This work was supported by an NRSA predoctoral grant 1F31DC012985 (to R.D.) and by NIH grants to J.C.
We would like to thank Dr. Linnea Weiss for helpful comments on the manuscript, Dr. Ryan Joseph for help compiling figures, and Dr. Frederic Marion-Poll for helpful technical advice. We would also like to acknowledge the helpful comments of four reviewers.

The following protocol complies with all the animal care guidelines of Yale University.
1. Reagents and Equipment Preparation
<ol>
	<li>Recording equipment setup (Figure 1A).
	<ol>
		<li fo:keep-together.within-page="always">Choose a room for rig setup that is free of large variations in temperature or humidity and also isolated from sources of electrical and mechanical noise, such as refrigerators and centrifuges. 
		<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51355/51355fig1highres.jpg" src="/files/ftp_upload/51355/51355fig1.jpg" /> 
		Figure 1.&#160;(A) Overview of recording rig setup. Stereomicroscope (a) is mounted on anti-vibration platform (b). Reference electrode holder (c) is mounted on platform opposite the headstage (d), via micromanipulators. An outlet plastic tube (e) delivering humidified air stream directed at the fly preparation is also mounted on the platform. The headstage is connected to the amplifier (f), which is connected to the digital acquisition system (DAS) (g), which is connected to a PC (h). (B) Configuration of electrodes and outlet tube: reference electrode on the left, recording electrode on the right, and air stream outlet tube directed at fly preparation. <a href="https://www.jove.com/files/ftp_upload/51355/51355fig1highres.jpg" target="_blank">Click here to view larger image.</a></li>
		<li>Mount stereomicroscope to center of anti-vibration table or platform.</li>
		<li>Attach micromanipulators for the reference electrode/insect preparation and headstage/recording electrode to the left and right of the microscope, respectively, using magnetic stands.</li>
		<li>Mount outlet plastic tube in a third micromanipulator to the rear of the microscope, oriented such that the tube opening is pointed toward location of fly prep (see Figure 1B).</li>
		<li>Using flexible plastic tubing, attach outlet plastic tube to a vacuum flask partially filled with water. Connect a small aquarium pump to bubble air through the water in the flask, generating a humidified air stream through the outlet plastic tube towards the fly.</li>
		<li>Mount fiber optic light source off the vibration table, orienting the outputs to illuminate the preparation by reflecting light via a piece of white card paper directly below the preparation. Ensure that the light source does not rest on the table. Note: the benefit of reflecting the light source on a paper disc is two-fold: it improves the contrast, making sensilla easier to visualize, and it prevents heating of the preparation that would result from direct light.</li>
		<li>Plug tastePROBE amplifier into the digital acquisition system (DAS), and the DAS into a personal computer, according to supplier manual. Plug foot pedal trigger in and arrange under workspace. Note: Electrically isolated wall sockets for the amplifier and DAS are highly desirable.</li>
		<li>Electrically ground microscope, micromanipulators, and light source by connecting metal components to table using alligator clips and lengths of insulated electrical wire and electrical tape. Electrically ground metal platform by connecting to building ground or DAS, which is grounded through power supply plug.</li>
		<li>Install appropriate acquisition software for the DAS of choice on the personal computer. Note: Ensure that the digital acquisition drivers are compatible with the operating system on the PC.</li>
		<li>Configure software amplification (10-100x), signal filtering (typically a Bessel bandpass filter set from 100 Hz-3,000 Hz), and sampling rate (at least 10 KHz). Note: Signal amplitudes from gustatory neurons are typically in the 0.5-2 mV range, so the display scale is set to facilitate their visualization. Note: The 100 Hz filter helps to exclude extraneous electrical noise; however, it changes the shape of spikes and can make advanced spike sorting more challenging. Alternatively, a 1 Hz filter can be used.</li>
		<li>Optionally, a Faraday cage can be set up around the whole vibration table. However, small sheets of aluminum foil are usually sufficient to reduce any noise generated by the external environment or investigator.</li>
	</ol>
	</li>
	<li fo:keep-together.within-page="always">Glass Electrode Preparation 
	<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51355/51355fig2highres.jpg" src="/files/ftp_upload/51355/51355fig2.jpg" /> 
	Figure 2.&#160;Reference and recording electrodes. Photograph under magnification of glass capillaries pulled into reference electrode, with (A) and without (B) tip broken, and recording electrode (C). White bar represents 2 mm. <a href="https://www.jove.com/files/ftp_upload/51355/51355fig2highres.jpg" target="_blank">Click here to view larger image.</a>
	<ol>
		<li>Pull the reference electrode from a glass capillary using a pipette puller instrument. Note: The exact settings of the pipette puller program will vary from instrument to instrument. Try to achieve a very long gradual taper. The pore size at the tip is not crucial because the tip will be broken before fly preparation (Figures 2A and 2B); however, make sure that the diameter of the tapered length of the electrode is neither too thin, which will not allow for sufficient immobilization of the labellum, nor too large, which could damage the gustatory neurons or rupture the salivary glands.</li>
		<li>Pull recording electrode from a borosilicate glass capillary with filament using a pipette puller instrument. Try to achieve a taper that is shallower than that of the reference electrode, and a pore diameter of approximately 10-15 &#956;m (Figure 2C)28.</li>
	</ol>
	</li>
	<li>Tastant solutions preparation
	<ol>
		<li>Use Beadle-Ephrussi Ringer solution (B&#38;E) as the reference electrode electrolyte. To make one liter of B&#38;E, dissolve 7.5 g NaCl, 0.35 g KCl, and 0.279 g CaCl2&#8729;2H2O in one liter of ultrapure water. Store smaller aliquots at -20 &#176;C.</li>
		<li>Use 30 mM tricholine citrate solution (TCC) as the recording electrode electrolyte and solvent for tastant solutions25, if bitter or sugar GRN responses are to be measured. Alternatively, 1-3 mM potassium chloride solution can be used if responses of the water cell are to be measured.</li>
		<li>To make tastant solutions, weigh appropriate amount of tastant in powder form and add to TCC to make an initial stock concentration. Use this to make serial dilutions from this initial stock to yield the desired concentration for testing. Note: If tastants do not readily dissolve in water, another solvent, such as ethanol, can be used to make initial stock concentration. An appropriate control solution of TCC and solvent without tastant should be used in this case.</li>
