila+melanogaster+males.">From the back or front: the courtship position is a matter of smell and sight in Drosophila melanogaster males.</a> J Neurogenet. 29 (1), 18-22 (2015).</li><li>Grosjean, Y., 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=An+olfactory+receptor+for+food-derived+odours+promotes+male+courtship+in+Drosophila.">An olfactory receptor for food-derived odours promotes male courtship in Drosophila.</a> Nature. 478 (7368), 236-240 (2011).</li><li>Wang, J. W., Wong, A. M., Flores, J., Vosshall, L. B., Axel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+calcium+imaging+reveals+an+odor-evoked+map+of+activity+in+the+fly+brain.">Two-photon calcium imaging reveals an odor-evoked map of activity in the fly brain.</a> Cell. 112 (2), 271-282 (2003).</li><li>Pellegrino, M., Nakagawa, T., Vosshall, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+sensillum+recordings+in+the+insects+Drosophila+melanogaster+and+Anopheles+gambiae.">Single sensillum recordings in the insects Drosophila melanogaster and Anopheles gambiae.</a> J Vis Exp. (36), e1-e5 (2010).</li></ol>

The authors have nothing to disclose.

Here, we described a procedure by which the responses of Or47b ORNs to palmitoleic acid can be robustly induced and recorded. We modified a conventional long-distance odor delivery method2,7,10 to troubleshoot the problem of insufficient pheromone odorant delivery. We addressed the issue of low odorant volatility by delivering the compound via odorant cartridges, the opening of which are positioned within millimeters of the prep...

