1. Odor dilutions
<ol>
<li>Most odorants are soluble in paraffin oil. However, DMSO or ethanol can also be used as alternative solvents for particular odors. Prepare appropriate dilutions (e.g. 1:10 volume : volume; v:v) from pure odorants in glass vials. Most odor dilutions are stable at room temperature, but for highly volatile compounds it is better to make working dilutions on a weekly basis. Each sensillum responds to different odors within a different concentration range. For Drosophila, a useful look-up table for appropriate concentrations to use with a given sensillum can be found in 6, 7. 
For the video experiment, we use paraffin oil as a solvent control and methyl acetate (10-6 v:v in paraffin oil) and 1-octen-3-ol (10-7 v:v in paraffin oil) for Drosophila and Anopheles recordings, respectively.</li>
<li>Using scissors, cut the chromatography paper into 3 mm x 5 cm strips so that they will fit inside Pasteur pipettes.</li>
<li>Pipette 30 &mu;L of the desired odor on a filter paper strip and insert it into the glass pipette. Cut ~3 cm of air line tubing and insert it into the open end of the pipette, closing it with a connector. The connector is used to seal the pipette that will be then attached to the air-line tubing of an air pump when it is time to deliver the odor during the experiment.</li>
</ol>
2. Odor delivery system
<ol>
<li>Using a small drill, cut a 10 mL plastic serological pipette (e.g. at the 4 mL mark) and create two holes (e.g. at the -1.5 mL and -0.5 mL marks) that will be used to hold the pipettes containing the odors. Insert a 200 &mu;L pipette tip in the barbed coupler and introduce the coupler into the blunt end of the 10 mL pipette. The pipette will be used as part of the delivery system for odorants.</li>
<li>Attach the pipette on a magnetic stand with a pipette clamp and position it near the microscope.</li>
</ol>
3. Sharpening electrodes
<ol>
<li>To sharpen electrodes, prepare a 0.5 M solution of potassium hydroxide (KOH) and filter it to remove fine particles (e.g. using a 45 &mu;m filter). Take a 20 mL syringe and make a small hole (~2 mm diameter) using a needle on the wall close to the tip (~1 cm from the tip), into which the electric wire is inserted (Figure 1A).</li>
<li>Fill the syringe with 0.5 M KOH, and clamp it on a stand under the microscope so that the tip is placed in the field of view (Figure 1B,
C). Insert the electric wire into the small hole on the syringe wall (Figure 1B), making sure that the wire is not directly in front of the
entrance of the syringe. Connect this wire to the cathode of a DC power supply (e.g. Wild Heerbrugg MTR32, see methods) or a pole of an AC power supply (e.g. Wild Heerbrugg-LEP 990018). Attach the electrode holder shaft onto the manual micromanipulator on the right side of the microscope, and attach an electric cable at the base of the electrode holder shaft with a crocodile clip. Connect the cable to the anode of the DC power supply or the other pole of the AC power supply (Figure 1A).</li>
<li>Insert the tungsten wire (~5 cm length) into the electrode holder, and attach it to the electrode holder shaft on the manipulator. Set the power supply to 6 V, and insert the tip of the wire into the syringe repeatedly to hone it, being careful to monitor the tip under the microscope during the process (Figure 1C). To get an ideal electrode for recording, place 90% of its length in the solution for up to 1 min, and pull it out slowly. Then insert only ~50% of the electrode to further thin for 30 s, and repeat it to get the tip of the wire sharpened (~10 times). The tip of the electrode should be fine enough to enter into the sensillum, but not so fine as to bend when it touches it during the recordings (steps 6 and 7). Although watching the electrode tip under the dissecting microscope while it is being sharpened is a good indication of its thickness, only looking at it under the recording microscope at high magnification will give a clear idea whether a given electrode is suitable for recording.</li>
</ol>
4. Insect prep: Drosophila antenna
<ol>
<li>Build a fly aspirator. Cut a piece of air line tubing long enough to hang comfortably around your neck (about 90 - 120 cm). Cut the tip of a 200 &mu;L pipette tip and insert it into one end of the tube. On the other end, position a ~1.5 cm x 1.5 cm piece of mesh so that it creates a physical barrier but does not prevent air from flowing out of the tube. Cut the tip of a 1 mL pipette tip and position the wider end on top of the tube opening, blocking the mesh in between. This portion will be used to pick up and manipulate adult vinegar flies (Figure 2).</li>
