Supported by USDA-NIFA/AFRI 2010-65105-20582, NSF 0918177, and Bedoukian Research Incorporation.

1. Insects
<ol>
<li>Navel orangeworm, Amyelois transitella (Walker) (Lepidoptera:Pyralidae) colony in our laboratory originated from moths collected from almond trees in Fresno, CA. Our insects colony is maintained in environmental chambers (Percival Inc, IA ) at 28 &plusmn; 2&deg;C, 75 &plusmn; 10% relative humidity, and under 16:8h (light:dark) photo regime.</li>
<li>Emerging moths are collected daily, sexed, and transferred to plastic boxes (669 mL lunchbox,13 x 13 cm; height, 4.5 cm, Rubbermaid) provided with up to 10 layers of water soaked paper towels (Thirsty Ultra Absorbent, 27.9 x 27.9 cm; Safeway). Box covers are perforated to allow air circulation. This arrangement provides ~100% RH.</li>
<li>For electrophysiological measurements we use 2-4 days old moths after emergence, which are kept under high humidity.</li>
</ol>
2. Electrophysiological Preparation
<ol>
<li>Moths (males or females) are gently pushed into a pipette tip (200 &mu;L, USA Scientific Inc) that is cut from the top to have a large (ca. 2 mm) tip diameter. A moth is gently pushed through the other end with a humidified tissue paper stub until the antennae and part of the head protrude from the tip.</li>
<li>Exposed head is immobilized by covering the head with a non-drying clay (Claytoon , Van Aken International, CA) leaving a small part of one eye and two antenna exposed. This preparation is placed on a platform of the EAG Micromanipulator MP-12 (Syntech, Germany) between the two electrode holders.</li>
<li>We use glass electrode made from 1.0 mm borosilicate capillary tubing with filament (WPI Inc, FL) and pulled in a Model P-97 micropipette puller (Sutter Instruments, CA) to obtain microelectrodes of less than a micrometer tip. Electrodes are back filled with sensillum lymph ringer (Kaissling and Thorson 1980). One electrode impales the exposed area of the eye under the microscope (Leica MZ75) and serves as reference, while the recording electrode accommodates the two antennae. To facilitate the contact, distal 1-2 antennal segments are cut before being inserted into the recording electrode. For recordings signals from individual sensilla, we slightly modify this setup wherein the recording electrode is impaled under the microscope (Olympus BX51WI; 800x magnification). Action potentials are recorded by inserting the glass electrode at the base of a sensillum. Signals are amplified by a high impedance pre-amplifier (Syntech, Germany) and fed into a UB-IDAC to be analyzed off-line with AUTOSPIKE software (Syntech, Germany). AC signals are band pass filtered between 100 -10,000 Hz and action potentials are extracted by computer using an AUTOSPIKE algorithm that distinguishes their peak-to-trough amplitudes from noise. Responses of individual neurons are calculated as the increase (or decrease) in action potential frequency (spikes per second) over the spontaneous frequency.</li>
</ol>
3. 	Chemical Stimuli and Stimulation Protocol 
<ol>
<li>Chemicals of highest purity are used. A major constituent of the female pheromone blend from the navel orangeworm, (Z,Z)-11,13-hexadecadienal was obtained from Bedoukian Research Inc, CT. Chemicals are diluted, w/v, with glass-distilled hexane to make stock solutions of 10 &mu;g/ &mu;L and decadic dilutions are made. An aliquot (10 &mu;L) of a stimulus chemical dissolved in hexane is loaded onto a Whatman filter paper strip (8x40 mm), the solvent is evaporated by gently shaking for 10 s under a fume hood and the strip is placed in a 5 mL polypropylene syringe (BD Syringes, NJ) from which various volumes are ejected. Hexane alone and an empty syringe serve as control.</li>
<li>The preparation is held in a humidified air stream delivered by the Syntech Stimulus controller (CS-55 model; Syntech, Germany) at 20 mL/sec to which a stimulus pulse of 4 mL/s is added for 500 ms. Signals are recorded for 10 s, starting 2 s before the onset of the stimulus pulse. Antennal preparation is stimulated with 500 ms pulse during which ca. 2 mL of the purified air from a 5 mL polypropylene syringe containing the stimulus is added onto the main air flow. A gap of at least 1 min or more, following high responses, is allowed between stimulations.</li>
</ol>
4. Representative Results
Antennae of restrained moths (Figure 1) stimulated with female pheromone component, (Z,Z)-11,13-hexadecadienal generate robust dose dependent responses (Figure 2). This major pheromone component also elicits dose dependent excitatory responses from long trichodea sensilla (Figure 3A) as displayed in traces of single unit recordings (Figure 3B).
<img src="/files/ftp_upload/2489/2489fig1.jpg" alt="Figure 1" /> 
Figure 1. Live navel orangeworm moth restrained and antenna exposed 
<img src="/files/ftp_upload/2489/2489fig2.jpg" alt="Figure 2" /> 
Figure 2. Dose dependent EAG responses elicited in response to (Z,Z)-11,13-hexadecadienal, a major female pheromone constituent.
<img src="/files/ftp_upload/2489/2489fig3.jpg" alt="Figure 3" /> 
Figure 3. Sensilla trichoid in male navel orangeworm moth respond to (Z,Z)-11,13-hexadecadienal in a dose dependent manner. (A) Antenna of male navel orangeworm moth is multi-segmented and each segment is adorned with a large number of hair like structures, sensilla. A scanning electron micrograph, showing details in the inset (Scale bars are 200 and 50 &mu;M, respectively). (B) Extracellular single-unit recordings from a trichoid sensillum.

