I am grateful to Dr. Cl&#233;ment Vinauger and Dr. Jeffrey Riffell for helpful discussions. The following reagents were obtained through BEI Resources, NIAID, NIH: Anopheles stephensi, Strain STE2, MRA-128, contributed by Mark Q. Benedict; Aedes aegypti, Strain ROCK, MRA-734, contributed by David W. Severson; Culex quinquefasciatus, Strain JHB, Eggs, NR-43025. The author thanks Dr. Jake Tu, Dr. Nisha Duggal, Dr. James Weger and Jeffrey Marano for providing Culex quinquefasciatus and Anopheles stephensi (strain: Liston) mosquito eggs. Aedes albopictus and Toxorhynchites rutilus septentrionalis are derived from field mosquitoes collected by the author in the New River Valley area (VA, USA). This work was supported by The Department of Biochemistry and The Fralin Life Science Institute.

1. Saline solution preparation
<ol>
	<li>Prepare the saline in advance and store in the fridge.</li>
	<li>Follow Beyenbach and Masia26 to prepare the solution. 
	&#8203;NOTE: Saline recipe in mM: 150.0 NaCl, 25.0 HEPES, 5.0 glucose, 3.4 KCl, 1.8 NaHCO3, 1.7 CaCl2, and 1.0 MgCl2. The pH is adjusted to 7.1 with 1 M NaOH. Do not add glucose or sucrose to the preparation at this time to increase shelf storage. Add the needed quantity to the saline right before running the EAGs (around 50 mL per experiment).</li>
</ol>
2. Odor preparation and storage
<ol>
	<li>Prepare the odorant mixtures or single compound dilutions in advance in 1.5 mL amber vials and store at -20 &#176;C to prevent compound degradation. 
	NOTE: Concentrations will depend on the test to be conducted. 0.1% or 1% are commonly used to determine whether a compound can be detected or not. For a dose-response curve, prepare serial dilutions of a given chemical and test them from that the lowest to the highest concentrations.</li>
	<li>Prepare the dilutions in water, ethanol, hexane, paraffin oil, or mineral oil, depending on the solubility of the tested chemical.</li>
	<li>Make sure to prepare a solvent control (a vial containing only the solvent) for the experiment.</li>
	<li>Remove the odorants from the freezer 30 min prior to starting the experiments to allow them to thaw. Vortex each vial prior to use to mix well the chemical and the solvent.</li>
	<li>Pipette 10 &#181;L of solution on to a piece of filter paper (0.5 cm x 2 cm) loaded inside a labeled glass syringe or Pasteur pipette.</li>
	<li>Load each compound or mixture in a specific Pasteur pipette or syringe to prevent contamination. 
	NOTE: Load 10 min before starting the experiment so the odor can diffuse in the syringe but not for longer to prevent degradation. Let the Pasteur pipette or syringe remain capped at this time to allow a good diffusion of the chemical before the experiment starts.</li>
	<li>After each EAG run, dispose the piece of the filter paper and replace it with a new one to prevent the paper from getting oversoaked and risking needle blockage. Replace the needles regularly (every 10 runs).</li>
</ol>
3. Mosquito separation
<ol>
	<li>Isolate the mosquitoes on the day of the experiments.</li>
	<li>Use mosquitoes that are at least 6 days old on the day of the experiments to increase the chances that the females are mated to enhance their response to host-related odorants. 
	NOTE: Adjust mosquito age at the time of the test depending on the project. Check and harmonize the physiological status (e.g., blood-fed, starved, never previously fed, etc.).</li>
	<li>Starve the mosquitoes up to 12 h (i.e., no access to sugar) to increase their motivation and sensitivity.</li>
	<li>Place the mosquito container in the fridge (4 &#176;C) until they stop flying so individuals can easily be delicately transferred to single cups with forceps. 
	NOTE: Species with a higher tolerance to cold can be put down using a CO2 fly pad. Ensure that the mosquitoes do not stay on it for a long time to prevent desiccation, which would decrease the responsiveness of the mosquito EAG preparation.</li>
	<li>Store the cups containing single mosquitoes at room temperature before the EAGs are performed and discard any mosquitoes that may not be used during the day.</li>
</ol>
4. Electrode holder and capillary preparation
<ol>
	<li>Capillary pulling, preparation, and storage
	<ol>
		<li>Use borosilicate capillaries with filaments (I.D: 0.78 mm, O.D: 1 mm). Pull them depending on the equipment27. 
		NOTE: Store the pulled capillaries in a Petri dish. Place the Petri dish on to pieces of wax or unscented modeling clay to prevent them from moving and breaking.</li>
		<li>Before running the EAG experiment, gently break the tip of 2 capillaries with a pair of forceps under the microscope. 
		NOTE: Ensure that one is slightly bigger than the other to fit either the neck (larger capillary) or the tips of the antennae (smaller capillary). Ensure that the cut is clean with no crack present on the capillary wall. This requires patience and practice.</li>
		<li>If still intact, reuse these capillaries after rinsing with de-ionized (DI) water after the experiment is over. Remove the excess water by gently applying a cleaning wipe against the tip. Place back in storage Petri dish. If the tip is crooked, discard the capillary.</li>
	</ol>
	</li>
	<li>Electrode holder and capillary mounting
	<ol>
		<li>Label the two electrode holders as &#34;recording&#34; and &#34;reference&#34; by using pieces of lab tape of different colors. This will help guide the mounting of the mosquito head and electrodes.</li>
		<li>Ensure that the electrode holders are clear inside and that no borosilicate debris are present.</li>
		<li>Chloridization: Soak the silver wires of the electrode holders in pure bleach for about 5 min. The wires turn from shiny light grey to matte dark grey.</li>
		<li>Loosen the rubber stopper and fill the inside of the capillary with 10% saline solution using a 20 G needle.</li>
		<li>Fill the borosilicate capillary with the saline solution using a syringe. Ensure that no bubbles are present in neither the electrode holder nor the pulled capillary. 
		&#8203;NOTE: To reduce the chances of having bubbles in the capillary, keep pushing saline in the capillary while gently pulling the needle out and use capillaries with a filament. It is possible to load the capillaries with a solution composed of 1:3 electrode gel and saline solution. This can help prevent evaporation of the saline and can be particularly useful when learning and practicing EAGs, as the experimenter will need more time to complete the different steps.</li>
		<li>After soaking, rinse the silver wires with DI water and insert them in the two capillaries. Ensure that the tip of the wire is less than 1 mm from the tip of the capillary. Ensure that the capillary passes the rubber ring inside the electrode holder without breaking. Gently tighten the rubber stopper. Verify that no air bubbles are present.</li>
		<li>Use the capillary with the wider opening on the reference electrode holder (neck), and the smaller opening on the recording electrode holder (antennae).</li>
		<li>Leave the two mounted electrode holders on a wet cleaning wipe to prevent the tip from drying out until ready to mount the head.</li>
	</ol>
	</li>
</ol>
5. EAG rig preparation (Figure 1)
<ol>
	<li>Ensure that the air table is up, that there is no blockage in the airline. Ensure that the tank of medical air is still full to avoid changing it in the middle of the experiment. Make sure that there are bubbles in the humidifier.</li>
	<li>Air and pulse delivery system
	<ol>
		<li>Turn on the medical air gas tank.</li>
		<li>Check the level of the two flowmeters. 
		&#8203;NOTE: The flowmeter controlling the main air current bathing the preparation during the whole experiment should be at 140 mL/min and the other related to the odor pulse should read 15 mL/min.</li>
