We thank Zina Berman for support, Lisa Baik for comments on the manuscript, and other members of the Carlson laboratory for discussion. This work was supported NIH grant K01 DC020145 to H.K.M.D; and NIH grants R01 DC02174, R01 DC04729, and R01 DC011697 to J.R.C.

The following protocol complies with all the animal care guidelines of Yale University.
1. Flies
<ol>
	<li>Place 10-15 newly emerged flies in fresh standard culture vials at 25 &#176;C and 60% relative humidity in a 12:12 h light-dark cycle.</li>
	<li>Use flies when 3-7 days old.</li>
</ol>
2. Chemosensory stimuli
<ol>
	<li>Obtain chemosensory stimuli of the highest available purity. Store them as recommended by the vendor until use.</li>
	<li>Dissolve chemosensory stimuli and dilute to the desired concentrations in water or another desired, non-toxic solvent such as paraffin oil. Stir the prepared solutions for a minimum of 1 h in the case of dissolved solid compounds.</li>
</ol>
3. Glass stimulus capillary
<ol>
	<li>Pull a glass capillary to hold the stimulus from a borosilicate glass capillary (100 mm length, 1 mm outer diameter, 0.58 mm inner diameter) using a pipette puller instrument. Aim to achieve a tip diameter between 3 &#181;m and 10 &#181;m.</li>
	<li>Fill the glass capillary with the preferred stimulus solution using a microloader pipette tip. Take care to avoid bubbles, which can be removed by gentle tapping.</li>
	<li>If the stimulus crystallizes at the tip, clean or replace the glass stimulus capillary.</li>
</ol>
4. Reference and recording electrodes
<ol>
	<li>Use tungsten rods (127 &#181;m diameter and 76.2 mm length) for both reference and recording electrodes. Sharpen the reference and recording electrodes to approximately 1 &#181;m diameter at the tip (the shapes of these electrodes are depicted in Delventhal et al.36) by dipping them repeatedly for several seconds in either a 10% KNO3 (~1 M) solution or a 10% KOH (1.8 M) solution. 
	NOTE: This solution requires current (0.3-3 mA) to facilitate this process.</li>
</ol>
5. Preparation of the fly for base recording
<ol>
	<li>Draw a single fly from the vial into an aspirator. Withdraw the aspirator and trap the animal by placing a finger over the end.</li>
	<li>Expel the fly into a 200 &#181;L plastic pipette tip. Keeping the end of the aspirator in the pipette tip, use the end to push the fly forward, headfirst, toward the narrow end of the pipette tip.</li>
	<li>Trim the pipette tip at each end (i.e., anterior and posterior to the animal) using a razor blade.</li>
	<li>Use either clay or a small piece of cotton to push the fly forward further, until half of the head protrudes from the end of the trimmed pipette tip. Use forceps to gently push until the labellum at the front of the head is exposed.</li>
	<li>Fix the trimmed pipette tip onto a glass microscope slide using clay (Figure 1).</li>
	<li>Under the stereomicroscope, position the labellum laterally on a cover slip so that one lobe, along with its 31 taste sensilla, is exposed (Figure 1). The cover slip keeps the labellum in place.</li>
</ol>
6. Electrophysiology rig
<ol>
	<li>Select a room for the rig setup that has stable temperature and relative humidity (&#60;70%) and is isolated from sources of electrical and mechanical noise, such as refrigerators and centrifuges.</li>
	<li>Place the microscope on the center of an antivibration table.</li>
	<li>Secure a manual micromanipulator to the antivibration table (Figure 2).</li>
	<li>Attach a stainless-steel shaft that holds the tungsten reference electrode to the manual micromanipulator (Figure 2).</li>