<li>Take a microscope slide and place a piece of dental wax roughly in the middle of the long side. On top of it position a cover glass slightly tilted upwards (~30 ), making sure that the wax is not directly underneath the uppermost part, which would prevent visualization of the vinegar fly under the microscope. Pull a glass electrode with the vertical puller. Its tip should be thin and flexible enough to fit in between the second and third antennal segment, and keep the antenna stable for the recordings. Position the glass electrode on another piece of wax and place it on the side of the cover glass, far enough that when the tip is lowered it reaches the corner of the cover glass (Figure 3A).</li>
<li>Working from a bottle or vial of adult flies of the desired genotype, pick up an adult vinegar fly using the fly aspirator. Although females are usually used because of their bigger size, males can also be used. Place a 200 &mu;L pipette tip on top of the 1 mL tip to avoid the fly from escaping. Blow into the tube, so that the fly is pushed towards the end of the 200 &mu;L pipette tip. Trim the wide end of the pipette tip with the razor blade a few millimeters away from the fly itself, then insert some wax to prevent the fly from backing out. Under a microscope, cut again near the head of the vinegar fly, paying attention not to damage the animal. With a small pipette tip, push the wax to force the fly head out so that about half of the eyes extrude from the tip (Figure 3B). Make sure that its legs do not come out as well, or they might move and interfere with the recordings.</li>
<li>Mount the fly on a piece of wax, its head facing upwards, and place it on the slide in front of the cover glass. Push the head slightly against the corner of the glass, so that the antennae extend and rest on the glass. Lower the tip of the glass capillary in between the second and third antennal segment (Figure 3B).</li>
<li>Different parts of the antenna will give access to different sensilla types. Depending on particular experimental needs, the antenna will need to be oriented differently to allow access to different sensilla types. To record from large basiconic sensilla as in our example, the arista is pushed down on the glass (Figure 3B).</li>
</ol>
5. Insect prep: Anopheles maxillary palps
<ol>
<li>Using an electric aspirator (Figure 2E), collect 40 to 60 3-5 days old mosquitoes (mixed males and females) in the small plastic cage (Figure 2F). Mosquitoes should have been reared under normal conditions, i.e. in an insect incubator or insectary at 25-28&deg;C with 70-80% humidity. Place the small plastic cage with the animals on ice for ~15 min until they are anesthetized by cold. Once the animals have stopped moving, transfer only 4-6 animals to the stage under the microscope at any one time, keeping the rest of the animals on ice during the procedures. Although females are usually used because of their sensitivity to CO2, males can also be employed. Select female or male mosquitoes, judging by the structure of their antenna (feather-like in females, filament-like in males), and remove their wings and legs with fine forceps to immobilize them. Keep them in a small plastic cup with a wet paper at the bottom, to prevent them from desiccating. </li>
<li>Put two pieces of double-sided tape (~1 cm in length) parallel to each other (~1 cm apart), at the center and at the side of the slide glass (Figure 3C). Using fine forceps, place one mosquito on the central tape under the dissecting microscope, and turn it sideways and stick its body and one eye on the tape.&nbsp; Adjust the position of the maxillary palps so that both palps extend parallel on the tape (Figure 3D). Fix the maxillary palps by placing thin strings (e.g. human hairs) both at the base and at the tip of palps (Figure 3D). The side tape is used as a repository for thin strings, while the central tape needs to be replaced after every recording (Figure 3C).</li>
</ol>
6. Recording Drosophila melanogaster
<ol>
<li>Place the slide under the microscope (Figure 4A) at low magnification and position the antenna roughly in the middle of the field of view (FOV) (Figure 4B). Gently lower the electrodes so that the reference electrode is located near the eye of the fly and the recording electrode is near the antenna (Figure 4C). Increase the magnification and re-position the antenna in the middle of the FOV (Figure 4D).</li>
<li>Place the odor delivery device close to the head of the fly, pointing at the antenna.</li>
<li>At low magnification (Figure 4C), insert the reference electrode into the eye of the fly. Lower the recording electrode on top of the antenna without touching its surface.</li>