		<li>Store aliquots long term at -20 &#176;C. Store one working aliquot of a tastant solution at 4 &#176;C for recording use for up to a week, depending on chemical properties of tastant.</li>
	</ol>
	</li>
</ol>
2. Drosophila&#160;Preparation
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51355/51355fig3highres.jpg" src="/files/ftp_upload/51355/51355fig3.jpg" /> 
Figure 3.&#160;Preparation of fly for recording. (A) Insertion position of reference electrode into dorsal thorax of fly. The white arrow indicates the reference electrode. (B) Intermediate position of reference electrode: advanced through neck and head, proboscis not yet extended. (C,D) Fly with reference electrode in final position with tip of electrode inside labellum, and proboscis fully extended. <a href="https://www.jove.com/files/ftp_upload/51355/51355fig3highres.jpg" target="_blank">Click here to view larger image.</a>
<ol>
	<li>Collect newly eclosed flies for recording from well-maintained fly cultures, grown under temperature- and humidity-controlled conditions, and age them 5-10 days in fresh culture vials before recording.</li>
	<li>Chill microscope plate on ice for 15-30 min before preparing fly.</li>
	<li>Backfill glass reference electrode with B&#38;E solution using a long, thin plastic needle of 0.5 mm diameter, such as a spinal needle and 1&#160;ml syringe and gently tap out any bubbles. Break small amount of tip off using forceps and use capillary action to draw out all remaining bubbles with a tissue, observing under dissecting microscope.</li>
	<li>Slide B&#38;E-filled reference electrode onto wire of reference electrode holder, taking care not to introduce air bubbles.</li>
	<li>Aspirate fly into a P200 pipette tip, using fly aspirator built from tubing, mesh, and pipette tip;29 place in ice bucket and chill for 30-60 sec.</li>
	<li>Remove microscope plate from ice, wipe off any moisture, and position underneath microscope. Gently tap fly out of pipette tip onto microscope plate. 
	Note: the fly should be sufficiently immobilized to manipulate easily.</li>
	<li>Under low magnification, gently remove the forelegs with one pair of forceps, while holding the thorax stable with the other pair of forceps. Position the fly on its ventral side, dorsal side facing up. Note: Always be careful to avoid touching the labellum with the forceps at all times during the preparation process to minimize mechanical damage.</li>
	<li>While holding the fly in place with one pair of forceps, insert the reference electrode at the midline of the posterior dorsal thorax. A suggested angle of entry is approximately forty-five degrees, in the direction of the head (Figure 3A).</li>
	<li>Secure the reference electrode holder with modeling clay such that the fly is visible underneath the microscope at high magnification. Maneuver and angle the glass electrode through the neck and head, by sliding the fly towards the reference electrode holder using two pairs of forceps. Note: Work quickly but smoothly; it is easier to complete this step while the fly is still immobilized from the cold (Figure 3B).</li>
	<li>Gently extend the proboscis with one pair of forceps, while sliding the fly further down the glass reference electrode, until the tip of the electrode is inside the labellum and the proboscis is fully extended (Figures 3C and 3D). Note: Take care not to puncture any part of the proboscis tissue or distend the edge of the labellum with the reference electrode, as this may damage the fly and/or taste neurons and affect the recording quality.</li>
</ol>
3. Recording from Labellar Sensilla
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/51355/51355fig4highres.jpg" src="/files/ftp_upload/51355/51355fig4.jpg" /> 
Figure 4.&#160;Recording from fly. (A) labellum of fly preparation on left with recording electrode aligned for contact on right, under high magnification. (B) recording electrode and single sensillum on labellum in contact, under high magnification. <a href="https://www.jove.com/files/ftp_upload/51355/51355fig4highres.jpg" target="_blank">Click here to view larger image.</a>
<ol>
	<li>Always ground yourself by touching the metal surface of the anti-vibration table or platform prior to touching any equipment during recording process! Note: It is extremely important not to deliver a static charge to the headstage as that can damage the circuitry.</li>
	<li>Secure reference electrode holder to micromanipulator mounted on air table of recording rig. Position one lobe of labellum in microscope field of view, under high magnification (typically at least 140X), and in line with humidified air stream.</li>
	<li>Turn on humidified air stream, computer, DAS, and amplifier. Open acquisition software.</li>
	<li>Rinse and fill glass recording electrode with desired tastant.
	<ol>
		<li>Rinse glass recording electrode with ultrapure water by using a syringe and plastic tubing28 to pull small amounts of water through the tube at least ten times.</li>
		<li>Rinse recording electrode with tastant at least five times. Fill recording electrode approximately one-third to halfway full with tastant and remove from tubing. If there are air bubbles, tap to release or simply refill the electrode.</li>
		<li>Slide electrode onto silver wire of the headstage quickly and smoothly so as not to introduce air bubbles.</li>
	</ol>
	</li>
	<li>Stimulate single sensillum with tastant-filled recording electrode.
	<ol>
		<li>Use the micromanipulator to bring the recording electrode aligned with sensillum of interest.</li>
		<li>Press foot pedal to trigger acquisition mode of the amplifier.</li>
		<li>Advance the recording electrode with the fine control knob of micromanipulator carefully until it makes contact with tip of sensillum and recording commences.</li>
		<li>Remove electrode after 1-2 sec.</li>
		<li>Repeat step 3.5 with other sensilla, if desired. Note: Wait at least 1 min in between presentations to the same sensillum. If recording with a single tastant for a prolonged period of time, the tastant solution may dry out and the solution in the tip may become more concentrated. This can be remedied by gently contacting the tip of the glass electrode with smooth paper to remove a small amount of liquid by capillary action.</li>
	</ol>
	</li>
	<li>To record responses to another tastant, rinse and load recording electrode with new tastant and repeat step 3.4. Note: Thoroughly rinsing the electrode between tastants is absolutely crucial to avoid cross-contamination.</li>
	<li>Save data files periodically with identifying information, such as date, genotype, and tastants. Note: It is important to keep a written record of the tastant and sensillum identity of each presentation during recording session for data analysis.</li>
</ol>