Drosophila ORNs respond to a vast number of odorants, with widely ranging carbon chain lengths and a variety of functional groups, including esters, alcohols, ketones, lactones, aldehydes, terpenes, organic acids, amines, sulfur compounds, heterocyclics, and aromatics2,3. Odorants varied in their physicochemical features can have markedly different volatilities, indicated by the vapor pressure of the compound. Notably, biologically relevant odorants for Drosophila melanogaster differ tremendously in their volatility. For example, Ir92a ORNs respond to ammonia4, which is highly volatile, with a vapor pressure of 6,432 mmHg at 20 &#176;C. In contrast, Or67d ORNs respond to a male pheromone, cis-vaccenyl acetate (cVA)5,6, the vapor pressure of which is 43 mmHg at 20 &#176;C.
Studying the olfactory response to odorants of low volatility is particularly challenging with conventional odor-delivery methods, in which odorants are delivered via a carrier air stream over a relatively long distance (i.e.&#160;several centimeters). As such, the reported olfactory responses to a given low-volatility odorant can vary greatly, depending on the design of the odor-delivery system. For example, the reported response of Or67d ORNs to a high dose of cVA ranges from ~407&#160;- &#62;200 spikes/s6. Moreover, the ineffective delivery of cVA with conventional delivery methods is likely attributed to false-negative results, leading to the interpretation that cVA by itself is not sufficient to activate Or67d ORNs8. This interpretation was later challenged by another study using a close-range odor-delivery method9. It is therefore imperative to develop a robust odor-delivery system for the effective presentation of odorants of low volatility.
Recently, we identified several long-chain cuticular fatty acids as ligands for Or47b ORNs. They are housed in the type 4 Antennal Trichoid Sensillum (at4). Among the long-chain fatty acid odorants, we found that palmitoleic acid functions as an aphrodisiac pheromone that promotes male courtship by activating Or47b ORNs1. However, in another study using a conventional odor-delivery method, methyl laurate was shown to elicit responses from Or47b ORNs, while palmitoleic acid evoked no response when presented from the same distance10. Compared to cVA, long-chain fatty acids are even less volatile, with vapor pressures less than 0.001 mmHg at 25 &#176;C11. The inherently low volatility of long-chain fatty acid odorants, which precludes efficient presentation to the antenna via conventional odor-delivery systems, likely accounted for the false-negative results10. This inconsistency highlights the inadequacy of conventional odor-delivery systems in presenting low-volatility odorants. It was previously shown that the effective delivery of fly cuticular odors requires close proximity between the odor source and the target tissue6. Thus, to fully characterize the effects of biologically active pheromones while mimicking the distance from which they are likely encountered by fruit flies in nature12,13, we agreed that minimal distance must be accorded high priority in our procedure.
Our method holds further advantages, including compatibility with standard electrophysiology rigs and techniques. Pre-existing rig setups require minimal modification to accommodate this protocol, and most SSR steps require only minor adjustments. This renders our technique readily accessible to researchers experienced in SSR. Furthermore, our technique allows for the presentation of low-volatility odorants with sharp onset and offset, correlating stimulus delivery with neuronal response. Finally, the hardware layout facilitates rapid exchanges between odorant cartridges, expediting data collection over a desired dosage range.
We begin by reviewing the preparation of reference and recording electrodes, Adult Hemolymph-Like (AHL) solution, odorant delivery cartridges, and the corresponding olfactometer. We then discuss the preparation of the palmitoleic acid odorant solutions, followed by the preparation of the fly for recording. We proceed to consider the criteria for selecting a trichoid sensillum to record and more closely examine the positioning of the odorant cartridge before presenting representative data acquired using this method. Finally, we conclude by exploring useful applications of this technique, some encountered issues, and their solutions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Prep Setup &#38; Miscellaneous Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Puller Instrument&#160;</td>
			<td>Sutter Instruments 
			Novato CA USA</td>
			<td>P97</td>
			<td>Pipette Puller</td>
		</tr>
		<tr>
			<td>Borosilicate Glass Capillaries</td>
			<td>World Precision Instruments 
			Sarasota FL USA</td>
			<td>1B100F-4</td>
			<td>to make holding rods</td>
		</tr>
		<tr>
			<td>Aluminosilicate Glass Capillaries&#160;</td>
			<td>Sutter Instruments 
			Novato CA USA</td>
			<td>AF100-64-10</td>
			<td>to make electrodes</td>
		</tr>
		<tr>
			<td>Superfrost Microscope Slides</td>
			<td>Fisher Scientific 
			Pittsburgh PA USA</td>
			<td>12-550-143</td>
			<td>for fly-prep station</td>
		</tr>
		<tr>
			<td>Permanent Double Sided Tape</td>
			<td>Scotch 
			St. Paul MN USA</td>
			<td>NA</td>
			<td>for fly-prep station</td>
		</tr>
		<tr>
			<td>Upright microscope</td>
			<td>Olympus 
			Shinjuku Tokyo Japan</td>
			<td>BX51</td>
			<td>for recording rig</td>
		</tr>
		<tr>
			<td>Plastalina modeling clay</td>
			<td>Van Aken 
			North Charleston SC USA</td>
			<td>B0019QZMQQ</td>
			<td>for prep station and to stablize the holding rod</td>
		</tr>
		<tr>
			<td>Rapid-Flow Sterile Disposable Filter Unit with SFCA Membrane, 0.45 mm</td>
			<td>Nalgene 
			Rochester NY USA</td>
			<td>#156-4045</td>
			<td>to sterilize AHL solution</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company&#160;&#160;&#160;</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Cartridge Materials</td>
			<td>&#160;&#160;&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;L pipette tip&#160;</td>
			<td>VWR 
			Radnor PA USA</td>
			<td>53508-810</td>
			<td>to make odor cartridges and fly prep</td>
		</tr>
		<tr>
			<td>Filter Paper</td>
			<td>Whatman 
			Maidstone Kent UK</td>
			<td>740-E</td>
			<td>to make odor cartridges&#160;</td>
		</tr>
		<tr>
			<td>Vacuum Desiccator&#160;</td>
			<td>Cole-Parmer 
			Vernon Hills IL USA</td>
			<td>VX-06514-30</td>
			<td>to vaporize ethanol solvent</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company&#160;&#160;&#160;</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Odorant Materials</td>
			<td>&#160;&#160;&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>cis-palmitoleic acid</td>
			<td>Cayman Chemical 
			Ann Arbor MI USA</td>
			<td>#10009871 (CAS # 373-49-9)</td>
			<td>Or47b odorant</td>
		</tr>
		<tr>
			<td>trans-palmitoleic acid</td>
			<td>Cayman Chemical 
			Ann Arbor MI USA</td>
			<td>#9001798 (CAS # 10030-73-6)</td>
			<td>Or47b odorant</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Spectrum Chemical MFG.&#160; 
			New Brunswick NJ USA</td>
			<td>E1028-500MLGL</td>
			<td>to dilute palmitoleic acid&#160;</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company&#160;&#160;&#160;</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Rig Setup Materials</td>
			<td>&#160;&#160;&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Odorant Cartridge Micromanipulator</td>
			<td>Siskiyou 
			Grants Pass OR USA</td>
			<td>MX130R</td>
			<td>to position the olfactometer</td>
		</tr>
		<tr>
			<td>Flow Vision software&#160;</td>
			<td>Alicat 
			Tuscon AZ USA</td>
			<td>FLOWVISIONSC</td>
			<td>software to control flow rate</td>
		</tr>
		<tr>
			<td>Mass Controller</td>
			<td>Alicat 
			Tuscon AZ USA</td>
			<td>MC-2SLPM-D</td>
			<td>to control the flow rate for humidified air</td>
		</tr>
		<tr>
			<td>Mass Controller</td>
			<td>Alicat 
			Tuscon AZ USA</td>
			<td>MC-500SCCM-D</td>
			<td>to control the flow rate for odor stimulation</td>
		</tr>
		<tr>
			<td>Clampex</td>
			<td>Molecular Devices 
			Sunnyvale CA USA</td>
			<td>Ver. 10.4</td>
			<td>Data acquisition software</td>
		</tr>
		<tr>
			<td>Air delivery tube</td>
			<td>Ace Glass 
			Vineland NJ USA</td>
			<td>8802-936&#160;</td>
			<td>to deliver humidified air</td>
		</tr>
		<tr>
			<td>50x objective lens&#160;</td>
			<td>Olympus 
			Shinjuku Tokyo Japan</td>
			<td>LMPLFL50X</td>
			<td>recording rig</td>
		</tr>
		<tr>
			<td>Clampfit 10</td>
			<td>Molecular Devices 
			Sunnyvale CA USA</td>
			<td>Ver. 10.4</td>
			<td>software for spike analysis&#160;</td>
		</tr>
		<tr>
			<td>Igor Pro 6</td>
			<td>WaveMetrics 
			Lake Oswego OR USA</td>
			<td>Ver. 6.37</td>
			<td>software for data analysis&#160;</td>
		</tr>
		<tr>
			<td>Audio Monitor</td>
			<td>ALA Scientific Instruments 
			Farmingdale NY USA</td>
			<td>NPIEXB-AUDIS-08B</td>
			<td>Aurally reports individual spikes</td>
		</tr>
		<tr>
			<td>Extracellular Amplifier</td>
			<td>ALA Scientific Instruments 
			Farmingdale NY USA</td>
			<td>NPIEXT-02F</td>
			<td>to increase the amplitude of electrical signals</td>
		</tr>
		<tr>
			<td>Valve Controller</td>
			<td>Warner Instruments&#160;&#160;&#160;</td>
			<td>VC-8</td>
			<td>to control the opening of the valve for odor stimulation</td>
		</tr>
		<tr>
			<td>Recording Electrode Micromanipulator</td>
			<td>Sutter Instruments 
			Novato CA USA</td>
			<td>MP-285</td>
			<td>to position recording electrode</td>
		</tr>
		<tr>
			<td>Headstage Amplifier</td>
			<td>ALA Scientific Instruments 
			Farmingdale NY USA</td>
			<td>EQ-16.0008</td>
			<td>to increase the amplitude of electrical signals</td>
		</tr>
		<tr>
			<td>Oscilloscope</td>
			<td>Tektronix 
			Beaverton OR USA</td>
			<td>TDS2000C</td>
			<td>Visual report of individual spikes</td>
		</tr>
	</tbody>
</table>