<li>Switching to high magnification, control the recording electrode with the micromanipulator and insert it into a selected sensillum (Figure 4D). Any point along the sensillum length is good for recording. Once inside the sensillum, the electrode can be pushed farther in (sometimes all the way through) to obtain a better signal to noise ratio. Once the electrode is in the sensillum, the spontaneous activity of the cells can be detected.</li>
</ol>
7. Recording Anopheles gambiae
<ol>
<li>Place the glass slide under the microscope at low magnification (10x) and position the maxillary palps roughly in the middle of the FOV and the head at the top (Figure 4E). Rotate the stage until one of the palps is at a right angle with the recording electrode.</li>
<li>Adjust the height of the electrodes so that the reference electrode is positioned just above the eye of the mosquito and the recording electrode is near the maxillary palps.</li>
<li>Place the odor delivery device so that it is as close as possible to the palps.</li>
<li>Insert the reference electrode into the eye at lower magnification (10x), and switching to the higher magnification (100x), insert the recording electrode into the peg sensillum on the palp (Figure 4F). Once the electrode is in the sensillum, the spontaneous activity of the cells can be detected.</li>
</ol>
8. Representative results
Depending on the sensillum and the quality of the recording, one can distinguish different numbers of olfactory neurons within a single trace. In the large basiconic sensilla of Drosophila melanogaster, for example, between 2 and 4 cells that differ in spike amplitude should appear during the recording 9, 10.
In our video experiment, the Drosophila ab2 sensillum shows two cells, an A cell (Figure 5, blue spikes) and a B cell (Figure 5, green spikes). Neither cell is activated during application of paraffin oil (Figure 5A), while only the A cell responds to the 10-6 dilution of methyl acetate (Figure 5B).
In the maxillary palp of Anopheles gambiae, the grooved peg sensillum contains three cells, but only two are easily discriminated (Figure 5C, blue and green spikes, respectively). In the video experiment we show how the B cell responds to a 10-7 dilution of 1-octen-3-ol (Figure 5D).
<img src="/files/ftp_upload/1725/1725fig1.jpg" alt="Figure 1" /> Figure 1. Electrode sharpener (A) General view of the electrode sharpener apparatus. (B) The syringe containing 0.5 M KOH (left) used to sharpen the electrode (right). (C) Close-up of the electrode tip next to the opening of the syringe.
<img src="/files/ftp_upload/1725/1725fig2.jpg" alt="Figure 2" /> Figure 2. How to prepare a fly aspirator and mosquito aspirator (A) Starting material: air line plastic tubing, two cut pipette tips, and mesh. (B) The fly aspirator once it is completed. (C) Detail of the end that is used to catch vinegar flies. (D) Detail of the other end of the fly aspirator. (E) The electric aspirator for mosquito collection consists of a main body and a detachable plastic cage. (F) The detachable plastic cage for mosquitoes.
<img src="/files/ftp_upload/1725/1725fig3.jpg" alt="Figure 3" /> Figure 3. Preparing a vinegar fly (Drosophila melanogaster) and a malaria mosquito (Anopheles gambiae) for recording (A) Picture of a vinegar fly mounted on the slide before positioning it under the microscope. (B) Close-up of the vinegar fly head with the antenna kept in place by the glass capillary. (C) Picture of a mosquito mounted on the slide. (D) Close-up of the mosquito head with proboscis and palps sticking on the tape.
<img src="/files/ftp_upload/1725/1725fig4.jpg" alt="Figure 4" /> Figure 4. Recording from a vinegar fly (Drosophila melanogaster) and a malaria mosquito (Anopheles gambiae) (A) View of the electrophysiology setup. (B) Close-up of the fly preparation mounted on the microscope. Notice the respective position of the recording electrode (left), the odor delivery system (middle pipette), and the recording electrode (right). (C) Image of the fly under the 10x objective. (D) Image of the fly antenna under the 100x objective; big basiconic sensilla (arrows), interspersed among non-sensory hairs (arrowheads). (E) 10x view of a mosquito mounted for recording. (F) High magnification view of the mosquito palp and a peg sensillum (arrow).
<img src="/files/ftp_upload/1725/1725fig5.jpg" alt="Figure 5" /> Figure 5. Examples of recordings from Drosophila melanogaster and Anopheles gambiae 
(A) The ab2 sensillum of Drosophila melanogaster houses two sensory neurons; the A cell (blue spikes) and B cell (green spikes). (B) The A and B cells during application of 10-6 methyl acetate. (C) The peg sensillum of Anopheles gambiae houses two sensory neurons; the A cell (blue spikes) and B cell (green spikes). (D) Application of 10-7 1-octen-3-ol to the peg sensillum.

