Name: electrophysiological recording from drosophila trichoid sensilla in response to odorants of low volatility
Uploaded: 2017-07-27T15:00:00.000+00:00
Duration: 7 min 49 s
Description: Watch this Scientific Journal Video about Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility at JoVE.com

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>

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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><tbody><tr><td>&lt;strong&gt;Prep Setup &amp;amp; Miscellaneous Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Pipette Puller Instrument&amp;nbsp;</td><td>Sutter Instruments&lt;br/&gt; Novato CA USA</td><td>P97</td><td>Pipette Puller</td></tr><tr><td>Borosilicate Glass Capillaries</td><td>World Precision Instruments&lt;br/&gt; Sarasota FL USA</td><td>1B100F-4</td><td>to make holding rods</td></tr><tr><td>Aluminosilicate Glass Capillaries&amp;nbsp;</td><td>Sutter Instruments&lt;br/&gt; Novato CA USA</td><td>AF100-64-10</td><td>to make electrodes</td></tr><tr><td>Superfrost Microscope Slides</td><td>Fisher Scientific&lt;br/&gt; 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&lt;br/&gt; St. Paul MN USA</td><td>NA</td><td>for fly-prep station</td></tr><tr><td>Upright microscope</td><td>Olympus&lt;br/&gt; Shinjuku Tokyo Japan</td><td>BX51</td><td>for recording rig</td></tr><tr><td>Plastalina modeling clay</td><td>Van Aken&lt;br/&gt; 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&lt;br/&gt; Rochester NY USA</td><td>#156-4045</td><td>to sterilize AHL solution</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&amp;nbsp;&amp;nbsp;&amp;nbsp;&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Cartridge Materials&lt;/strong&gt;</td><td>&lt;strong&gt;&amp;nbsp;&amp;nbsp;&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td></tr><tr><td>200 &amp;micro;L pipette tip&amp;nbsp;</td><td>VWR&lt;br/&gt; 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&lt;br/&gt; Maidstone Kent UK</td><td>740-E</td><td>to make odor cartridges&amp;nbsp;</td></tr><tr><td>Vacuum Desiccator&amp;nbsp;</td><td>Cole-Parmer&lt;br/&gt; Vernon Hills IL USA</td><td>VX-06514-30</td><td>to vaporize ethanol solvent</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&amp;nbsp;&amp;nbsp;&amp;nbsp;&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Odorant Materials&lt;/strong&gt;</td><td>&lt;strong&gt;&amp;nbsp;&amp;nbsp;&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td></tr><tr><td>cis-palmitoleic acid</td><td>Cayman Chemical&lt;br/&gt; 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&lt;br/&gt; 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.&amp;nbsp;&lt;br/&gt; New Brunswick NJ USA</td><td>E1028-500MLGL</td><td>to dilute palmitoleic acid&amp;nbsp;</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&amp;nbsp;&amp;nbsp;&amp;nbsp;&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Rig Setup Materials&lt;/strong&gt;</td><td>&lt;strong&gt;&amp;nbsp;&amp;nbsp;&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td></tr><tr><td>Odorant Cartridge Micromanipulator</td><td>Siskiyou&lt;br/&gt; Grants Pass OR USA</td><td>MX130R</td><td>to position the olfactometer</td></tr><tr><td>Flow Vision software&amp;nbsp;</td><td>Alicat&lt;br/&gt; Tuscon AZ USA</td><td>FLOWVISIONSC</td><td>software to control flow rate</td></tr><tr><td>Mass Controller</td><td>Alicat&lt;br/&gt; 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&lt;br/&gt; 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&lt;br/&gt; Sunnyvale CA USA</td><td>Ver. 10.4</td><td>Data acquisition software</td></tr><tr><td>Air delivery tube</td><td>Ace Glass&lt;br/&gt; Vineland NJ USA</td><td>8802-936&amp;nbsp;</td><td>to deliver humidified air</td></tr><tr><td>50X objective lens&amp;nbsp;</td><td>Olympus&lt;br/&gt; Shinjuku Tokyo Japan</td><td>LMPLFL50X</td><td>recording rig</td></tr><tr><td>Clampfit 10</td><td>Molecular Devices&lt;br/&gt; Sunnyvale CA USA</td><td>Ver. 10.4</td><td>software for spike analysis&amp;nbsp;</td></tr><tr><td>Igor Pro 6</td><td>WaveMetrics&lt;br/&gt; Lake Oswego OR USA</td><td>Ver. 6.37</td><td>software for data analysis&amp;nbsp;</td></tr><tr><td>Audio Monitor</td><td>ALA Scientific Instruments&lt;br/&gt; 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&lt;br/&gt; 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&amp;nbsp;&amp;nbsp;&amp;nbsp;</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&lt;br/&gt; Novato CA USA</td><td>MP-285</td><td>to position recording electrode</td></tr><tr><td>Headstage Amplifier</td><td>ALA Scientific Instruments&lt;br/&gt; 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&lt;br/&gt; 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

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 ...