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Exp. , e1725 (2010).</li><li>Axon Instruments. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Axon Guide for Electrophysiology &amp; Biophysics Laboratory Techniques. , (1993).</li><li>Fujishiro, N., Kijima, H., Morita, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impulse+frequency+and+action+potential+amplitude+in+labellar+chemosensory+neurones+of+Drosophila+melanogaster.">Impulse frequency and action potential amplitude in labellar chemosensory neurones of Drosophila melanogaster.</a> Journal of insect physiology. 30, 317-325 (1984).</li><li>Marion-Poll, F., Tobin, T. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Software+filter+for+detecting+spikes+superimposed+on+a+fluctuating+baseline.">Software filter for detecting spikes superimposed on a fluctuating baseline.</a> Journal of neuroscience. 37, 1-6 (1991).</li><li>Meunier, N., Marion-Poll, F., Lansky, P., Rospars, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimation+of+the+individual+firing+frequencies+of+two+neurons+recorded+with+a+single+electrode.">Estimation of the individual firing frequencies of two neurons recorded with a single electrode.</a> Chem Senses. 28, 671-679 (2003).</li><li>Meunier, N., Marion-Poll, F., Rospars, J. P., Tanimura, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+coding+of+bitter+taste+in+Drosophila.">Peripheral coding of bitter taste in Drosophila.</a> Journal of neurobiology. 56, 139-152 (2003).</li></ol>