<ol>
	<li>Boeckh, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elektrophysiologische+Untersuchungen+an+einzelnen+Geruchsrezeptoren+auf+der+Antenne+des+TotengrAbers+(Necrophorus+Coleoptera).">Elektrophysiologische Untersuchungen an einzelnen Geruchsrezeptoren auf der Antenne des TotengrAbers (Necrophorus Coleoptera).</a> Z. Vergl. Physiol. 46, 212-248 (1962).</li><li>Schneider, D., Hecker, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zur+Elektrophysiologic+der+Antenne+des+Seidenspinners+Bombyx+mori+bei+Reizung+mit+angereicherten+Extrakten+des+Sexuallockstoffes.">Zur Elektrophysiologic der Antenne des Seidenspinners Bombyx mori bei Reizung mit angereicherten Extrakten des Sexuallockstoffes.</a> Z. Naturforschg. 11b, 121-124 (1956).</li><li>Touhara, K., Vosshall, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensing+odorants+and+pheromones+with+chemosensory+receptors.">Sensing odorants and pheromones with chemosensory receptors.</a> Annu Rev Physiol. 71, 307-332 (2009).</li><li>Sato, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insect+olfactory+receptors+are+heteromeric+ligand-gated+ion+channels.">Insect olfactory receptors are heteromeric ligand-gated ion channels.</a> Nature. 452, 1002-1006 (2008).</li><li>Wicher, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+odorant+receptors+are+both+ligand-gated+and+cyclic-nucleotide-activated+cation+channels.">Drosophila odorant receptors are both ligand-gated and cyclic-nucleotide-activated cation channels.</a> Nature. 452, 1007-1011 (2008).</li><li>Hallem, E. A., Ho, M. 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=The+molecular+basis+of+odor+coding+in+the+Drosophila+antenna.">The molecular basis of odor coding in the Drosophila antenna.</a> Cell. 117, 965-979 (2004).</li><li>Hallem, E. A., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coding+of+odors+by+a+receptor+repertoire.">Coding of odors by a receptor repertoire.</a> Cell. 125, 143-160 (2006).</li><li>Boeckh, J., Kaissling, K. E., Schneider, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insect+olfactory+receptors.">Insect olfactory receptors.</a> Cold Spring Harb Symp Quant Biol. 30, 263-280 (1965).</li><li>de Bruyne, M., Foster, K., 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=Odor+coding+in+the+Drosophila+antenna.">Odor coding in the Drosophila antenna.</a> Neuron. 30, 537-552 (2001).</li><li>Lu, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odor+coding+in+the+maxillary+palp+of+the+malaria+vector+mosquito+Anopheles+gambiae.">Odor coding in the maxillary palp of the malaria vector mosquito Anopheles gambiae.</a> Curr Biol. 17, 1533-1544 (2007).</li><li>Hallem, E. A., Fox, A. N., Zwiebel, L. J., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Olfaction:+mosquito+receptor+for+human-sweat+odorant.">Olfaction: mosquito receptor for human-sweat odorant.</a> Nature. 427, 212-213 (2004).</li><li>Couto, A., Alenius, M., Dickson, B. J. M. o. l. e. c. u. l. a. r. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=anatomical,+and+functional+organization+of+the+Drosophila+olfactory+system.">anatomical, and functional organization of the Drosophila olfactory system.</a> Curr. Biol. 15, 1535-1547 (2005).</li><li>Fishilevich, E., Vosshall, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+and+functional+subdivision+of+the+Drosophila+antennal+lobe.">Genetic and functional subdivision of the Drosophila antennal lobe.</a> Curr Biol. 15, 1548-1553 (2005).</li></ol>