	<li>Connect motorized manipulators-one with a holder for the recording electrode probe and a second one with a holder connected to a stainless-steel shaft for the glass stimulus capillary-to the same table using stands (Figure 2).</li>
	<li>Connect the recording electrode probe to an Intelligent Data Acquisition Controller (IDAC) system or another amplifier/digitizer system.</li>
	<li>Link this IDAC system to the computer at the workstation.</li>
	<li>Ground the manual and motorized manipulators to the same location within the rig.</li>
	<li>Install appropriate acquisition software for the IDAC system on the computer. Ensure that the digital acquisition drivers are compatible with the operating system (e.g., Windows XP-7, -8, or -10) on the computer.</li>
</ol>
7. Recording from taste sensilla
<ol>
	<li>Place the preparation slide on the microscope stage with a low magnification (e.g.,&#160;10x) objective in position. Move the stage until the labellum is in focus at the center of the field of view at both low-magnification and high-magnification (e.g., 50x) objectives.</li>
	<li>Insert the reference electrode into the eye using the low magnification objective. To insert the reference electrode, target the eye on the side of the fly opposite the side with the recording electrode, for example, if the recording electrode is approaching from the right, place the reference electrode in the left eye. Utilize a manual micromanipulator for precise insertion.</li>
	<li>Bring the tip of the glass stimulus capillary into focus at the center of the field of view of both low-magnification and high-magnification objectives using a motorized micromanipulator (Figure 3).</li>
	<li>Under low magnification, bring the recording electrode close to the labellum using a second motorized micromanipulator.</li>
	<li>Under high magnification, insert the recording electrode into the base of a taste sensillum using the motorized micromanipulator until the sound of neuronal firing activity from the audio output of the IDAC system is heard.</li>
	<li>Once a stable signal has been established, start recording the signal using the software that comes with the IDAC system (Figure 4A-D). To start recording, press the Start recording button.</li>
	<li>Bring the tip of the stimulus glass capillary to cover the tip of the taste sensillum using the motorized manipulator.</li>
	<li>To end the stimulus, remove the glass stimulus capillary from the sensillum using the motorized manipulator.</li>
	<li>Mark the start and end of the stimulation manually using a pedal. The pedal is connected to the IDAC, and its communication with the software is facilitated through the IDAC to mark the start/end of the stimulus.</li>
</ol>
8. Analysis
<ol>
	<li>Use the different functions of the software that comes with the IDAC system to sort spike populations by amplitude (when possible) and analyze response dynamics.
	<ol>
		<li>To count spikes, left click on the recording of interest, bringing up a window to select from. Choose To Spikes, initiating another window named Convert waves to spikes. Enter a name in the New field and press the OK button.</li>
		<li>Entering the name in the New field in step 8.1.1 leads to the Amplitude Histogram view. Choose the amplitude to count, then close this view. Left click to add a counter.</li>
		<li>Examine the spikes manually to confirm conclusions based on analysis with software. 
		NOTE: The software also allows the exportation of data in different formats for further analysis.</li>
	</ol>
	</li>
</ol>