The authors have nothing to disclose.

Labellar sensilla vary in the ease of recording due to differences in morphology and anatomical organization. Sometimes a sensillum does not respond to any tastants, even one that is known to elicit a positive response. The frequency with which this occurs varies depending on sensillum type. L sensilla are most consistently responsive and are relatively easy to access due to their length. In general, S sensilla are consistently responsive, but their short length and position on the labellum make good contact challenging....

The sense of taste allows an insect to detect a vast range of soluble chemicals and plays an important role in the acceptance of a nutritious substance, or the rejection of a noxious or toxic one. Taste is also thought to play a role in mate selection, through the detection of pheromones1-5. These important and diverse functions have made the insect taste system a compelling target of investigation into how sensory systems translate environmental cues into relevant behavioral outputs.
The primary unit of the Drosophila melanogaster taste system is the taste hair, or sensillum. Molecules enter the sensillum via a pore at its tip2,6. Sensilla are found on the labellum, the legs, the wing margin, and the pharynx6.&#160;On the labellum, the number and location of sensilla is stereotyped. There are three morphological classes of sensilla based on length: the long (L), intermediate (I), and short (S) sensilla7,8. Each sensillum contains either two (I-type) or four (L- and S- type) gustatory receptor neurons (GRNs)9. Different GRNs respond to different categories of taste stimuli: bitter, sugar, salt and osmolarity7,10 and express different subsets of gustatory receptors8,11-13. Only I and S-type sensilla contain bitter-responsive GRNs8,10. The GRNs project to the subesophageal ganglion (SOG) and their activation by taste molecules is relayed to the higher central nervous system for decoding, resulting in a behavioral response6. The relatively small number of neurons and the amenability to molecular and behavioral analysis make the Drosophila taste system an excellent model for the investigation of gustatory systems in general. The relative ease with which the system can be manipulated via genetic mutation or the GAL4-UAS expression system also serves as a valuable tool14,15.
Because these sensilla protrude from the surface of the labellum, they make excellent targets for electrophysiology. The firing of the GRNs can be monitored using extracellular recording. Historically, the side-wall recording method, which uses a glass electrode inserted into the sensillum to record neuronal activity,26 has been used. However, this method is technically challenging to perform, and it is difficult to record for long from each preparation. The tip-recording method, which measures the response of the neurons with an electrode that simultaneously delivers a tastant, has since become the method of choice9,16. It has been utilized to investigate the taste system of Drosophila melanogaster8,10,17,18 as well as a number of other insect species19-23. It has been greatly facilitated by the development of the tastePROBE amplifier, which overcame one of the major drawbacks of the tip-recording method by compensating for the large potential difference between the reference electrode and the insect sensillum, allowing the GRN action potentials to be recorded without excessive amplification or filtering24. Another important development was the use of tricholine citrate as the recording electrolyte25. TCC suppresses responses from the osmolarity-sensitive GRN and does not stimulate the salt-sensitive GRN, making responses generated by bitter and sugar tastants much easier to analyze25.
Here we describe how tip recording of Drosophila labellar sensilla is currently performed in the Carlson laboratory. This protocol will explain how to establish a suitable electrophysiology rig, how to prepare the fly, and how to perform taste recordings. We also present some representative data obtained by recording from subsets of Drosophila sensilla, as well as some common issues and potential solutions that may be encountered when using this technique.

