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>

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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)&#160; / 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&#8221;</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 ...

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4.4K 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

Protocol for Mosquito Rearing (A. gambiae)

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night cycle. For this procedure, you'll need mosquito cages, 10% sterile sucrose solution, paper towels, beaker, whatman filter paper, glass feeders, human blood and serum, water bath, parafilm, distilled water, clean plastic trays, mosquito food (described below), mosquito net to cover the trays, vacuum, and a collection chamber to collect adults.

Protocol for Mosquito Rearing A. gambiae

This video illustrates the general techniques used to rear Anopheles gambiae in the laboratory. The methods for caring for laboratory mosquitoes are demonstrated through all stages of the organism's life cycle from larvae to pupae to blood-feeding adults.

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night ...

Building a Better Mosquito: Identifying the Genes Enabling Malaria and Dengue Fever Resistance in A. gambiae and A. aegypti Mosquitoes

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections.   Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections. Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus ...

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing and adult tissue. An advance in this concept is Genetic Inducible Fate Mapping (GIFM), linking gene expression, cell fate, and cell behaviors in vivo, to create fate maps based on genetic lineage.
GIFM exploits X-CreER lines where X is a gene or set of gene regulatory elements that confers spatial expression of a modified bacteriophage protein, Cre recombinase (CreERT). CreERT contains a modified estrogen receptor ligand binding domain which renders CreERT sequestered in the cytoplasm in the absence of the drug tamoxifen. The binding of tamoxifen releases CreERT, which translocates to the nucleus and mediates recombination between DNA sequences flanked by loxP sites. In GIFM, recombination typically occurs between a loxP flanked Stop cassette preceding a reporter gene such as GFP.
Mice are bred to contain either a region- or cell type-specific CreER and a conditional reporter allele. Untreated mice will not have marking because the Stop cassette in the reporter prevents further transcription of the reporter gene. We administer tamoxifen by oral gavage to timed-pregnant females, which provides temporal control of CreERT release and subsequent translocation to the nucleus removing the Stop cassette from the reporter. Following recombination, the reporter allele is constitutively and heritably expressed. This series of events marks cells such that their genetic history is indelibly recorded. The recombined reporter thus serves as a high fidelity genetic lineage tracer that, once on, is uncoupled from the gene expression initially used to drive CreERT.
We apply GIFM in mouse to study normal development and ascertain the contribution of genetic lineages to adult cell types and tissues. We also use GIFM to follow cells on mutant genetic backgrounds to better understand complex phenotypes that mimic salient features of human genetic disorders.
This video article guides researchers through experimental methods to successfully apply GIFM. We demonstrate the method using our well characterized Wnt1-CreERT;mGFP mice by administering tamoxifen at embryonic day (E)8.5 via oral gavage followed by dissection at E12.5 and analysis by epifluorescence stereomicroscopy. We also demonstrate how to micro-dissect fate mapped domains for explant preparation or FACS analysis and dissect adult fate-mapped brains for whole mount fluorescent imaging. Collectively, these procedures allow researchers to address critical questions in developmental biology and disease models.

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Genetic Inducible Fate Mapping (GIFM) marks and tracks cells with fine spatial and temporal control in vivo and elucidates how cells from a specific genetic lineage contribute to developing and adult tissues. Demonstrated here are the techniques required to fate map E12.5 mouse embryos for epifluorescent and explant analysis.

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing ...

A Step-by-step Method for the Reconstitution of an ABC Transporter into Nanodisc Lipid Particles