electrophysiological recording from drosophila trichoid sensilla in response to odorants of low volatility

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, oviposition, and mating. Myriad chemicals of varying volatilities have been identified as natural odorants that activate insect Olfactory Receptor Neurons (ORNs). However, studying the olfactory responses to low-volatility odorants has been hampered by an inability to effectively present such stimuli using conventional odor-delivery methods. Here, we describe a procedure that permits the effective presentation of low-volatility odorants for in vivo Single-Sensillum Recording (SSR). By minimizing the distance between the odor source and the target tissue, this method allows for the application of biologically salient but hitherto inaccessible odorants, including palmitoleic acid, a stimulatory pheromone with a demonstrated effect on ORNs involved in courtship and mating behavior1. Our procedure thus affords a new avenue to assay a host of low-volatility odorants for the study of insect olfaction and pheromone communication.

Our technique was successfully applied to determine the relative efficacy of the trans (Figure 5A) versus cis (Figure 5B) isomers of palmitoleic acid. Our representative data demonstrates that trans-palmitoleic acid is a more effective ligand for Or47b ORNs when compared to the cis isoform (Figure 5C). A single neuron was recorded from each fly, with twelve flies r...

Watch this Scientific Journal Video about Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility at JoVE.com

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

The overall goal of this protocol is to demonstrate how to present odorants of low volatility for single-sensillum recording from Drosophila olfactory receptor neurons that respond to long-chain cuticular pheromones.

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, ...

electrophysiological-recording-from-drosophila-trichoid-sensilla

Research

JoVE Journal

Neuroscience

8.8K Views.  University of California San Diego. The overall goal of this protocol is to demonstrate how to present odorants of low volatility for single-sensillum recording from Drosophila olfactory receptor neurons that respond to long-chain cuticular pheromones.