Olfactory cues are used by organisms to identify food sources, potential mates, and predators. Olfactory sensory neurons (OSNs) are the first relay center between external stimuli and higher centers of the brain where the information is further processed. In Drosophila melanogaster and Anopheles gambiae, OSNs are easily accessible and their electrical activity can be monitored while stimulated by odor puffs.
The single sensillum recording (SSR) technique explained in this vi...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Paraffin oil</td>
<td>Odors</td>
<td>Fluka</td>
<td>76235</td>
<td>&nbsp;</td>
<tr>
<td>High purity odors (&gt;98%)</td>
<td>Odors</td>
<td>Sigma-Aldrich</td>
<td>&nbsp;</td>
<td>Methyl acetate 
#296996 
1-octen-3-ol 
#74950 </td>
</tr>
<tr>
<td>Filter paper strips</td>
<td>Odors</td>
<td>Fisherbrand</td>
<td>05-714-1</td>
<td>Chromatography paper</td>
</tr>
<tr>
<td>Connectors</td>
<td>Odors</td>
<td>Cole-Parmer</td>
<td>EW-06365-40</td>
<td>1/16x1/8"</td>
</tr>
<tr>
<td>Glass vials</td>
<td>Odors</td>
<td>Agilent Technologies</td>
<td>5182-0556</td>
<td>&nbsp;</td>
<tr>
<td>Air line plastic tubing</td>
<td>Odor Delivery</td>
<td>Python Products</td>
<td>500PAL</td>
<td>&nbsp;</td>
<tr>
<td>1 serological pipette</td>
<td>Odor Delivery</td>
<td>Corning</td>
<td>4101</td>
<td>10 mL</td>
</tr>
<tr>
<td>Plastic tubing</td>
<td>Odor Delivery</td>
<td>Cole-Parmer</td>
<td>EW-06418-0</td>
<td>0.050"x0.090"OD</td>
</tr>
<tr>
<td>Disposable borosilicate glass Pasteur pipettes</td>
<td>Odor Delivery</td>
<td>FisherBrand</td>
<td>13-678-20A</td>
<td>5-3/4 inches</td>
</tr>
<tr>
<td>Programmable stimulus controller</td>
<td>Odor Delivery</td>
<td>Syntech</td>
<td>CS-55</td>
<td>&nbsp;</td>
<tr>
<td>Anti-vibration table</td>
<td>Electrophysiology Equipment</td>
<td>TMC</td>
<td>63533</td>
<td>36&#x201D;Wx30&#x201D;Dx29&#x201D;H</td>
</tr>
<tr>
<td>Faraday cage</td>
<td>Electrophysiology Equipment</td>
<td>TMC</td>
<td>MI8133303</td>
<td>&nbsp;</td>
<tr>
<td>Inverted microscope</td>
<td>Electrophysiology Equipment</td>
<td>Nikon</td>
<td>E600FN ECLIPSE</td>
<td>Recording microscope</td>
</tr>
<tr>
<td>10x and 100x objectives</td>
<td>Electrophysiology Equipment</td>
<td>Nikon</td>
<td>10x Plan Fluor 100x L Plan</td>
<td>&nbsp;</td>
<tr>
<td>Dissecting microscope</td>
<td>Electrophysiology Equipment</td>
<td>Nikon</td>
<td>EZ645</td>
<td>electrode sharpening/insect prep microscope</td>
</tr>
<tr>
<td>Magnetic stands</td>
<td>Electrophysiology Equipment</td>
<td>Newport</td>
<td>MODEL 150</td>
<td>&nbsp;</td>
<tr>
<td>IDAC</td>
<td>Electrophysiology Equipment</td>
<td>Syntech</td>
<td>IDAC-4</td>
<td>&nbsp;</td>
<tr>
<td>Acquisition software</td>
<td>Electrophysiology Equipment</td>
<td>Syntech</td>
<td>Autospike</td>