<table border="0" cellpadding="0" cellspacing="0" width="783">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="84">
			<td height="84" style="height:84px;width:168px;">Stereo Zoom Microscope</td>
			<td style="width:187px;">Olympus&#160;</td>
			<td style="width:211px;">SZX12 DFPLFL1.6x PF eyepieces: WHN10x-H/22</td>
			<td style="width:219px;">capable of ~150x magnification with long working distance table mount stand</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Anti-vibration Table</td>
			<td style="width:187px;">Kinetic Systems</td>
			<td>BenchMate2210</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Micromanipulators</td>
			<td style="width:187px;">Narishige</td>
			<td>NMN-21</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Magnetic stands</td>
			<td style="width:187px;">ENCO</td>
			<td>Model #625-0930</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Reference Electrode Holder</td>
			<td style="width:187px;">Harvard Apparatus</td>
			<td style="width:211px;">ESP/W-F10N</td>
			<td>Can be mounted on 5ml serological pipette for extended range</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">&#160;Silver Wire</td>
			<td style="width:187px;">World Precision Instruments</td>
			<td style="width:211px;">AGW1510</td>
			<td>0.3-0.5mm diameter</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Retort Stand</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>generic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Outlet Plastic Tube</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>generic, 1cm diameter</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Flexible Plastic Tubing</td>
			<td style="width:187px;">Nalgene&#160;</td>
			<td>8000-0060</td>
			<td style="width:219px;">VI grade 1/4 in internal diameter&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">500 ml Conical Flask</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>generic,&#160; with side arm</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Aquarium Pump</td>
			<td style="width:187px;">Aquatic Gardens</td>
			<td style="width:211px;">Airpump 2000</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Fiber Optic Light Source</td>
			<td style="width:187px;">Dolan-Jenner Industries</td>
			<td>Fiber-Lite 2100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">White Card/Paper</td>
			<td style="width:187px;">Whatman</td>
			<td style="width:211px;">1001-110</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Digital Acquisition System</td>
			<td style="width:187px;">Syntech</td>
			<td style="width:211px;">IDAC-4</td>
			<td>Alternative: National Instruments NI-6251 &#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Headstage</td>
			<td style="width:187px;">Syntech</td>
			<td style="width:211px;">DTP-1</td>
			<td>Tasteprobe</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Tasteprobe Amplifier</td>
			<td style="width:187px;">Syntech</td>
			<td style="width:211px;">DTP-1</td>
			<td>Tasteprobe</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Alligator Clips</td>
			<td style="width:187px;">Grainger</td>
			<td style="width:211px;">1XWN7</td>
			<td>Any brand is fine</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Insulated Electrical Wire</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>Generic</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Gold Connector Pins</td>
			<td style="width:187px;">World Precision Instruments</td>
			<td align="right" style="width:211px;">5482</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Personal Computer</td>
			<td style="width:187px;">Dell&#160;</td>
			<td style="width:211px;">Vostro</td>
			<td>Check for compatibility with digital acquisition system and software</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Acquisition Software</td>
			<td style="width:187px;">Syntech</td>
			<td style="width:211px;">Autospike</td>
			<td>Autospike works with IDAC-4; alternatively, use Labview with NI-6251</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Aluminum Foil and/or Faraday Cage</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>Electro-magnetic noise shielding</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;"></td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Borosilicate Glass Capillaries</td>
			<td style="width:187px;">World Precision Instruments</td>
			<td style="width:211px;">1B100F-4</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Pipette Puller</td>
			<td style="width:187px;">Sutter Instrument Company</td>
			<td style="width:211px;">Model P-87 Flaming/Brown Micropipette Puller</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;"></td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Beadle and Ephrussi Ringer Solution</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>See recipe in protocol section</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Tricholine citrate, 65%&#160;</td>
			<td style="width:187px;">Sigma</td>
			<td style="width:211px;">T0252-100G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;"></td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Stereo Microscope</td>
			<td style="width:187px;">Olympus</td>
			<td style="width:211px;">VMZ 1x-4x</td>
			<td style="width:219px;">Capable of 10x-40x magnification</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Ice Bucket</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>Generic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">p200 Pipette Tips</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>Generic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Spinal Needle</td>
			<td style="width:187px;">Terumo</td>
			<td style="width:211px;">SN*2590</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">1ml Syringe</td>
			<td style="width:187px;">Beckton-Dickenson</td>
			<td align="right" style="width:211px;">301025</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Fly Aspirator</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>Assembled from P1000 pipette tips, flexible plastic tubing, and mesh</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Modeling Clay</td>
			<td style="width:187px;"></td>
			<td style="width:211px;"></td>
			<td>Generic</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:168px;">Forceps</td>
			<td style="width:187px;">Fine Science Tools By Dumont</td>
			<td style="width:211px;">11252-00</td>
			<td>#5SF (super-fine tips)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">10ml Syringe&#160;</td>
			<td style="width:187px;">Beckton-Dickinson</td>
			<td align="right" style="width:211px;">301029</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:168px;">Plastic Tubing</td>
			<td style="width:187px;">Tygon</td>
			<td style="width:211px;">R-3603</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Figure 5A shows the response of an L sensillum to a sugar, sucrose. The same sensillum does not respond to a bitter compound, berberine. Figure 5B shows that an I type sensillum, which contains a bitter responsive neuron, displays larger amplitude spikes in response to berberine, and smaller amplitude spikes in response to sucrose. L sensilla display a minimal background response to the solvent control, TCC, while I sensilla display virtually no response to TCC (Figure 5...