	</ol>
	</li>
	<li>If doing GC-EAD, turn on the machine, gas tanks and create/load the file/method.</li>
	<li>Turn on the computers, the software applications, valve power supply, and verify the internet connection for the software application to work.
	<ol>
		<li>Software application: A short script can be written to deliver the pulse.</li>
		<li>EAG software: use any electrophysiology software.</li>
		<li>Implement the parameters in the software (e.g., amplifier, duration of recording, duration of the pulses, etc.).</li>
	</ol>
	</li>
	<li>Deliver a control pulse to verify that the valve delivering the pulses is functional.</li>
	<li>Set the power supply at 5.2 V. Verify the amplifier parameters. 
	NOTE: The parameters used for the data presented here are: low cut-off filter of 0.1 Hz; high cut-off filter of 500 Hz; Gain of x100.</li>
</ol>
6. Mosquito head preparation and mounting (Figure 2)
<ol>
	<li>Place an aluminum plate on ice and place a piece of wet cleaning wipe over it.</li>
	<li>Put a small dollop of electrode gel in a corner.</li>
	<li>Place a mosquito cup on ice and let the mosquito cool down for a couple of minutes, or until it is stops flying. 
	NOTE: Some species are cold-resistant and might require a quick anesthesia over a CO2 fly pad to go down. The least the mosquito stays on either, the better.</li>
	<li>Place the mosquito on its back and clip the tip of each antenna (only a small portion of the last segment) with micro scissors.</li>
	<li>Use forceps to drag the mosquito next to the electrode gel dollop and dip each antenna&#39;s tip gently in the gel. Avoid dipping more than just the very last segment in the electrode gel.</li>
	<li>Using forceps, pull the mosquito antennae out while maintaining them next to each other. Let them come out together from the gel. Make sure that the antennae do not touch the cleaning wipe&#39;s surface, or they might separate.</li>
	<li>Place the mosquito on its side and chop the head using micro scissors or a razor blade. 
	NOTE: Once the head is chopped, proceed quickly to the next steps and to the EAG rig to start the recordings. The preparation should stay responsive for about 30 min.</li>
	<li>Take the reference electrode and gently deep the tip in the gel. Keep in contact with neck tissues and let the head stick on it.</li>
	<li>Move the electrode holders under the EAG microscope and view through the microscope to place the head (i.e., reference) electrode on a micromanipulator. Ensure that the antennae are at the center.</li>
	<li>Grab the recording electrode, place it in front of the antennae tips. Move and align it as close as possible to the tips using the micromanipulator. Using the microscope, move the tip of the recording electrode towards the antennae.</li>
	<li>Connect both electrode holders to the amplifier before inserting the tips to prevent them from moving after insertion.</li>
	<li>Insert the antennae tips in the recording electrode. Ensure that they just contact the saline and the electrode gel and are visible by transparency through the capillary. The antenna goes in by the &#34;suction effect&#34;.</li>
	<li>Adjust the position of the head and tips with forceps under the microscope, if needed.</li>
	<li>Place the airline tubing close to the mosquito head preparation (distance: 1 cm). 
	NOTE: If the head falls off, return to the dissection station and remount the head or prepare a new one if it has been lost or if it has been more than 5 min since the head was cut off. A good connection between the capillary and the neck/antennae is essential for low noise and reliable recording. Ideally, the antennae tips will be at less than 1 mm from the wire of the recording electrode once inserted.</li>
	<li>Turn off the light source, if used.</li>
	<li>Position the vacuum line near the mosquito head preparation (distance: 20 cm) and align with the main airline. 
	&#8203;NOTE: The vacuum will help remove chemicals surrounding the head preparation after the stimulus, which could lead to EAG responses after the pulses were applied.</li>
</ol>
7. Recordings
<ol>
	<li>After insertion of the tips of the antennae, turn on the amplifier and the noise reducer. Observe the baseline signal and ensure that it is not noisy. 
	NOTE: Observe whether large oscillations in the electrical signal are present. Adjust the position of the head and antennae tips as needed until the signal is clean. Use alligator clips to ground anything that introduces noise to the Faraday cage or air table. A baseline signal of less than 0.01 mV in amplitude is ideal for detecting and discriminating minute EAG responses.</li>
	<li>Once the noise level is satisfying, insert the first odor syringe to test in the airline hole.</li>
	<li>Close the Faraday cage. Do not stay in front of the preparation, to reduce noise.</li>
	<li>Click on Record on EAG software.</li>
	<li>Deliver the pulse(s) using the software application. 
	NOTE: The number and duration of the pulses will vary depending on the experiments. Here, single 1 s. pulses per odorant have been used. Odorants were separated by 45 s.</li>
	<li>Note the response of the mosquito antennae in the lab notebook. 
	NOTE: If the odorant is detected by the mosquito antennae, a clear deflection in the signal is observed (see Figure 3A).</li>
	<li>Proceed with the next odor or concentration. Do not forget to randomize the presentation of odorants unless a dose-response curve is performed. 
	NOTE: A negative control and a positive control should be used in the experiments. This will ensure that the responses observed are indeed olfactory responses and not due to mechanical or electric noise.</li>
	<li>At the end of the recording, apply a positive control to verify that the antennae are still responsive. 
	NOTE: Use 0.1% or 1% benzaldehyde as all mosquito species tested so far have been responsive to this compound.</li>
	<li>Proceed with the next mosquito preparation.</li>
</ol>
8. Cleaning
<ol>
	<li>Turn off the amplifier, the noise reducer, the airline, and the computer.</li>
	<li>Return the odorants to the freezer.</li>
	<li>Remove the filter papers from the glass syringes and clean with 100% ethanol if residue is visible on the walls. Let dry on a cleaning wipe overnight.</li>
	<li>Clean the electrode holders with DI water to remove any possible trace of salt. Dry by gently applying against a piece of cleaning wipe.</li>
	<li>Place mosquito remains in the freezer and dispose 24 h later. 
	&#8203;NOTE: If working with infected mosquitoes, follow safety requirements at your&#160;institution.</li>
</ol>
9. Data analyses
<ol>
	<li>Measure either manually or automatically the EAG responses. 
	NOTE: The EAG amplitude (-mV) is measured here. Average if multiple pulses have been applied for each compound. Depending on the software used, EAGs can be automatically detected and measured. However, it is essential to inspect each response individually to verify the shape of the response and assess the possible carry-over, delayed response, etc. The ideal EAG response is aligned with the pulse, shows a clear deflection, and is repeatable between mosquito preparations (Figure 3).</li>
	<li>Present the raw data to show minimal variability, low noise signal, and clear responses (Figure 3B). 
	NOTE: The data can also be normalized (e.g., Z-score). The negative control value (e.g., mineral oil) (i.e., the baseline) can be subtracted from the response and, if not, should be presented in the figures. A positive control should also be presented.</li>
	<li>Perform statistical analysis using any statistics software28.</li>
</ol>