<td>&nbsp;</td>
<tr>
<td>1 macromanipulator</td>
<td>Electrophysiology Equipment</td>
<td>NARISHIGE</td>
<td>MN-151</td>
<td>Joystick manipulator 
Used for positioning reference electrode
</td>
</tr>
<tr>
<td>1 micromanipulator</td>
<td>Electrophysiology Equipment</td>
<td>EXFO</td>
<td>PCS-6000</td>
<td>Used for positioning recording electrode</td>
</tr>
<tr>
<td>Crocodile clip</td>
<td>Electrophysiology Equipment</td>
<td>Pomona</td>
<td>AL-B-12-0</td>
<td>&nbsp;</td>
<tr>
<td>Electric cable</td>
<td>Electrophysiology Equipment</td>
<td>Pomona</td>
<td>B-36-0</td>
<td>Test Cable Assembly</td>
</tr>
<tr>
<td>2 electrode holders</td>
<td>Electrophysiology Equipment</td>
<td>Syntech</td>
<td>N/A</td>
<td>Electrode holders (set of 2) for tungsten wire electrode</td>
</tr>
<tr>
<td>AC probe</td>
<td>Electrophysiology Equipment</td>
<td>Syntech</td>
<td>N/A</td>
<td>Universal single ended probe (10xAC)</td>
</tr>
<tr>
<td>Tungsten electrodes</td>
<td>Electrophysiology Equipment</td>
<td>Microprobes</td>
<td>M210</td>
<td>straight tungsten rods, 0.005&#x201C;x3&#x201C;</td>
</tr>
<tr>
<td>Potassium hydroxide</td>
<td>Electrophysiology Equipment</td>
<td>Sigma-Aldrich</td>
<td>221473</td>
<td>&nbsp;</td>
<tr>
<td>Syringe</td>
<td>Electrophysiology Equipment</td>
<td>BD</td>
<td>301625</td>
<td>20 mL</td>
</tr>
<tr>
<td>Power supply</td>
<td>Electrophysiology Equipment</td>
<td>WILD HEERBRUGG 6V 40W</td>
<td>e.g MTR32</td>
<td>&nbsp;</td>
<tr>
<td>Vertical puller</td>
<td>Insect prep</td>
<td>Narishige</td>
<td>PB-7</td>
<td>&nbsp;</td>
<tr>
<td>Razor blade</td>
<td>Insect prep</td>
<td>VWR</td>
<td>55411-050</td>
<td>&nbsp;</td>
<tr>
<td>Dental wax</td>
<td>Insect prep</td>
<td>Patterson</td>
<td>091-1503</td>
<td>&nbsp;</td>
<tr>
<td>Microscope slide</td>
<td>Insect prep</td>
<td>FisherBrand</td>
<td>12-550A</td>
<td>&nbsp;</td>
<tr>
<td>Cover glass</td>
<td>Insect prep</td>
<td>FisherBrand</td>
<td>12-541A</td>
<td>18X18 #1.5</td>
</tr>
<tr>
<td>Polypropylene mesh</td>
<td>Insect prep</td>
<td>Small Parts inc.</td>
<td>CMP-0500-B</td>
<td>&nbsp;</td>
<tr>
<td>Glass electrode</td>
<td>Insect prep</td>
<td>Frederick Haer & Co.</td>
<td>27-32-0-075</td>
<td>Capillary tubing borosilicate 1.5mm OD x 1.12mm ID x 75 mm</td>
</tr>
<tr>
<td>Double-sided tape (3M)</td>
<td>Insect prep</td>
<td>3M</td>
<td>MMM6652P3436</td>
<td>Double-sided tape (3M)</td>
</tr>
<tr>
<td>Forceps</td>
<td>Insect prep</td>
<td>Fine Science Tools</td>
<td>021x0053</td>
<td>Dumont #5 Mirror Finish Forceps</td>
</tr>
<tr>
<td>Small plastic cup</td>
<td>Insect prep</td>
<td>VWR</td>
<td>89009-662</td>
<td>7 x 5.7 (23/4 x 21/4)</td>
</tr>
<tr>
<td>Electric aspirator</td>
<td>Insect prep</td>
<td>Gempler&#x2019;s</td>
<td>RHM200</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