A Novel Method for Enhanced Analysis of Insect Taste Neuron Activity

<ol>
	<li>Joseph, R. M., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+chemoreceptors:+a+molecular+interface+between+the+chemical+world+and+the+brain.">Drosophila chemoreceptors: a molecular interface between the chemical world and the brain.</a> Trends Genet. 31 (12), 683-695 (2015).</li><li>Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+sensory+receptors-a+set+of+molecular+Swiss+Army+Knives.">Drosophila sensory receptors-a set of molecular Swiss Army Knives.</a> Genetics. 217 (1), 1-34 (2021).</li><li>Dweck, H. K. M., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+logic+and+evolution+of+bitter+taste+in+Drosophila.">Molecular logic and evolution of bitter taste in Drosophila.</a> Curr Biol. 30 (1), 17-30 (2020).</li><li>Weiss, L. A., Dahanukar, A., Kwon, J. Y., Banerjee, D., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+and+cellular+basis+of+bitter+taste+in+Drosophila.">The molecular and cellular basis of bitter taste in Drosophila.</a> Neuron. 69 (2), 258-272 (2011).</li><li>He, Z., Luo, Y., Shang, X., Sun, J. S., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemosensory+sensilla+of+the+Drosophila+wing+express+a+candidate+ionotropic+pheromone+receptor.">Chemosensory sensilla of the Drosophila wing express a candidate ionotropic pheromone receptor.</a> PLoS Biol. 17 (5), e2006619 (2019).</li><li>Ling, F., Dahanukar, A., Weiss, L. A., Kwon, J. Y., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+and+cellular+basis+of+taste+coding+in+the+legs+of+Drosophila.">The molecular and cellular basis of taste coding in the legs of Drosophila.</a> J Neurosci. 34 (21), 7148-7164 (2014).</li><li>Falk, R., Bleiser-Avivi, N., Atidia, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Labellar+taste+organs+of+Drosophila+melanogaster.">Labellar taste organs of Drosophila melanogaster.</a> J Morphol. 150 (2), 327-341 (1976).</li><li>Hiroi, M., Meunier, N., Marion-Poll, F., 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=Two+antagonistic+gustatory+receptor+neurons+responding+to+sweet-salty+and+bitter+taste+in+Drosophila.">Two antagonistic gustatory receptor neurons responding to sweet-salty and bitter taste in Drosophila.</a> J Neurobiol. 61 (3), 333-342 (2004).</li><li>Shanbhag, S. R., Park, S. K., Pikielny, C. W., Steinbrecht, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gustatory+organs+of+Drosophila+melanogaster:+fine+structure+and+expression+of+the+putative+odorant-binding+protein+PBPRP2.">Gustatory organs of Drosophila melanogaster: fine structure and expression of the putative odorant-binding protein PBPRP2.</a> Cell Tissue Res. 304 (3), 423-437 (2001).</li><li>Siddiqi, O., Rodrigues, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+analysis+of+a+complex+chemoreceptor.">Genetic analysis of a complex chemoreceptor.</a> Basic Life Sci. 16, 347-359 (1980).</li><li>Clyne, P. J., Warr, C. G., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Candidate+taste+receptors+in+Drosophila.">Candidate taste receptors in Drosophila.</a> Science. 287 (5459), 1830-1834 (2000).</li><li>Koh, T. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+IR20a+clade+of+ionotropic+receptors+are+candidate+taste+and+pheromone+receptors.">The Drosophila IR20a clade of ionotropic receptors are candidate taste and pheromone receptors.</a> Neuron. 83 (4), 850-865 (2014).</li><li>S&#225;nchez-Alca&#241;iz, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+expression+atlas+of+variant+ionotropic+glutamate+receptors+identifies+a+molecular+basis+of+carbonation+sensing.">An expression atlas of variant ionotropic glutamate receptors identifies a molecular basis of carbonation sensing.</a> Nat Commun. 9 (1), 4252 (2018).</li><li>Dweck, H. K. M., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diverse+mechanisms+of+taste+coding+in+Drosophila.">Diverse mechanisms of taste coding in Drosophila.</a> Sci Adv. 9 (46), (2023).</li><li>Chyb, S., Dahanukar, A., Wickens, A., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Gr5a+encodes+a+taste+receptor+tuned+to+trehalose.">Drosophila Gr5a encodes a taste receptor tuned to trehalose.</a> Proc Natl Acad Sci U S A. 100, 14526-14530 (2003).</li><li>Dahanukar, A., Lei, Y. T., Kwon, J. Y., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+Gr+genes+underlie+sugar+reception+in+Drosophila.">Two Gr genes underlie sugar reception in Drosophila.</a> Neuron. 56 (3), 503-516 (2007).</li><li>Delventhal, R., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bitter+taste+receptors+confer+diverse+functions+to+neurons.">Bitter taste receptors confer diverse functions to neurons.</a> Elife. 5, e11181 (2016).</li><li>Dweck, H. K. M., Talross, G. J. S., Luo, Y., Ebrahim, S. A. M., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ir56b+is+an+atypical+ionotropic+receptor+that+underlies+appetitive+salt+response+in+Drosophila.">Ir56b is an atypical ionotropic receptor that underlies appetitive salt response in Drosophila.</a> Curr Biol. 32 (8), 1776-1787 (2022).</li><li>Hiroi, M., Marion-Poll, F., 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=Differentiated+response+to+sugars+among+labellar+chemosensilla+in+Drosophila.">Differentiated response to sugars among labellar chemosensilla in Drosophila.</a> Zoolog Sci. 19 (9), 1009-1018 (2002).</li><li>Jeong, Y. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+odorant-binding+protein+required+for+suppression+of+sweet+taste+by+bitter+chemicals.">An odorant-binding protein required for suppression of sweet taste by bitter chemicals.</a> Neuron. 79 (4), 725-737 (2013).</li><li>Jiao, Y., Moon, S. J., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Drosophila+gustatory+receptor+required+for+the+responses+to+sucrose,+glucose,+and+maltose+identified+by+mRNA+tagging.">A Drosophila gustatory receptor required for the responses to sucrose, glucose, and maltose identified by mRNA tagging.</a> Proc Natl Acad Sci U S A. 104 (35), 14110-14115 (2007).</li><li>Jiao, Y., Moon, S. J., Wang, X., Ren, Q., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gr64f+is+required+in+combination+with+other+gustatory+receptors+for+sugar+detection+in+Drosophila.">Gr64f is required in combination with other gustatory receptors for sugar detection in Drosophila.</a> Curr Biol. 18 (22), 1797-1801 (2008).</li><li>Kim, S. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+TRPA1+channel+mediates+chemical+avoidance+in+gustatory+receptor+neurons.">Drosophila TRPA1 channel mediates chemical avoidance in gustatory receptor neurons.</a> Proc Natl Acad Sci U S A. 107 (18), 8440-8445 (2010).</li><li>Lacaille, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+inhibitory+sex+pheromone+tastes+bitter+for+Drosophila+males.">An inhibitory sex pheromone tastes bitter for Drosophila males.</a> PLoS One. 2 (7), e661 (2007).</li><li>Lee, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gustatory+receptors+required+for+avoiding+the+insecticide+L-canavanine.">Gustatory receptors required for avoiding the insecticide L-canavanine.</a> J Neurosci. 32 (4), 1429-1435 (2012).</li><li>Lee, Y., Kim, S. H., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Avoiding+DEET+through+insect+gustatory+receptors.">Avoiding DEET through insect gustatory receptors.</a> Neuron. 67 (4), 555-561 (2010).</li><li>Lee, Y., Moon, S. J., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+gustatory+receptors+required+for+the+caffeine+response+in+Drosophila.">Multiple gustatory receptors required for the caffeine response in Drosophila.</a> Proc Natl Acad Sci U S A. 106 (11), 4495-4500 (2009).</li><li>Lee, Y., Moon, S. J., Wang, Y., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Drosophila+gustatory+receptor+required+for+strychnine+sensation.">A Drosophila gustatory receptor required for strychnine sensation.</a> Chem Senses. 40 (7), 525-533 (2015).</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> J Neurobiol. 56 (2), 139-152 (2003).</li><li>Moon, S. J., K&#246;ttgen, M., Jiao, Y., Xu, H., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+taste+receptor+required+for+the+caffeine+response+in+vivo.">A taste receptor required for the caffeine response in vivo.</a> Curr Biol. 16 (18), 1812-1817 (2006).</li><li>Moon, S. J., Lee, Y., Jiao, Y., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Drosophila+gustatory+receptor+essential+for+aversive+taste+and+inhibiting+male-to-male+courtship.">A Drosophila gustatory receptor essential for aversive taste and inhibiting male-to-male courtship.</a> Current Biology. 19, 1623-1627 (2009).</li><li>Rimal, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanism+of+acetic+acid+gustatory+repulsion+in+Drosophila.">Mechanism of acetic acid gustatory repulsion in Drosophila.</a> Cell Rep. 26 (6), 1432-1442 (2019).</li><li>Shim, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+full+repertoire+of+Drosophila+gustatory+receptors+for+detecting+an+aversive+compound.">The full repertoire of Drosophila gustatory receptors for detecting an aversive compound.</a> Nat Commun. 6, 8867 (2015).</li><li>Xiao, S., Baik, L. S., Shang, X., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Meeting+a+threat+of+the+Anthropocene:+Taste+avoidance+of+metal+ions+by+Drosophila.">Meeting a threat of the Anthropocene: Taste avoidance of metal ions by Drosophila.</a> Proc Natl Acad Sci U S A. 119 (25), e2204238119 (2022).</li><li>Zhang, Y. V., Ni, J., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+basis+for+attractive+salt-taste+coding+in+Drosophila.">The molecular basis for attractive salt-taste coding in Drosophila.</a> Science. 340 (6138), 1334-1338 (2013).</li><li>Delventhal, R., Kiely, A., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+recording+from+Drosophila+labellar+taste+sensilla.">Electrophysiological recording from Drosophila labellar taste sensilla.</a> J Vis Exp. (84), e51355 (2014).</li><li>Marella, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+taste+responses+in+the+fly+brain+reveals+a+functional+map+of+taste+category+and+behavior.">Imaging taste responses in the fly brain reveals a functional map of taste category and behavior.</a> Neuron. 49 (2), 285-295 (2006).</li><li>Thorne, N., Amrein, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atypical+expression+of+Drosophila+gustatory+receptor+genes+in+sensory+and+central+neurons.">Atypical expression of Drosophila gustatory receptor genes in sensory and central neurons.</a> J Comp Neurol. 506 (4), 548-568 (2008).</li><li>Wang, Z., Singhvi, A., Kong, P., Scott, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taste+representations+in+the+Drosophila+brain.">Taste representations in the Drosophila brain.</a> Cell. 117 (7), 981-991 (2004).</li></ol>