<ol>
	<li>World Health Organization. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=World+health+statistics+2019:+monitoring+health+for+the+SDGs,+sustainable+development+goals.">World health statistics 2019: monitoring health for the SDGs, sustainable development goals.</a> World Health Organization. , (2019).</li><li>Takken, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+olfaction+in+host-seeking+of+mosquitoes:+a+review.">The role of olfaction in host-seeking of mosquitoes: a review.</a> International Journal of Tropical Insect Science. 12 (1-2-3), 287-295 (1991).</li><li>Zwiebel, L. J., Takken, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Olfactory+regulation+of+mosquito-host+interactions.">Olfactory regulation of mosquito-host interactions.</a> Insect Biochemistry and Molecular Biology. 34 (7), 645-652 (2004).</li><li>Potter, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stop+the+biting:+targeting+a+mosquito's+sense+of+smell.">Stop the biting: targeting a mosquito's sense of smell.</a> Cell. 156 (5), 878-881 (2014).</li><li>Paluch, G., Bartholomay, L., Coats, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosquito+repellents:+a+review+of+chemical+structure+diversity+and+olfaction.">Mosquito repellents: a review of chemical structure diversity and olfaction.</a> Pest Management Science. 66 (9), 925-935 (2010).</li><li>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=Electrophysiological+investigation+on+the+antennal+receptors+of+the+silk+moth+during+chemical+and+mechanical+stimulation.">Electrophysiological investigation on the antennal receptors of the silk moth during chemical and mechanical stimulation.</a> Experientia. 13 (2), 89-91 (1957).</li><li>Raguso, R. A., Light, D. M., Pickersky, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram+responses+of+Hyles+lineata+(Sphingidae:+Lepidoptera)+to+volatile+compounds+from+Clarkia+breweri+(Onagraceae)+and+other+moth-pollinated+flowers.">Electroantennogram responses of Hyles lineata (Sphingidae: Lepidoptera) to volatile compounds from Clarkia breweri (Onagraceae) and other moth-pollinated flowers.</a> Journal of Chemical Ecology. 22 (10), 1735-1766 (1996).</li><li>Schweitzer, E. S., Sanes, J. R., Hildebrand, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ontogeny+of+electroantennogram+responses+in+the+moth,+Manduca+sexta.">Ontogeny of electroantennogram responses in the moth, Manduca sexta.</a> Journal of Insect Physiology. 22 (7), 955-960 (1976).</li><li>Martel, V., Anderson, P., Hansson, B. S., Schlyter, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+modulation+of+olfaction+by+physiological+state+in+the+Egyptian+leaf+worm+Spodoptera+littoralis+(Lepidoptera:+Noctuidae).">Peripheral modulation of olfaction by physiological state in the Egyptian leaf worm Spodoptera littoralis (Lepidoptera: Noctuidae).</a> Journal of Insect Physiology. 55 (9), 793-797 (2009).</li><li>Spaethe, J., Brockmann, A., Halbig, C., Tautz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Size+determines+antennal+sensitivity+and+behavioral+threshold+to+odors+in+bumblebee+workers.">Size determines antennal sensitivity and behavioral threshold to odors in bumblebee workers.</a> Naturwissenschaften. 94 (9), 733-739 (2007).</li><li>Suchet, C., 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=Floral+scent+variation+in+two+Antirrhinum+majus+subspecies+influences+the+choice+of+na&#239;ve+bumblebees.">Floral scent variation in two Antirrhinum majus subspecies influences the choice of na&#239;ve bumblebees.</a> Behavioral Ecology and Sociobiology. 65 (5), 1015-1027 (2011).</li><li>De Jong, R., Pham-Del&#232;gue, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram+responses+related+to+olfactory+conditioning+in+the+honeybee+(Apis+mellifera+ligustica).">Electroantennogram responses related to olfactory conditioning in the honeybee (Apis mellifera ligustica).</a> Journal of Insect Physiology. 37 (4), 319-324 (1991).</li><li>Patte, F., Etcheto, M., Marfaing, P., Laffort, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram+stimulus-response+curves+for+59+odourants+in+the+honeybee,+Apis+mellifica.">Electroantennogram stimulus-response curves for 59 odourants in the honeybee, Apis mellifica.</a> Journal of Insect Physiology. 35 (9), 667-675 (1989).</li><li>Alcorta, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+electroantennogram+in+Drosophila+melanogaster+and+its+use+for+identifying+olfactory+capture+and+transduction+mutants.">Characterization of the electroantennogram in Drosophila melanogaster and its use for identifying olfactory capture and transduction mutants.</a> Journal of Neurophysiology. 65 (3), 702-714 (1991).</li><li>Park, K. C., Ochieng, S. A., Zhu, J., Baker, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odor+discrimination+using+insect+electroantennogram+responses+from+an+insect+antennal+array.">Odor discrimination using insect electroantennogram responses from an insect antennal array.</a> Chemical Senses. 27 (4), 343-352 (2002).</li><li>Du, Y. J., Millar, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram+and+oviposition+bioassay+responses+of+Culex+quinquefasciatus+and+Culex+tarsalis+(Diptera:+Culicidae)+to+chemicals+in+odors+from+Bermuda+grass+infusions.">Electroantennogram and oviposition bioassay responses of Culex quinquefasciatus and Culex tarsalis (Diptera: Culicidae) to chemicals in odors from Bermuda grass infusions.</a> Journal of Medical Entomology. 36 (2), 158-166 (1999).</li><li>Costantini, C., 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=Electroantennogram+and+behavioural+responses+of+the+malaria+vector+Anopheles+gambiae+to+human-specific+sweat+components.">Electroantennogram and behavioural responses of the malaria vector Anopheles gambiae to human-specific sweat components.</a> Medical and Veterinary Entomology. 15 (3), 259-266 (2001).</li><li>Collins, L. E., Blackwell, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram+studies+of+potential+oviposition+attractants+for+Toxorhynchites+moctezuma+and+T.