The nanodisc is a discoidal particle (~ 10-12 nm large) that trap membrane proteins into a small patch of phospholipid bilayer. The nanodisc is a particularly attractive option for studying membrane proteins, especially in the context of ligand-receptor interactions. The method pioneered by Sligar and colleagues is based on the amphipathic properties of an engineered highly a-helical scaffold protein derived from the apolipoprotein A1. The hydrophobic faces of the scaffold protein interact with the fatty acyl side-chains of the lipid bilayer whereas the polar regions face the aqueous environment. Analyses of membrane proteins in nanodiscs have significant advantages over liposome because the particles are small, homogeneous and water-soluble. In addition, biochemical and biophysical methods normally reserved to soluble proteins can be applied, and from either side of the membrane. In this visual protocol, we present a step-by-step reconstitution of a well characterized bacterial ABC transporter, the MalE-MalFGK2 complex. The formation of the disc is a self-assembly process that depends on hydrophobic interactions taking place during the progressive removal of the detergent. We describe the essential steps and we highlight the importance of choosing a correct protein-to-lipid ratio in order to limit the formation of aggregates and larger polydisperse liposome-like particles. Simple quality controls such as gel filtration chromatography, native gel electrophoresis and dynamic light scattering spectroscopy ensure that the discs have been properly reconstituted.

Nanodiscs are small discoid particles that incorporate membrane proteins into a small patch of phospholipid bilayer. We provide a visual protocol that shows the step-by-step incorporation of the MalFGK2 transporter into a disc.

The nanodisc is a discoidal particle (~ 10-12 nm large) that trap membrane proteins into a small patch of phospholipid bilayer. The nanodisc is a ...

A Practical Guide to Phylogenetics for Nonexperts

Many researchers, across incredibly diverse foci, are applying phylogenetics to their research question(s). However, many researchers are new to this topic and so it presents inherent problems. Here we compile a practical introduction to phylogenetics for nonexperts.&#160;We outline in a step-by-step manner, a pipeline for generating reliable phylogenies from gene sequence datasets. We begin with a user-guide for similarity search tools via online interfaces as well as local executables. Next, we explore programs for generating multiple sequence alignments followed by protocols for using software to determine best-fit models of evolution. We then outline protocols for reconstructing phylogenetic relationships via maximum likelihood and Bayesian criteria&#160;and finally describe tools for visualizing phylogenetic trees. While this is not by any means an exhaustive description of phylogenetic approaches, it does provide the reader with practical starting information on key software applications commonly utilized by phylogeneticists. The vision for this article would be that it could serve as a practical training tool for researchers embarking on phylogenetic studies&#160;and also serve as an educational resource that could be incorporated into a classroom or teaching-lab.

Here we describe a step-by-step pipeline for generating reliable phylogenies from nucleotide or amino acid sequence datasets. This guide aims to serve researchers or students new to phylogenetic analysis.&#160;

Many researchers, across incredibly diverse foci, are applying phylogenetics to their research question(s). However, many researchers are new to this ...

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 ...

A Guide to Modern Quantitative Fluorescent Western Blotting with Troubleshooting Strategies

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins within complex biological samples. Since then, western blotting methodology has become a common component of the molecular biologists experimental repertoire. A cursory search of PubMed using the term &#8220;western blot&#8221; suggests that in excess of two hundred and twenty thousand published manuscripts have made use of this technique by the year 2014. Importantly, the last ten years have seen technical imaging advances coupled with the development of sensitive fluorescent labels which have improved sensitivity and yielded even greater ranges of linear detection. The result is a now truly Quantifiable Fluorescence based Western Blot (QFWB) that allows biologists to carry out comparative expression analysis with greater sensitivity and accuracy than ever before. Many &#8220;optimized&#8221; western blotting methodologies exist and are utilized in different laboratories. These often prove difficult to implement due to the requirement of subtle but undocumented procedural amendments. This protocol provides a comprehensive description of an established and robust QFWB method, complete with troubleshooting strategies.

The advancement of western blotting using fluorescence has allowed detection of subtle changes in protein expression enabling quantitative analyses. Here we describe a robust methodology for detection of a range of proteins across a variety of species and tissue types. A strategy to overcome common technical problems is also provided.

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins ...