Video: Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) pacemakers or clocks. Malfunctions in the human circadian system are associated with numerous diseases or disorders. Much progress towards our understanding of the mechanisms underlying circadian rhythms has emerged from genetic screens whereby an easily measured behavioral rhythm is used as a read-out of clock function. Studies using Drosophila have made seminal contributions to our understanding of the cellular and biochemical bases underlying circadian rhythms. The standard circadian behavioral read-out measured in Drosophila is locomotor activity. In general, the monitoring system involves specially designed devices that can measure the locomotor movement of Drosophila. These devices are housed in environmentally controlled incubators located in a darkroom and are based on using the interruption of a beam of infrared light to record the locomotor activity of individual flies contained inside small tubes. When measured over many days, Drosophila exhibit daily cycles of activity and inactivity, a behavioral rhythm that is governed by the animal's endogenous circadian system. The overall procedure has been simplified with the advent of commercially available locomotor activity monitoring devices and the development of software programs for data analysis. We use the system from Trikinetics Inc., which is the procedure described here and is currently the most popular system used worldwide. More recently, the same monitoring devices have been used to study sleep behavior in Drosophila. Because the daily wake-sleep cycles of many flies can be measured simultaneously and only 1 to 2 weeks worth of continuous locomotor activity data is usually sufficient, this system is ideal for large-scale screens to identify Drosophila manifesting altered circadian or sleep properties.

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

We describe procedures for recording daily locomotor activity rhythms of Drosophila and subsequent data analysis. Locomotor activity rhythms are a reliable behavioral output of animal circadian clocks and are used as the standard readout of clock function when studying circadian mutants or examining how the environment regulates the circadian system.

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Vertical T-maze Choice Assay for Arthropod Response to Odorants

Given the economic importance of insects and arachnids as pests of agricultural crops, urban environments or as vectors of plant and human diseases, various technologies are being developed as control tools. A subset of these tools focuses on modifying the behavior of arthropods by attraction or repulsion. Therefore, arthropods are often the focus of behavioral investigations. Various tools have been developed to measure arthropod behavior, including wind tunnels, flight mills, servospheres, and various types of olfactometers. The purpose of these tools is to measure insect or arachnid response to visual or more often olfactory cues. The vertical T-maze oflactometer described here measures choices performed by insects in response to attractants or repellents. It is a high throughput assay device that takes advantage of the positive phototaxis (attraction to light) and negative geotaxis (tendency to walk or fly upward) exhibited by many arthropods. The olfactometer consists of a 30 cm glass tube that is divided in half with a Teflon strip forming a T-maze. Each half serves as an arm of the olfactometer enabling the test subjects to make a choice between two potential odor fields in assays involving attractants. In assays involving repellents, lack of normal response to known attractants can also be measured as a third variable.

A vertical, T-maze olfactometer is described for assaying the behavioral response of arthropods. The olfactometer allows the experimenter to measure choices performed by test subjects when subjected to two potential odor fields. Both attraction to and repulsion from odorants can be measured with this device.

Given the economic importance of insects and arachnids as pests of agricultural crops, urban environments or as vectors of plant and human diseases, ...

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

This protocol describes extracellular recording of the action potential responses fired by labellar taste neurons in Drosophila.

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

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

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

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

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

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

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, enabling the use of D. melanogaster molecular genetics to study inositol-lipid signaling and Transient Receptor Potential (TRP) channels at the single-molecule level. A handful of labs have mastered this powerful technique, which enables the analysis of the physiological responses to light under highly controlled conditions. This technique allows control over the intracellular and extracellular media; the membrane voltage; and the fast application of pharmacological compounds, such as a variety of ionic or pH indicators, to the intra- and extracellular media. With an exceptionally high signal-to-noise ratio, this method enables the measurement of dark spontaneous and light-induced unitary currents (i.e.&#160;spontaneous and quantum bumps) and macroscopic Light-induced Currents (LIC) from single D. melanogaster photoreceptors. This protocol outlines, in great detail, all the key steps necessary to perform this technique, which includes both electrophysiological and optical recordings. The fly retina dissection procedure for the attainment of intact and viable ex vivo isolated ommatidia in the bath chamber is described. The equipment needed to perform whole-cell and fluorescence imaging measurements are also detailed. Finally, the pitfalls in using this delicate preparation during extended experiments are explained.

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell recordings from Drosophila melanogaster photoreceptors enable the measurement of spontaneous dark bumps, quantum bumps, macroscopic responses to light, and current-voltage relationships under various conditions. In combination with D. melanogaster genetic manipulation tools, this method enables the study of the ubiquitous inositol-lipid signaling pathway and its target, the TRP channel.