+amboinensis+mosquitoes.">Electroantennogram studies of potential oviposition attractants for Toxorhynchites moctezuma and T. amboinensis mosquitoes.</a> Physiological Entomology. 23 (3), 214-219 (1998).</li><li>Seenivasagan, T., Sharma, K. R., Sekhar, K., Ganesan, K., Prakash, S., Vijayaraghavan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram,+flight+orientation,+and+oviposition+responses+of+Aedes+aegypti+to+the+oviposition+pheromone+n-heneicosane.">Electroantennogram, flight orientation, and oviposition responses of Aedes aegypti to the oviposition pheromone n-heneicosane.</a> Parasitology Research. 104 (4), 827-833 (2009).</li><li>Puri, S. N., Mendki, M. J., Sukumaran, D., Ganesan, K., Prakash, S., Sekhar, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroantennogram+and+behavioral+responses+of+Culex+quinquefasciatus+(Diptera:+Culicidae)+females+to+chemicals+found+in+human+skin+emanations.">Electroantennogram and behavioral responses of Culex quinquefasciatus (Diptera: Culicidae) females to chemicals found in human skin emanations.</a> Journal of Medical Entomology. 43 (2), 207-213 (2014).</li><li>Cooperband, M. F., McElfresh, J. S., Millar, J. G., Carde, R. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Attraction+of+female+Culex+quinquefasciatus+Say+(Diptera:+Culicidae)+to+odors+from+chicken+feces.">Attraction of female Culex quinquefasciatus Say (Diptera: Culicidae) to odors from chicken feces.</a> Journal of Insect Physiology. 54 (7), 1184-1192 (2008).</li><li>Dekker, T., Ignell, R., Ghebru, M., Glinwood, R., Hopkins, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+mosquito+repellent+odours+from+Ocimum+forskolei.">Identification of mosquito repellent odours from Ocimum forskolei.</a> Parasites &amp; Vectors. 4 (1), 183 (2011).</li><li>Choo, Y. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reverse+chemical+ecology+approach+for+the+identification+of+an+oviposition+attractant+for+Culex+quinquefasciatus.">Reverse chemical ecology approach for the identification of an oviposition attractant for Culex quinquefasciatus.</a> Proceedings of the National Academy of Sciences. 115 (4), 714-719 (2018).</li><li>Wolff, G. H., Lahond&#232;re, C., Vinauger, C., Riffell, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuromodulation+and+differential+learning+across+mosquito+species.">Neuromodulation and differential learning across mosquito species.</a> bioRxiv. , 755017 (2019).</li><li>Lahond&#232;re, C., 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+olfactory+basis+of+orchid+pollination+by+mosquitoes.">The olfactory basis of orchid pollination by mosquitoes.</a> Proceedings of the National Academy of Sciences. 117 (1), 708-716 (2020).</li><li>Beyenbach, K., Masia, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+conductances+of+principal+cells+in+Malpighian+tubules+of+Aedes+aegypti.">Membrane conductances of principal cells in Malpighian tubules of Aedes aegypti.</a> Journal of Insect Physiology. 48, 375-386 (2002).</li><li>Oesterle, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Pipette Cookbook. , (2018).</li><li>R Core Team. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=R:+A+language+and+environment+for+statistical+computing.">R: A language and environment for statistical computing.</a> R Foundation for Statistical Computing. , (2018).</li><li>Afify, A., Betz, J. F., Riabinina, O., Lahond&#232;re, C., Potter, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Commonly+used+insect+repellents+hide+human+odors+from+Anopheles+mosquitoes.">Commonly used insect repellents hide human odors from Anopheles mosquitoes.</a> Current Biology. 29 (21), 3669-3680 (2019).</li><li>Martin, F., Riveron, J., Alcorta, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Environmental+temperature+modulates+olfactory+reception+in+Drosophila+melanogaster.">Environmental temperature modulates olfactory reception in Drosophila melanogaster.</a> Journal of Insect Physiology. 57 (12), 1631-1642 (2011).</li><li>Qiu, Y. T., Gort, G., Torricelli, R., Takken, W., van Loon, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+blood-feeding+on+olfactory+sensitivity+of+the+malaria+mosquito+Anopheles+gambiae:+application+of+mixed+linear+models+to+account+for+repeated+measurements.">Effects of blood-feeding on olfactory sensitivity of the malaria mosquito Anopheles gambiae: application of mixed linear models to account for repeated measurements.</a> Journal of Insect Physiology. 59 (11), 1111-1118 (2013).</li><li>Taylor, B., Jones, M. D. 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+circadian+rhythm+of+flight+activity+in+the+mosquito+Aedes+aegypti+(L.):+the+phase-setting+effects+of+light-on+and+light+off.">The circadian rhythm of flight activity in the mosquito Aedes aegypti (L.): the phase-setting effects of light-on and light off.</a> Journal of Experimental Biology. 51 (1), 59-70 (1969).</li><li>Eilerts, D. F., VanderGiessen, M., Bose, E. A., Broxton, K., Vinauger, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odor-specific+daily+rhythms+in+the+olfactory+sensitivity+and+behavior+of+Aedes+aegypti+mosquitoes.">Odor-specific daily rhythms in the olfactory sensitivity and behavior of Aedes aegypti mosquitoes.</a> Insects. 9 (4), 147 (2018).</li><li>Krishnan, B., Dryer, S. E., Hardin, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circadian+rhythms+in+olfactory+responses+of+Drosophila+melanogaster.">Circadian rhythms in olfactory responses of Drosophila melanogaster.</a> Nature. 400 (6742), 375-378 (1999).</li><li>Pelletier, J., Guidolin, A., Syed, Z., Cornel, A. J., Leal, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Knockdown+of+a+mosquito+odorant-binding+protein+involved+in+the+sensitive+detection+of+oviposition+attractants.">Knockdown of a mosquito odorant-binding protein involved in the sensitive detection of oviposition attractants.</a> Journal of Chemical Ecology. 36 (3), 245-248 (2010).</li></ol>