single sensillum recordings in the insects drosophila melanogaster and anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Watch this Scientific Journal Video about Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae at JoVE.com

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

single-sensillum-recordings-insects-drosophila-melanogaster-anopheles

Research

JoVE Journal

Neuroscience

25.0K Views.  Rockefeller University. Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

Video: Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" (or "startle") response during the adult stage is mediated by the multi-component neuronal circuit called the Giant Fiber System (GFS). The comparative large size of the neurons, their distinctive morphology and simple connectivity make the GFS an attractive model system for studying neuronal circuitry. The GFS pathway is composed of two bilaterally symmetrical Giant Fiber (GF) interneurons whose axons descend from the brain along the midline into the thoracic ganglion via the cervical connective. In the mesothoracic neuromere (T2) of the ventral ganglia the GFs form electro-chemical synapses with 1) the large medial dendrite of the ipsilateral motorneuron (TTMn) which drives the tergotrochanteral muscle (TTM), the main extensor for the mesothoracic femur/leg, and 2) the contralateral peripherally synapsing interneuron (PSI) which in turn forms chemical (cholinergic) synapses with the motorneurons (DLMns) of the dorsal longitudinal muscles (DLMs), the wing depressors. The neuronal pathway(s) to the dorsovental muscles (DVMs), the wing elevators, has not yet been worked out (the DLMs and DVMs are known jointly as indirect flight muscles - they are not attached directly to the wings, but rather move the wings indirectly by distorting the nearby thoracic cuticle) (King and Wyman, 1980; Allen et al., 2006). The di-synaptic activation of the DLMs (via PSI) causes a small but important delay in the timing of the contraction of these muscles relative to the monosynaptic activation of TTM (~0.5 ms) allowing the TTMs to first extend the femur and propel the fly off the ground. The TTMs simultaneously stretch-activate the DLMs which in turn mutually stretch-activate the DVMs for the duration of the flight. The GF pathway can be activated either indirectly by applying a sensory (e.g."air-puff" or "lights-off") stimulus, or directly by a supra-threshold electrical stimulus to the brain (described here). In both cases, an action potential reaches the TTMs and DLMs solely via the GFs, PSIs, and TTM/DLM motoneurons, although the TTMns and DLMns do have other, as yet unidentified, sensory inputs. Measuring "latency response" (the time between the stimulation and muscle depolarization) and the "following to high frequency stimulation" (the number of successful responses to a certain number of high frequency stimuli) provides a way to reproducibly and quantitatively assess the functional status of the GFS components, including both central synapses (GF-TTMn, GF-PSI, PSI-DLMn) and the chemical (glutamatergic) neuromuscular junctions (TTMn-TTM and DLMn-DLM). It has been used to identify genes involved in central synapse formation and to assess CNS function.

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

The Giant Fiber System is a simple neuronal circuit of adult Drosophila melanogaster containing the largest neurons in the fly. We describe the protocol for monitoring synaptic transmission through this pathway by recording post synaptic potentials in dorsal longitudinal (DLM) and tergotrochanteral (TTM) muscles following direct stimulation of the Giant Fiber interneurons.

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

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

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

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

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

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such behavior is spontaneous locomotor activity. Here we describe our protocol that utilizes Drosophila population monitors and a tracking system that allows continuous monitoring of the spontaneous locomotor activity of flies for several days at a time. This method is simple, reliable, and objective and can be used to examine the effects of aging, sex, changes in caloric content of food, addition of drugs, or genetic manipulations that mimic human diseases.

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster are useful in studying genetic or environmental manipulations that affect behaviors such as spontaneous locomotor activity.&#160; Here we describe a protocol that utilizes monitors with infrared beams and data analysis software to quantify spontaneous locomotor activity.&#160;

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such ...

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy and cheap methods to distinguish between individuals in a trial. A standard technique for this is CO2 anaesthesia and then labelling or wing clipping each fly. However, this is invasive and has been shown to affect behavior. Other techniques have used coloration to identify flies. This article presents a simple and non-invasive method for labelling Drosophila that allows them to be individually identified within experiments, using food coloring. This method is used in trials where two males compete to mate with a female. Dyeing allowed quick and easy identification. There was, however, some difference in the strength of the coloration across the three species tested. Data is presented showing the dye has a lower impact on mating behavior than CO2 in Drosophila melanogaster. The impact of CO2 anaesthesia is shown to depend on the species of Drosophila, with D. pseudoobscura and D. subobscura showing no impact, whereas D. melanogaster males had reduced mating success. The dye method presented is applicable to a wide range of experimental designs.

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

This protocol describes a simple, cost effective way to individually identify Drosophila or other insects. Demonstration data investigating mating success across three species of Drosophila show that this method is comparable or better than the use of CO2 anaesthesia.

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy ...

Conditions Affecting Social Space in Drosophila melanogaster

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a straightforward manner. As we previously demonstrated 1, in a two-dimensional chamber, we first force the flies to form a tight group, subsequently allowing them to take their preferred distance from each other. After the flies have settled, we measure the distance to the closest neighbor (or social space), processing a static picture with free online software (ImageJ). The analysis of the distance to the closest neighbor allows researchers to determine the effects of genetic and environmental factors on social interaction, while controlling for potential confounding factors. Diverse factors such as climbing ability, time of day, sex, and number of flies, can modify social spacing of flies. We thus propose a series of experimental controls to mitigate these confounding effects. This assay can be used for at least two purposes. First, researchers can determine how their favorite environmental shift (such as isolation, temperature, stress or toxins) will impact social spacing 1,2. Second, researchers can dissect the genetic and neural underpinnings of this basic form of social behavior 1,3. Specifically, we used it as a diagnostic tool to study the role of orthologous genes thought to be involved in social behavior in other organisms, such as candidate genes for autism in humans 4.