The authors have no conflicts of interest to disclose.

In recordings from some types of sensilla, it can be challenging to differentiate the spikes of different neurons. For example, the sugar neurons and mechanosensory neurons of S and I sensilla produce spikes of similar amplitudes, making it difficult to distinguish them4,14. We find that the use of a very sharp tungsten recording electrode reduces the firing of the mechanosensory neuron, as does the judicious placement of the recording electrode. Insertion of the...

Insects, including drosophilid flies, are endowed with a sophisticated taste system that enables them to extract complex chemical information from their surroundings. This system allows them to discern the chemical composition of various substances, distinguishing between those that are nutritious and those that are harmful1,2.
At the core of this system are specialized structures known as taste hairs or sensilla, strategically located on various body parts. In drosophilid flies, these sensilla are located on the labellum, which is the major taste organ of the fly head1,2,3,4, as well as on the legs and wings1,2,5,6. The labellum is located at the tip of the proboscis and contains two lobes4,7,8. Each lobe is covered with 31 taste sensilla categorized as short, long, and intermediate4,7,8. These sensilla each house 2-4 taste neurons1,2,9,10. These taste neurons express members of at least four different gene families, namely the Gustatory receptor (Gr), Ionotropic receptor (Ir), Pickpocket (Ppk), and Transient receptor potential (Trp) genes1,2,11,12,13. This diversity of receptors and channels equips insects with the ability to recognize a wide array of chemical compounds, including both nonvolatile and volatile cues1,2,14.
For over 50 years, scientists have quantified the response of taste neurons and their receptors using a technique called tip recording3,4,6,8,13,15,16,17,18,19,20,21,22,23,24,25,26,27,28, 
29,30,31,32,33,34,35. However, this method has major limitations. First, neural activity can be measured only during contact with the stimulus, and not before or after contact. This limitation precludes the measurement of spontaneous spiking activity and prevents the measurement of OFF responses. Second, only tastants that are soluble in aqueous solutions can be tested.
These limitations can be overcome by a rarely used alternative electrophysiological technique called &#34;base recording.&#34; Here we describe this technique, which we have adapted from a method used by Marion-Poll and colleagues24, and show the crucial taste coding features that it can now conveniently measure14.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td>BX51WI</td>
			<td>equipped with a 50X objective (LMPLFLN 50X, Olympus) and 10X eyepieces.&#160;</td>
		</tr>
		<tr>
			<td>Antivibration Table</td>
			<td>TMC</td>
			<td>63-7590E</td>
			<td></td>
		</tr>
		<tr>
			<td>motorized Micromanipulators</td>
			<td>Harvard Apparatus and M&#228;rzh&#228;user Micromanipulators</td>
			<td>Micromanipulator PM 10 Piezo Micromanipulator</td>
			<td></td>
		</tr>
		<tr>
			<td>manual Micromanipulators</td>
			<td>M&#228;rzh&#228;user Micromanipulators</td>
			<td>MM33 Micromanipulator</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic stands</td>
			<td>ENCO</td>
			<td>Model #625-0930</td>
			<td></td>
		</tr>
		<tr>
			<td>Reference&#160; and recording Electrode Holder</td>
			<td>Ockenfels Syntech GmbH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stimulus glass capillary Holder</td>
			<td>Ockenfels Syntech GmbH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Universal Single Ended Probe</td>
			<td>Ockenfels Syntech GmbH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4-CHANNEL USB ACQUISITION CONTROLLER , IDAC-4</td>
			<td>Ockenfels Syntech GmbH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stimulus Controllers</td>
			<td>Ockenfels Syntech GmbH</td>
			<td>Stimulus Controller CS 55</td>
			<td></td>
		</tr>
		<tr>
			<td>Personal Computer</td>
			<td>Dell</td>
			<td>Vostro</td>
			<td>Check for compatibility with digital acquisition system and software</td>
		</tr>
		<tr>
			<td>Tungsten Rod</td>
			<td>A-M Systems</td>
			<td>Cat#716000</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum Foil and/or Faraday Cage</td>
			<td></td>
			<td></td>
			<td>Electromagnetic noise shielding</td>
		</tr>
		<tr>
			<td>Borosilicate Glass Capillaries</td>
			<td>World Precision Instruments</td>
			<td>1B100F-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Puller</td>
			<td>Sutter Instrument Company</td>
			<td>Model P-97 Flaming/Brown Micropipette Puller</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Olympus</td>
			<td>VMZ 1x-4x</td>
			<td>For fly preparation</td>
		</tr>
		<tr>
			<td>p200 Pipette Tips</td>
			<td></td>
			<td></td>
			<td>Generic</td>
		</tr>
		<tr>
			<td>Microloader tips&#160;</td>
			<td>Eppendorf</td>
			<td>E5242956003</td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml Syringe</td>
			<td></td>
			<td></td>
			<td>Generic</td>
		</tr>
		<tr>
			<td>Crocodile clips</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Power Transformers</td>
			<td>STACO ENERGY PRODUCTS</td>
			<td>STACO 3PN221B</td>
			<td>Assembled from P1000 pipette tips, flexible plastic tubing, and mesh</td>
		</tr>
		<tr>
			<td>Modeling Clay</td>
			<td></td>
			<td></td>
			<td>Generic</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td></td>
			<td></td>
			<td>Generic</td>
		</tr>
		<tr>
			<td>Plastic Tubing</td>
			<td>Saint Gobain</td>
			<td>Tygon S3&#8482; E-3603</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard culture vials</td>
			<td>Archon Scientific</td>
			<td></td>
			<td>Narrow 1-oz polystyrene vails, each with 10 mL of glucose medium, preloaded with cellulose acetate plugs</td>
		</tr>
		<tr>
			<td>Berberine chloride (BER)</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# Y0001149</td>
			<td></td>
		</tr>
		<tr>
			<td>Denatonium benzoate (DEN)</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# D5765</td>
			<td></td>
		</tr>
		<tr>
			<td>N,N-Diethyl-m- toluamide (DEET)</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# 36542</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Figure 4A shows spontaneous spikes that arise from a sensillum. They fall into two classes based on amplitude, with the larger spikes deriving from the neuron that is sensitive to bitter compounds and the smaller spikes from the neuron that responds to sugars. The relationship between spike amplitude and functional specificity has been corroborated by genetic experiments4,14,37,<sup class...