The author has nothing to disclose.

Olfactory mediated behaviors are affected by many factors, including physiological (e.g., age, time of day) and environmental (e.g., temperature, relative humidity)30. Thus, when conducting EAGs, it is essential to use insects that are in the same physiological status (i.e., monitoring for age, starving, mating)31 and to also maintain a warm and humid environment around the preparation to avoid desiccation. A temperature around 25 &#176;C is ideal and 60% to 80% humidity fo...

Mosquitoes are the deadliest organisms on earth, claiming the lives of more than one million people per year and place more than half the world population at risk of exposure to the pathogens they transmit, while biting1. These insects rely on a wide range of cues (i.e., thermal, visual, mechanical, olfactory, auditory) to locate a host to feed on (both plant and animal), for mating and oviposition, as well as to avoid predators at both the larval and adult stages2,3. Among these senses, olfaction plays a critical role in the above mentioned behaviors, in particular for medium to long-range detection of odorant molecules2,3. Odors emitted by a host or an oviposition site are detected by various specific olfactory receptors (e.g., GRs, ORs, IRs) located on the mosquito palps proboscis, tarsi, and antennae2,3.
As olfaction is a key component of their host-seeking (plant and animal), mating and oviposition behaviors, it thus constitutes an ideal target to study to develop new tools for mosquito control4. Research on repellents (e.g., DEET, IR3535, picaridin) and baits (e.g., BG sentinel human lure) is extremely prolific5, but because of the current challenges in mosquito control (e.g., insecticide resistance, invasive species), it is essential to develop new efficient control methods informed by the mosquito biology.
Many techniques (e.g., olfactometer, landing assays, electrophysiology) have been used to assess the bioactivity of compounds or mixtures of compounds in mosquitoes. Among them, electroantennography (or electroantennograms (EAGs)) can be used to determine whether the odorants are detected by the mosquito antennae. This technique was initially developed by Schneider6 and has been used in many different insect genera since then, including moths7,8,9, bumblebees10,11, honeybees12,13, and fruit flies14,15 to name a few. Electroantennography has also been employed using various protocols, including single or multiple antennae in mosquitoes16,17,18,19,20,21,22,23,24,25.
Mosquitoes are relatively small and delicate insects with rather thin antennae. While performing EAGs on larger insects such as moths or bumblebees is relatively easy because of their larger size and thicker antennae, conducting EAGs in mosquitoes can be challenging. In particular, maintaining a good signal-to-noise ratio and a lasting responsive preparation are two major requirements for data reproducibility and reliability.
The step-by-step guide to low noise EAGs proposed here directly offers solutions to these limitations and make this protocol applicable to several mosquito species from various genera, including Aedes, Anopheles, Culex, and Toxorhynchites, and describes the technique for both females and males. Electroantennography offers a quick yet reliable way to screen and determine bioactive compounds that can then be leveraged in bait development after valence has been determined with behavioral assays.