Watch this Scientific Journal Video about Electrophysiological Recording From Drosophila Labellar Taste Sensilla at JoVE.com

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

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

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 ...

electrophysiological-recording-from-drosophila-labellar-taste-sensilla

Research

JoVE Journal

Neuroscience

16.2K vues.  Yale University.  Ce protocole décrit l'enregistrement extracellulaire des réponses potentielles d'action tirés par les neurones gustatifs labellar chez la drosophile. 

Vidéo: Electrophysiological Recording From Drosophila Labellar Taste Sensilla

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 ...

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, ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

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 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 ...

Taste Preference Assay for Adult Drosophila

Olfactory and gustatory perception of the environment is vital for animal survival. The most obvious application of these chemosenses is to be able to distinguish good food sources from potentially dangerous food sources. Gustation requires physical contact with a chemical compound which is able to signal through taste receptors that are expressed on the surface of neurons. In insects, these gustatory neurons can be located across the animal's body allowing taste to play an important role in many different behaviors. Insects typically prefer compounds containing sugars, while compounds that are considered bitter tasting are avoided. Given the basic biological importance of taste, there is intense interest in understanding the molecular mechanisms underlying this sensory modality. We describe an adult Drosophila taste assay which reflects the preference of the animals for a given tastant compound. This assay may be applied to animals of any genetic background to examine the taste preference for a desired soluble compound.

Taste Preference Assay for Adult Drosophila

Taste is an important sensory process which facilitates attraction to beneficial substances and avoidance of toxic substances. This protocol describes a simple ingestion assay for determining Drosophila gustatory preference for a given chemical compound.

Olfactory and gustatory perception of the environment is vital for animal survival.  The most obvious application of these chemosenses is to be able to ...

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.

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, ...

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.

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.

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 of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...