Read this Scientific Article Protocol about Author Spotlight: Advances in Chemoreception at JoVE.com

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.

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 ...

base-recording-technique-for-analyzing-responses-taste-neurons

Research

JoVE Journal

Neuroscience

911 Views.  Yale University. 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. 

Video: Author Spotlight: Advances in Chemoreception – From Insect Odor Receptors to Non-Coding RNAs

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Recording Behavioral Responses to Reflection in Crayfish

Social behavior depends on sensory input from the visual, mechanical and olfactory systems. One important issue concerns the relative roles of each sensory modality in guiding behavior. The role of visual inputs has been examined by isolating visual stimuli from mechanical and chemosensory stimuli. In some studies (Bruski &amp; Dunham, 1987: Delgado-Morales et al., 2004) visual inputs have been removed with blindfolds or low light intensity, and effects of remaining sensory modalities have been elucidated. An alternative approach is to study the effects of visual inputs in the absence of any appropriate mechanical and chemosensory cues. This approach aims to identify the exclusive role of visual inputs. 
We have used two methods to provide visual stimuli to crayfish without providing chemical and mechanical cues. In one method, crayfish are videotaped in an aquarium where half of the walls are covered in mirrors to provide a reflective environment, and the other half are covered in a non-reflective (matte finish) plastic. This gives the crayfish a choice between reflective and non-reflective environments. The reflective environment provides visual cues in the form of reflected images of the crayfish as it moves throughout half of the tank; these visual cues are missing from the non-reflective half of the tank. An alternative method is to videotape the behavior of crayfish in an aquarium separated by a smaller chamber at each end, with a crayfish in one small chamber providing visual cues and an inert object in the opposite small chamber providing visual input from a non-moving, non-crayfish source. 
Our published results indicate that responses of crayfish to the reflective environment depend on socialization and dominance rank. Socialized crayfish spent more time in the reflective environment and exhibited certain behaviors more frequently there than in the non-reflective environment; isolated crayfish showed no such differences. Crayfish that were housed in same-sex pairs developed a social rank of either dominant or subordinate. Responses to reflection differed between dominant and subordinate crayfish (May &amp; Mercier, 2006; May &amp; Mercier, 2007). Dominant crayfish spent more time on the reflective side, entered reflective corners more frequently and spent more time in reflective corners compared to the non-reflective side. Subordinate crayfish walked in reverse more often on the reflective side than on the non-reflective side. Preliminary data suggest similar effects from visual cues provided by a crayfish in a small adjoining chamber (May et al., 2008).