<table><tbody><tr><td>Air table Clean Bench</td><td>TMC</td><td>https://www.techmfg.com/products/labtables/cleanbench63series/accessoriess</td><td>Noise reducer</td></tr><tr><td>Analog-to-digital board</td><td>National Instruments</td><td>BNC-2090A</td><td></td></tr><tr><td>Benchtop Flowbuddy Complete</td><td>Genesee Scientific</td><td>59-122BC</td><td>To anesthesize mosquitoes</td></tr><tr><td>Borosillicate glass capillary</td><td>Sutter Instrument</td><td>B100-78-10</td><td>To make the recording and references capillaries</td></tr><tr><td>Chemicals</td><td>Sigma Aldrich</td><td>Benzaldehyde: 418099-100 mL; Butyric acid: B103500-100mL; 1-Hexanol: 471402-100mL; Mineral oil: M8410-1L</td><td>Chemicals used for the experiments presented here</td></tr><tr><td>CO2</td><td>Airgas or Praxair</td><td>N/A</td><td>To anesthesize mosquitoes</td></tr><tr><td>Cold Light Source</td><td>Volpi</td><td>NCL-150</td><td></td></tr><tr><td>Disposable syringes</td><td>BD</td><td>1 mL (309628)&amp;nbsp; / 3 mL (309657)</td><td></td></tr><tr><td>Electrode cables</td><td>World Precision Instruments</td><td>5371</td><td></td></tr><tr><td>Electrode gel salt free</td><td>Parkerlabs</td><td>12-08-Spectra-360</td><td></td></tr><tr><td>Faraday cage</td><td>TMC</td><td>https://www.techmfg.com/products/electric-and-magnetic-field-cancellation/faradaycages</td><td>Noise reducer</td></tr><tr><td>Flowmeters</td><td>Bel-art</td><td>65 mm (H40406-0010) / 150 mm (H40407-0075)</td><td>One of each</td></tr><tr><td>GCMS vials and caps</td><td>Thermo-fisher scientific</td><td>2-SVWKA8-CPK</td><td>To prepare odorant dilutions</td></tr><tr><td>Glass syringes (Fortuna)</td><td>Sigma Aldrich</td><td>Z314307</td><td>For odor delivery to the EAG prep</td></tr><tr><td>Humbug</td><td>Quest Scientific</td><td>http://www.quest-sci.com/</td><td>Noise reducer</td></tr><tr><td>2 mm Jack Holder, Narrow, 90 deg., With Wire</td><td>A-M Systems</td><td>675748</td><td>Electrode holder</td></tr><tr><td>Magnetic bases</td><td>Kanetec</td><td>MB-FX</td><td>x 2</td></tr><tr><td>MATLAB + Toolboxes</td><td>Mathworks</td><td>https://www.mathworks.com/products/matlab.html</td><td>For delivering the pulses</td></tr><tr><td>Medical air</td><td>Airgas or Praxair</td><td>N/A</td><td>For main airline</td></tr><tr><td>Microscope</td><td>Nikkon</td><td>SMZ-800N</td><td></td></tr><tr><td>Micromanipulators Three-Axis Coarse/Fine Compact Micromanipulator</td><td>Narishige</td><td>MHW-3</td><td>x 2</td></tr><tr><td>Microelectrode amplifier with headstage</td><td>A-M Systems</td><td>Model 1800</td><td></td></tr><tr><td>Mosquito rearing supplies</td><td>Bioquip</td><td>https://www.bioquip.com/Search/WebCatalog.asp</td><td></td></tr><tr><td>Needles</td><td>BD</td><td>25G (305127) / 21G (305165)</td><td></td></tr><tr><td>Pasteur pipettes</td><td>Fisher Scientific</td><td>13-678-6A</td><td>For odor delivery to the EAG prep</td></tr><tr><td>PTFE Tubing of different diameters</td><td>Mc Master Carr</td><td>N/A</td><td>To connect solenoid valve, flowmeter, airline ect.</td></tr><tr><td>30V/5A DC Power Supply</td><td>Dr. Meter</td><td>PS-305DM</td><td></td></tr><tr><td>R version 3.5.1</td><td>R project</td><td>https://www.r-project.org/</td><td>For data analyses</td></tr><tr><td>Relay for solenoid valve</td><td>N/A</td><td>Custom made</td><td></td></tr><tr><td>Silver wire 0.01&amp;rdquo;</td><td>A-M Systems</td><td>782500</td><td></td></tr><tr><td>Solenoid valve (3-way)</td><td>The Lee Company</td><td>LHDA0533115H</td><td></td></tr><tr><td>WinEDR software</td><td>Strathclyde Electrophysiology Software</td><td>WinEDR V3.9.1</td><td>For EAG recording</td></tr><tr><td>Whatman paper</td><td>Cole Parmer</td><td>UX-06648-03</td><td>To load chemical in glass syringe / Pasteur pipette</td></tr></tbody></table>

a step-by-step guide to mosquito electroantennography

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when acquiring a blood-meal. To locate a host to feed on, mosquitoes rely on a wide range of sensory cues, including visual, mechanical, thermal, and olfactory. The study details a technique, electroantennography (EAG), that allows researchers to assess whether the mosquitoes can detect individual chemicals and blends of chemicals in a concentration-dependent manner. When coupled with gas-chromatography (GC-EAG), this technique allows to expose the antennae to a full headspace/complex mixture and determines which chemicals present in the sample of interest, the mosquito can detect. This is applicable to host body odors as well as plant floral bouquets or other ecologically relevant odors (e.g., oviposition sites odorants). Here, we described a protocol that permits long durations of preparation responsiveness time and is applicable to both female and male mosquitoes from multiple genera, including Aedes, Culex, Anopheles, and Toxorhynchites mosquitoes. As olfaction plays a major part in mosquito-host interactions and mosquito biology in general, EAGs and GC-EAG can reveal compounds of interest for the development of new disease vector control strategies (e.g., baits). Complemented with behavioral assays, the valence (e.g., attractant, repellent) of each chemical can be determined.

Electroantennography is a powerful tool to determine whether a chemical or blend of chemicals is detected by an insect antenna. It can also be used to determine the detection threshold for a given chemical using a gradual increase of concentration (i.e., dose curve response, Figure 4B). Moreover, it is useful to test the effects of repellent on the response to host-related odors29.
Positive and negative controls should always be used in EAG...

Watch this Scientific Journal Video about A Step-by-Step Guide to Mosquito Electroantennography at JoVE.com

A Step-by-Step Guide to Mosquito Electroantennography

The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males.

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when ...

a-step-by-step-guide-to-mosquito-electroantennography

Research

JoVE Journal

Biology

4.5K Views.  Virginia Polytechnic Institute and State University. The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males. 

Video: A Step-by-Step Guide to Mosquito Electroantennography

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

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

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

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

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

Neuroscience

Early Metamorphic Insertion Technology for Insect Flight Behavior Monitoring

Early Metamorphosis Insertion Technology (EMIT) is a novel methodology for integrating microfabricated neuromuscular recording and actuation platforms on insects during their metamorphic development. Here, the implants are fused within the structure and function of the neuromuscular system as a result of metamorphic tissue remaking. The implants emerge with the insect where the development of tissue around the electronics during pupal development results in a bioelectrically and biomechanically enhanced tissue interface. This relatively more reliable and stable interface would be beneficial for many researchers exploring the neural basis of the insect locomotion with alleviated traumatic effects caused during adult stage insertions. In this article, we implant our electrodes into the indirect flight muscles of Manduca sexta. Located in the dorsal-thorax, these main flight powering dorsoventral and dorsolongitudinal muscles actuate the wings and supply the mechanical power for up and down strokes. Relative contraction of these two muscle groups has been under investigation to explore how the yaw maneuver is neurophysiologically coordinated. To characterize the flight dynamics, insects are often tethered with wires and their flight is recorded with digital cameras. We also developed a novel way to tether Manduca sexta on a magnetically levitating frame where the insect is connected to a commercially available wireless neural amplifier. This set up can be used to limit the degree of freedom to yawing &#8220;only&#8221; while transmitting the related electromyography signals from dorsoventral and dorsolongitudinal muscle groups.

We present a novel surgical procedure to implant electrodes in Manduca sexta during its early metamorphic stages. This technique allows mechanically stable and electrically reliable coupling with the neuromuscular tissue to study flight neurophysiology dynamics. We also present a novel magnetic levitation platform for tethered studies of insect yaw.