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

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

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

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

Biology

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory bulb1-5. Olfactory information in the external world is transmitted to the antennal lobe by olfactory sensory neurons (OSNs), which segregate to distinct regions of neuropil called glomeruli according to the specific olfactory receptor they express. Here, OSN axons synapse with both local interneurons (LNs), whose processes can innervate many different glomeruli, and projection neurons (PNs), which convey olfactory information to higher olfactory brain regions.
Optical imaging of the activity of OSNs, LNs and PNs in the antennal lobe - traditionally using synthetic calcium indicators (e.g. calcium green, FURA-2) or voltage-sensitive dyes (e.g. RH414) - has long been an important technique to understand how olfactory stimuli are represented as spatial and temporal patterns of glomerular activity in many species of insects6-10. Development of genetically-encoded neural activity reporters, such as the fluorescent calcium indicators G-CaMP11,12 and Cameleon13, the bioluminescent calcium indicator GFP-aequorin14,15, or a reporter of synaptic transmission, synapto-pHluorin16 has made the olfactory system of the fruitfly, Drosophila melanogaster, particularly accessible to neurophysiological imaging, complementing its comprehensively-described molecular, electrophysiological and neuroanatomical properties2,4,17. These reporters can be selectively expressed via binary transcriptional control systems (e.g. GAL4/UAS18, LexA/LexAop19,20, Q system21) in defined populations of neurons within the olfactory circuitry to dissect with high spatial and temporal resolution how odor-evoked neural activity is represented, modulated and transformed22-24.
Here we describe the preparation and analysis methods to measure odor-evoked responses in the Drosophila antennal lobe using G-CaMP25-27. The animal preparation is minimally invasive and can be adapted to imaging using wide-field fluorescence, confocal and two-photon microscopes.

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

We describe an established technique to measure and analyze odor-evoked calcium responses in the antennal lobe of living Drosophila melanogaster.

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory ...

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 ...

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila express gustatory receptors1,2, ion channels3-6, and ionotropic receptors7 that are involved in the detection of volatile and non-volatile sensory cues. These neurons directly contact tastants such as food, noxious substances and pheromones and therefore influence many complex behaviors such as feeding, egg-laying and mating. Electrode recordings and calcium imaging have been widely used in insects to quantify the neuronal responses evoked by these tastants. However, electrophysiology requires specialized equipment and obtaining measurements from a single taste sensillum can be technically challenging depending on the cell-type, size, and position. In addition, single neuron resolution in Drosophila can be difficult to achieve since taste sensilla house more than one type of chemosensory neuron. The live calcium imaging method described here allows responses of single gustatory neurons in live flies to be measured. This method is especially suitable for imaging neuronal responses to lipid pheromones and other ligand types that have low solubility in water-based solvents.

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

The forelegs and proboscis of Drosophila contain a rich repertoire of gustatory sensory neurons. Here, we present a method using calcium imaging to measure physiological responses from sensory neurons in the foreleg and proboscis of live flies upon exogenous application of a gustatory pheromone.

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila ...

Immunology and Infection

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To complicate matters even further, the odors detected by the insect nervous system usually are not single compounds but mixtures whose composition and concentration ratios vary. This leads to an almost infinite amount of different olfactory stimuli which have to be evaluated by the nervous system.
To understand which aspects of an odor stimulus determine its evaluation by the fly, it is therefore desirable to efficiently examine odor-guided behavior towards many odorants and odor mixtures. To directly correlate behavior to neuronal activity, behavior should be quantified in a comparable time frame and under identical stimulus conditions as in neurophysiological experiments. However, many currently used olfactory bioassays in Drosophila neuroethology are rather specialized either towards efficiency or towards resolution.
Flywalk, an automated odor delivery and tracking system, bridges the gap between efficiency and resolution. It allows the determination of exactly when an odor packet stimulated a freely walking fly, and to determine the animal&#180;s dynamic behavioral reaction.