We have developed two methods for studying effects of visual cues on behavior in the absence of tactile and chemical cues. One method involves videotaping responses of crayfish to reflective walls in an aquarium; the other examines effects of visual inputs provided by a live crayfish behind a transparent partition.

Social behavior depends on sensory input from the visual, mechanical and olfactory systems.  One important issue concerns the relative roles of each ...

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 ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Preparation of Drosophila Central Neurons for in situ Patch Clamping

Short generation times and facile genetic techniques make the fruit fly Drosophila melanogaster an excellent genetic model in fundamental neuroscience research. Ion channels are the basis of all behavior since they mediate neuronal excitability. The first voltage gated ion channel cloned was the Drosophila voltage gated potassium channel Shaker1,2. Toward understanding the role of ion channels and membrane excitability for nervous system function it is useful to combine powerful genetic tools available in Drosophila with in situ patch clamp recordings. For many years such recordings have been hampered by the small size of the Drosophila CNS. Furthermore, a robust sheath made of glia and collagen constituted obstacles for patch pipette access to central neurons. Removal of this sheath is a necessary precondition for patch clamp recordings from any neuron in the adult Drosophila CNS. In recent years scientists have been able to conduct in situ patch clamp recordings from neurons in the adult brain3,4 and ventral nerve cord of embryonic5,6, larval7,8,9,10, and adult Drosophila11,12,13,14. A stable giga-seal is the main precondition for a good patch and depends on clean contact of the patch pipette with the cell membrane to avoid leak currents. Therefore, for whole cell in situ patch clamp recordings from adult Drosophila neurons must be cleaned thoroughly. In the first step, the ganglionic sheath has to be treated enzymatically and mechanically removed to make the target cells accessible. In the second step, the cell membrane has to be polished so that no layer of glia, collagen or other material may disturb giga-seal formation. This article describes how to prepare an identified central neuron in the Drosophila ventral nerve cord, the flight motoneuron 5 (MN515), for somatic whole cell patch clamp recordings. Identification and visibility of the neuron is achieved by targeted expression of GFP in MN5. We do not aim to explain the patch clamp technique itself.

Preparation of Drosophila Central Neurons for in situ Patch Clamping

In situ patch clamp recordings are used for electrophysiological characterization of neurons in intact circuitry. In the Drosophila genetic model patch clamping is difficult because the CNS is small and surrounded by a robust sheath. This article describes the procedure to remove the sheath and clean neurons for subsequent patch clamp recordings.

Short generation times and facile genetic techniques make the fruit fly Drosophila melanogaster an excellent genetic model in fundamental neuroscience ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

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 ...

A Low-cost Method for Analyzing Seizure-like Activity and Movement in Drosophila

Video tracking systems have been used widely to analyze Drosophila melanogaster movement and detect various abnormalities in locomotive behavior. While these systems can provide a wealth of behavioral information, the cost and complexity of these systems can be prohibitive for many labs. We have developed a low-cost assay for measuring locomotive behavior and seizure movement in D. melanogaster. The system uses a web-cam to capture images that can be processed using a combination of inexpensive and free software to track the distance moved, the average velocity of movement and the duration of movement during a specified time-span. To demonstrate the utility of this system, we examined a group of D. melanogaster mutants, the Bang-sensitive (BS) paralytics, which are 3-10 times more susceptible to seizure-like activity (SLA) than wild type flies. Using this novel system, we were able to detect that the BS mutant bang senseless (bss) exhibits lower levels of exploratory locomotion in a novel environment than wild type flies. In addition, the system was used to identify that the drug metformin, which is commonly used to treat type II diabetes, reduces the intensity of SLA in the BS mutants.

A Low-cost Method for Analyzing Seizure-like Activity and Movement in Drosophila

Using a webcam and a combination of free and inexpensive software programs, locomotive patterns in Drosophila melanogaster can be analyzed to detect differences in the speed, distance, and time of various motor activities.

Video tracking systems have been used widely to analyze Drosophila melanogaster movement and detect various abnormalities in locomotive behavior. While ...