Early Metamorphosis Insertion Technology (EMIT) is a novel methodology for integrating microfabricated neuromuscular recording and actuation platforms on ...

Behavior

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" ...

Electrophysiological Measurements from a Moth Olfactory System

Insect olfactory systems provide unique opportunities for recording odorant-induced responses in the forms of electroantennograms (EAG) and single sensillum recordings (SSR), which are summed responses from all odorant receptor neurons (ORNs) located on the antenna and from those housed in individual sensilla, respectively. These approaches have been exploited for getting a better understanding of insect chemical communication. The identified stimuli can then be used as either attractants or repellents in management strategies for insect pests.

Insect olfactory systems provide unique opportunities for recording odorant-induced responses in the forms of electroantennograms (EAG) and single sensillum recordings (SSR), which are summed responses from all odorant receptor neurons (ORNs) located on the antenna and from those housed in individual sensilla, respectively.

Insect olfactory systems provide unique opportunities for recording odorant-induced responses in the forms of electroantennograms (EAG) and single ...

Using Insect Electroantennogram Sensors on Autonomous Robots for Olfactory Searches

Robots designed to track chemical leaks in hazardous industrial facilities1 or explosive traces in landmine fields2 face the same problem as insects foraging for food or searching for mates3: the olfactory search is constrained by the physics of turbulent transport4. The concentration landscape of wind borne odors is discontinuous and consists of sporadically located patches. A pre-requisite to olfactory search is that intermittent odor patches are detected. Because of its high speed and sensitivity5-6, the olfactory organ of insects provides a unique opportunity for detection. Insect antennae have been used in the past to detect not only sex pheromones7 but also chemicals that are relevant to humans, e.g., volatile compounds emanating from cancer cells8 or toxic and illicit substances9-11. We describe here a protocol for using insect antennae on autonomous robots and present a proof of concept for tracking odor plumes to their source. The global response of olfactory neurons is recorded in situ in the form of electroantennograms (EAGs). Our experimental design, based on a whole insect preparation, allows stable recordings within a working day. In comparison, EAGs on excised antennae have a lifetime of 2 hr. A custom hardware/software interface was developed between the EAG electrodes and a robot. The measurement system resolves individual odor patches up to 10 Hz, which exceeds the time scale of artificial chemical sensors12. The efficiency of EAG sensors for olfactory searches is further demonstrated in driving the robot toward a source of pheromone. By using identical olfactory stimuli and sensors as in real animals, our robotic platform provides a direct means for testing biological hypotheses about olfactory coding and search strategies13. It may also prove beneficial for detecting other odorants of interests by combining EAGs from different insect species in a bioelectronic nose configuration14 or using nanostructured gas sensors that mimic insect antennae15.

We describe a protocol for using insect antennae in the form of electroantennograms (EAGs) on autonomous robots. Our experimental design allows stable recordings within a day and resolves individual odor patches up to 10 Hz. The efficiency of EAG sensors for olfactory searches is demonstrated in driving a robot toward an odor source.

Robots designed to track chemical leaks in hazardous industrial facilities1 or explosive traces in landmine fields2 face the same problem as insects ...

Electroantennography-based Bio-hybrid Odor-detecting Drone using Silkmoth Antennae for Odor Source Localization

Small drones with chemical or biosensor devices that can detect airborne odorant molecules have attracted considerable attention owing to their applicability in environmental and security monitoring and search-and-rescue operations. Small drones with commercial metal-oxide-semiconductor (MOX) gas sensors have been developed for odor source localization; however, their real-time-odor-detection performance has proven inadequate. However, biosensing technologies based on insect olfactory systems exhibit relatively high sensitivity, selectivity, and real-time response with respect to odorant molecules compared to commercial MOX gas sensors. In such devices, excised insect antennae function as portable odorant biosensor elements and have been found to deliver excellent sensing performance. This study presents experimental protocols for odorant-molecule detection in the air using a small autonomous bio-hybrid drone based on a mountable electroantennography (EAG) device incorporating silkmoth antennae.
We developed a mountable EAG device including sensing/processing parts with a Wi-Fi module. The device was equipped with a simple sensor enclosure to enhance the sensor directivity. Thus, odor source localization was conducted using the spiral-surge algorithm, which does not assume an upwind direction. The experimental bio-hybrid odor-detecting drone identified real-time odorant-concentration differences in a pseudo-open environment (outside a wind tunnel) and localized the source. The developed drone and associated system can serve as an efficient odorant molecule-detection tool and a suitable flight platform for developing odor source localization algorithms owing to its high programmability.

This study introduces experimental protocols for a bio-hybrid odor-detecting drone based on silkmoth antennae. The operation of an experimental electroantennogram device with silkmoth antennae is presented, in addition to the structure of a bio-hybrid drone designed for odor source localization using the spiral-surge algorithm.

Small drones with chemical or biosensor devices that can detect airborne odorant molecules have attracted considerable attention owing to their ...

Engineering

Electroantennographic Bioassay as a Screening Tool for Host Plant Volatiles

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host plant.1,2 When a host plant is an important agronomic commodity feeding damage by insect pests can inflict serious economic losses to growers. Accordingly, kairomones can be used as attractants to lure or confuse these insects and, thus, offer an environmentally friendly alternative to pesticides for insect control.3 Unfortunately, plants can emit a vast number volatiles with varying compositions and ratios of emissions dependent upon the phenology of the commodity or the time of day. This makes identification of biologically active components or blends of volatile components an arduous process. To help identify the bioactive components of host plant volatile emissions we employ the laboratory-based screening bioassay electroantennography (EAG). EAG is an effective tool to evaluate and record electrophysiologically the olfactory responses of an insect via their antennal receptors. The EAG screening process can help reduce the number of volatiles tested to identify promising bioactive components. However, EAG bioassays only provide information about activation of receptors. It does not provide information about the type of insect behavior the compound elicits; which could be as an attractant, repellent or other type of behavioral response. Volatiles eliciting a significant response by EAG, relative to an appropriate positive control, are typically taken on to further testing of behavioral responses of the insect pest. The experimental design presented will detail the methodology employed to screen almond-based host plant volatiles4,5 by measurement of the electrophysiological antennal responses of an adult insect pest navel orangeworm (Amyelois transitella) to single components and simple blends of components via EAG bioassay. The method utilizes two excised antennae placed across a "fork" electrode holder. The protocol demonstrated here presents a rapid, high-throughput standardized method for screening volatiles. Each volatile is at a set, constant amount as to standardize the stimulus level and thus allow antennal responses to be indicative of the relative chemoreceptivity. The negative control helps eliminate the electrophysiological response to both residual solvent and mechanical force of the puff. The positive control (in this instance acetophenone) is a single compound that has elicited a consistent response from male and female navel orangeworm (NOW) moth. An additional semiochemical standard that provides consistent response and is used for bioassay studies with the male NOW moth is (Z,Z)-11,13-hexdecadienal, an aldehyde component from the female-produced sex pheromone.6-8

A method to rapidly screen host plant volatiles by measurement of the electrophysiological response of adult navel orangeworm (Amyelois transitella) antennae to single components and blends via electroantennographic analysis is demonstrated.