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

The automated tracking system Flywalk is used for high-resolution quantification of odor-guided behavior in Drosophila melanogaster.

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To ...

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 ...

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an excellent model system for such investigations due to its complex behaviors, powerful genetic techniques, and compact nervous system. Laboratory behavioral assays have long been used with Drosophila to simulate properties of the natural environment and study the neural mechanisms underlying the corresponding behaviors (e.g. phototaxis, chemotaxis, sensory learning and memory)1-3. With the recent availability of large collections of transgenic Drosophila lines that label specific neural subsets, behavioral assays have taken on a prominent role to link neurons with behaviors4-11. Versatile and reproducible paradigms, together with the underlying computational routines for data analysis, are indispensable for rapid tests of candidate fly lines with various genotypes. Particularly useful are setups that are flexible in the number of animals tested, duration of experiments and nature of presented stimuli. The assay of choice should also generate reproducible data that is easy to acquire and analyze. Here, we present a detailed description of a system and protocol for assaying behavioral responses of Drosophila flies in a large four-field arena. The setup is used here to assay responses of flies to a single olfactory stimulus; however, the same setup may be modified to test multiple olfactory, visual or optogenetic stimuli, or a combination of these. The olfactometer setup records the activity of fly populations responding to odors, and computational analytical methods are applied to quantify fly behaviors. The collected data are analyzed to get a quick read-out of an experimental run, which is essential for efficient data collection and the optimization of experimental conditions.

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

We describe here a behavioral setup and data analysis method for assaying olfactory responses of up to 100 vinegar flies (Drosophila melanogaster). This system may be used with single or multiple olfactory stimuli, and adaptable for optogenetic activation or silencing of neuronal subsets.

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an ...

Single Sensillum Recordings for Locust Palp Sensilla Basiconica

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially for the detection of non-volatile chemical cues through sensilla chaetica (previously named terminal sensilla or crested sensilla). There is now increasing evidence that these palps also have an olfactory function. An odorant receptor (LmigOR2) and an odorant-binding protein (LmigOBP1) have been localized in the neurons and accessory cells, respectively, in the sensilla basiconica of the palps. Single sensillum recording (SSR) is used for recording the responses of odorant receptor neurons, which is an effective method for screening active ligands on specific odorant receptors. SSR is used in functional studies of odorant receptors in palp sensilla. The structure of the sensilla basiconica located on the dome of the palps differs somewhat from the structure of those on the antennae. Therefore, when performing an SSR elicited by odorants, some specific advice may be helpful for obtaining optimum results. In this paper, a detailed and highly effective protocol for an SSR from insect palp sensilla basiconica is introduced.

This paper describes a detailed and highly effective protocol for single sensillum recordings from the sensilla basiconica on the palps of insect mouthparts.

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially ...

A Novel Method for Enhanced Analysis of Insect Taste Neuron Activity

Base Recording: A Technique for Analyzing Responses of Taste Neurons in Drosophila

Insects taste the external world through taste hairs, or sensilla, that have pores at their tips. When a sensillum comes into contact with a potential food source, compounds from the food source enter through the pore and activate neurons within. For over 50 years, these responses have been recorded using a technique called tip recording. However, this method has major limitations, including the inability to measure neural activity before or after stimulus contact and the requirement for tastants to be soluble in aqueous solutions. We describe here a technique that we call base recording, which overcomes these limitations. Base recording allows the measurement of taste neuron activity before, during, and after the stimulus. Thus, it allows extensive analysis of OFF responses that occur after a taste stimulus. It can be used to study hydrophobic compounds such as long-chain pheromones that have very low solubility in water. In summary, base recording offers the advantages of single-sensillum electrophysiology as a means of measuring neuronal activity - high spatial and temporal resolution, without the need for genetic tools - and overcomes key limitations of the traditional tip recording technique.

Author Spotlight: Advances in Chemoreception &#8211; From Insect Odor Receptors to Non-Coding RNAs

A rarely used method of electrophysiological recording, base recording, allows analysis of features of taste coding that cannot be examined by conventional recording methods. Base recording also allows the analysis of taste responses to hydrophobic stimuli that cannot be studied using traditional electrophysiological methods.

Insects taste the external world through taste hairs, or sensilla, that have pores at their tips. When a sensillum comes into contact with a potential ...

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

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Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

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

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

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

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

Single Sensillum Recordings for Locust Palp Sensilla Basiconica

Base Recording: A Technique for Analyzing Responses of Taste Neurons in Drosophila