Conditions Affecting Social Space in Drosophila melanogaster

The effect of genes and environment on social space of Drosophila melanogaster can be quantified through a powerful but straightforward analytical paradigm. We show here different factors that can affect this social space, and thus need to be taken into consideration when designing experiments in this paradigm.

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a ...

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense organs provide an excellent model system for the study of mechanosensation. This protocol describes a simple touch behavior in fruit flies which can be used to identify mutations that interfere with mechanosensation. The tactile stimulation of a macrochaete bristle on the thorax of flies elicits a grooming reflex from either the first or third leg. Mutations that interfere with mechanotransduction (such as NOMPC), or with other aspects of the reflex arc, can inhibit the grooming response. A traditional screen of adult behaviors would have missed mutants that have essential roles during development. Instead, this protocol combines the touch screen with mosaic analysis with a repressible cell marker (MARCM) to allow for only limited regions of homozygous mutant cells to be generated and marked by the expression of green fluorescent protein (GFP). By testing MARCM clones for abnormal behavioral responses, it is possible to screen a collection of lethal p-element mutations to search for new genes involved in mechanosensation that would have been missed by more traditional methods.

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

In order to identify novel mutations affecting mechanosensation, we designed an assay that measures the behavioral response to tactile stimulation of fly bristles in mutant clones generated by the MARCM method. The combination of techniques allows for the identification of mechanosensitive mutations that would otherwise be lethal.

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

For most animals, feeding is an essential behavior for securing survival, and it influences development, locomotion, health and reproduction. Ingestion of the right type and quantity of food therefore has a major influence on quality of life. Research on feeding behavior focuses on the underlying processes that ensure actual feeding and unravels the role of factors regulating internal energy homeostasis and the neuronal bases of decision-making. The model organism Drosophila melanogaster, with its great variety of genetically traceable tools for labeling and manipulating single neurons, allows mapping of neuronal networks and identification of molecular signaling cascades involved in the regulation of food intake. This report demonstrates the CApillary FEeder assay (CAFE) and shows how to measure food intake in a group of flies for time spans ranging from hours to days. This easy-to-use assay consists of glass capillaries filled with liquid food that flies can freely access and feed on. Food consumption in the assay is accurately determined using simple measurement tools. Herein we describe step-by-step the method from setup to successful execution of the CAFE assay, and provide practical examples to analyze the food intake of a group of flies under controlled conditions. The reader is guided through possible limitations of the assay, and advantages and disadvantages of the method compared to other feeding assays in D. melanogaster are evaluated.

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

The CApillary FEeder (CAFE) assay is a simple, budget-friendly, highly reliable method for investigating mechanisms underlying food intake. Used with the highly versatile genetic model organism Drosophila melanogaster, it provides a powerful means of gaining new insights into regulatory mechanisms of food intake.

For most animals, feeding is an essential behavior for securing survival, and it influences development, locomotion, health and reproduction. Ingestion of ...

Measuring and Altering Mating Drive in Male Drosophila melanogaster

Despite decades of investigation, the neuronal and molecular bases of motivational states remain mysterious. We have recently developed a novel, reductionist, and scalable system for in-depth investigation of motivation using the mating drive of male Drosophila&#160;melanogaster (Drosophila), the methods for which we detail here. The behavioral paradigm centers on the finding that male mating drive decreases alongside fertility over the course of repeated copulations and recovers over ~3 d. In this system, the powerful neurogenetic tools available in the fly converge with the genetic accessibility and putative wiring diagram available for sexual behavior. This convergence allows rapid isolation and interrogation of small neuronal populations with specific motivational functions. Here we detail the design and execution of the satiety assay that is used to measure and alter courtship motivation in the male fly. Using this assay, we also demonstrate that low male mating drive can be overcome by stimulating dopaminergic neurons. The satiety assay is simple, affordable, and robust to influences of genetic background. We expect the satiety assay to generate many new insights into the neurobiology of motivational states.

Measuring and Altering Mating Drive in Male Drosophila melanogaster

This article describes a behavioral assay that uses male mating drive in Drosophila melanogaster to study motivation. Using this method, researchers can utilize advanced fly neurogenetic techniques to uncover the genetic, molecular, and cellular mechanisms that underlie this motivation.