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host ...

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of these disease vectors. We have recently identified a bitter sensing neuron in the labellum of female Aedes aegypti that responds to DEET and other repellents, as well as bitter quinine, through direct electrophysiological investigation. These gustatory receptor neuron responses prompted our sequencing of total mRNA from both male and female labella and tarsi samples to elucidate the putative chemoreception genes expressed in these contact chemoreception tissues. Samples of tarsi were divided into pro-, meso- and metathoracic subtypes for both sexes. We then validated our dataset by conducting qRT-PCR on the same tissue samples and used statistical methods to compare results between the two methods. Studies addressing molecular function may now target specific genes to determine those involved in repellent perception by mosquitoes. These receptor pathways may be used to screen novel repellents towards disruption of host-seeking behavior to curb the spread of harmful viruses.

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Using two methods to estimate gene expression in the major gustatory appendages of Aedes aegypti, we have identified the set of genes putatively underlying the neuronal responses to bitter and repulsive compounds, as determined by electrophysiological examination.

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of ...

Studying the Activity of Neuropeptides and Other Regulators of the Excretory System in the Adult Mosquito

Studies of insect physiology, particularly in those species that are vectors of pathogens causing disease in humans and other vertebrates, provide the foundation to develop novel strategies for pest control. Here, a series of methods are described that are routinely utilized to determine the functional roles of neuropeptides and other neuronal factors (i.e., biogenic amines) on the excretory system of the mosquito, Aedes aegypti. The Malpighian tubules (MTs), responsible for primary urine formation, can continue functioning for hours when removed from the mosquito, allowing for fluid secretion measurements following hormone treatments. As such, the Ramsay assay is a useful technique to measure secretion rates from isolated MTs. Ion-selective microelectrodes (ISME) can sequentially be used to measure ion concentrations (i.e., Na+ and K+) in the secreted fluid. This assay allows for the measurement of several MTs at a given time, determining the effects of various hormones and drugs. The Scanning Ion-selective Electrode Technique uses ISME to measure voltage representative of ionic activity in the unstirred layer adjacent to the surface of ion transporting organs to determine transepithelial transport of ions in near real time. This method can be used to understand the role of hormones and other regulators on ion absorption or secretion across epithelia. Hindgut contraction assays are also a useful tool to characterize myoactive neuropeptides, that may enhance or reduce the ability of this organ to remove excess fluid and waste. Collectively, these methods provide insight into how the excretory system is regulated in adult mosquitoes. This is important because functional coordination of the excretory organs is crucial in overcoming challenges such as desiccation stress after eclosion and before finding a suitable vertebrate host to obtain a bloodmeal.

This protocol outlines methodologies behind the Ramsay assay, ion-selective microelectrodes, Scanning Ion-selective Electrode Technique (SIET) and in vitro contraction assays, applied to study the adult mosquito excretory system, comprised of the Malpighian tubules and hindgut, to collectively measure ion and fluid secretion rates, contractile activity, and transepithelial ion transport.

Studies of insect physiology, particularly in those species that are vectors of pathogens causing disease in humans and other vertebrates, provide the ...

Non-invasive Electropenetrography to Study Mosquito Probing Behaviors

AC-DC Electropenetrography for the Study of Probing and Ingestion Behaviors of Culex tarsalis Mosquitoes on Human Hands

Mosquitoes transmit pathogens that negatively affect human and animal health. A greater understanding of their blood-feeding biology and interactions with hosts and pathogens could be exploited to develop new targets for controlling mosquito-borne diseases. Unfortunately, probing (i.e., biting) behaviors of mosquitoes are poorly understood because they occur inside host tissues. Here, a non-invasive procedure is described for using AC-DC electropenetrography (EPG) to indirectly visualize and quantify mosquito feeding behaviors by recording changes in electrical signals generated when probing and ingesting on human hands. Thin gold wires are attached to the mosquitoes using conductive silver glue and connected to the EPG instrument. The human host holds a substrate voltage probe in their hand. Probing of the host by the wired mosquito completes the electrical circuit, and electrical signals are recorded on a computer as &#34;waveforms&#34; that can then be measured and enumerated for analysis. This methodology has been used to characterize the probing and ingestion behaviors of Aedes aegypti and Culex tarsalis mosquitoes and can be applied to other mosquito species. EPG can be used to study the effects of pathogens, insecticides, and other factors on mosquito feeding behaviors.

Procedures are described for studying the probing and ingestion behaviors of Culex tarsalis&#160;mosquitoes on human hands using AC-DC electropenetrography.

Mosquitoes transmit pathogens that negatively affect human and animal health. A greater understanding of their blood-feeding biology and interactions with ...

A Step-by-Step Guide to Mosquito Electroantennography

Summary

Explore More Videos

Chapters in this video

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Early Metamorphic Insertion Technology for Insect Flight Behavior Monitoring

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

Electrophysiological Measurements from a Moth Olfactory System

Using Insect Electroantennogram Sensors on Autonomous Robots for Olfactory Searches

Electroantennography-based Bio-hybrid Odor-detecting Drone using Silkmoth Antennae for Odor Source Localization

Electroantennographic Bioassay as a Screening Tool for Host Plant Volatiles

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Studying the Activity of Neuropeptides and Other Regulators of the Excretory System in the Adult Mosquito

AC-DC Electropenetrography for the Study of Probing and Ingestion Behaviors of Culex tarsalis Mosquitoes on Human Hands