Endotracheal Intubation of Rabbits Using a Polypropylene Guide Catheter

Endotracheal intubation in rabbits can be challenging due to their unusual anatomy. Achieving a patent airway during anesthesia is critical for the avoidance of airway obstruction, prevention of gastric tympany, and to allow ventilatory support. Due to the difficulty of intubation, alternative methods such as the use of laryngeal mask airways or laryngeal tubes have been explored. However, these methods do not result in direct access to the trachea and thus may present a risk for development of complications. In addition, lack of direct intubation of the trachea can result in personnel exposure to waste anesthetic gases. Numerous methods for endotracheal intubation have been described, including blind placement, use of a fiberoptic laryngoscope or endoscope, and cricoid placement. Despite these numerous publications, many still struggle to achieve success. Here we provide a detailed description of an intubation technique that can be taught with minimal training with a short time to proficiency. Briefly, after administration of injectable anesthesia and proper positioning of the rabbit, a polypropylene catheter is placed into the trachea by direct visualization using a laryngoscope. The catheter is then used as a guide to direct the endotracheal tube into the trachea. This method allows for intubation without the need for expensive equipment and can be performed by a single individual without the need for an assistant. In conclusion, this technique can be easily taught and performed at very little cost in any clinical or research setting.

Endotracheal intubation in rabbits is challenging due to their unusual anatomy. Here we present a technique for direct intubation of the trachea using a polypropylene catheter as a guide. This method utilizes relatively inexpensive supplies, requires minimal training, and can be easily performed in any clinical setting.

Endotracheal intubation in rabbits can be challenging due to their unusual anatomy. Achieving a patent airway during anesthesia is critical for the ...

A Magnetic-Bead-Based Mosquito DNA Extraction Protocol for Next-Generation Sequencing

A recently published DNA extraction protocol using magnetic beads and an automated DNA extraction instrument suggested that it is possible to extract high quality and quantity DNA from a well-preserved individual mosquito sufficient for downstream whole genome sequencing. However, reliance on an expensive automated DNA extraction instrument can be prohibitive for many laboratories. Here, the study provides a budget-friendly magnetic-bead-based DNA extraction protocol, which is suitable for low to medium throughput. The protocol described here was successfully tested using individual Aedes aegypti mosquito samples. The reduced costs associated with high quality DNA extraction will increase the application of high throughput sequencing to resource limited labs and studies.

Described here is a DNA extraction protocol using magnetic beads to produce high quality DNA extractions from mosquitoes. These extractions are suitable for a downstream next-generation sequencing approach.

A recently published DNA extraction protocol using magnetic beads and an automated DNA extraction instrument suggested that it is possible to extract high ...

A Step-By-Step Method to Detect Neutralizing Antibodies Against AAV using a Colorimetric Cell-Based Assay

Recombinant adeno-associated viruses (rAAV) have proven to be a safe and successful vector for transferring genetic material to treat various health conditions in both the laboratory and the clinic. However, pre-existing neutralizing antibodies (NAbs) against AAV capsids pose an ongoing challenge for the successful administration of gene therapies in both large animal experimental models and human populations. Preliminary screening for host immunity against AAV is necessary to ensure the efficacy of AAV-based gene therapies as both a research tool and as a clinically viable therapeutic agent. This protocol describes a colorimetric in vitro assay to detect neutralizing factors against AAV serotype 6 (AAV6). The assay utilizes the reaction between an AAV encoding an alkaline phosphatase (AP) reporter gene and its substrate NBT/BCIP, which generates an insoluble quantifiable purple stain upon combination. 
In this protocol, serum samples are combined with an AAV expressing AP and incubated to permit potential neutralizing activity to occur. Virus serum mixture is subsequently added to cells to allow for viral transduction of any AAVs that have not been neutralized. The NBT/BCIP substrate is added and undergoes a chromogenic reaction, corresponding to viral transduction and neutralizing activity. The proportion of area colored is quantitated using a free software tool to generate neutralizing titers. This assay displays a strong positive correlation between coloration and viral concentration. Assessment of serum samples from sheep before and after administration of a recombinant AAV6 led to a dramatic increase in neutralizing activity (125 to &gt;10,000-fold increase). The assay displayed adequate sensitivity to detect neutralizing activity in &gt;1:32,000 serum dilutions. This assay provides a simple, rapid, and cost-effective method to detect NAbs against AAVs.

A comprehensive laboratory protocol and analysis workflow are described for a rapid, cost-effective, and straightforward colorimetric cell-based assay to detect neutralizing elements against AAV6.

Recombinant adeno-associated viruses (rAAV) have proven to be a safe and successful vector for transferring genetic material to treat various health ...