<ol>
	<li>Kaissling, K. -. E., Thorson, J., Sattelle, D. B., Hall, L. M., 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=Insect+olfactory+sensilla:+structural,+chemical+and+electrical+aspects+of+the+functional+organization.">Insect olfactory sensilla: structural, chemical and electrical aspects of the functional organization.</a> Receptors for Neurotransmitters. Hormones and Pheromones in Insects. , 261-282 (1980).</li></ol>

No conflicts of interest declared.

This novel method developed for restraining a live navel orangeworm moth to measure olfactory signals has proved to be robust and very reliable. We routinely employ this method for isolating and identifying novel attractants from natural host substrates like almonds and pistachios.

<table><tbody><tr><td>Microscope </td><td>Olympus Corporation</td><td>BX51WI model</td><td></td></tr><tr><td>Stereo microscope</td><td>Leica Microsystems</td><td>MZ75</td><td></td></tr><tr><td>1.0 mm borosilicate capillary tubing with filament </td><td>World Precision Instruments, Inc.</td><td>1B100F-3</td><td></td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument Co.</td><td>P-97</td><td></td></tr><tr><td>Stimulus controller </td><td>Syntech</td><td>CS-55 model</td><td></td></tr><tr><td>High Impedance pre-amplifiers (Universal Single ended probe)</td><td>Syntech</td><td></td><td></td></tr><tr><td>Amplifier / data-acquisition system (USB-IDAC)</td><td>Syntech</td><td></td><td></td></tr><tr><td>EAG Micromanipulator MP-12 </td><td>Syntech</td><td></td><td></td></tr><tr><td>(Z,Z)-11,13-hexadecadienal</td><td>Bedoukian Research Inc, CT.</td><td></td><td></td></tr><tr><td>Whatman filter paper</td><td>Whatman, GE Healthcare</td><td>1001070</td><td></td></tr><tr><td>5 mL polypropylene syringe </td><td>BD Biosciences</td><td>309633</td><td></td></tr><tr><td>pipette tip (200 &amp;mu;L)</td><td>USA Scientific, Inc.</td><td>1111-0806</td><td></td></tr><tr><td>669 mL lunchbox,13 x 13 cm; height, 4.5 cm, </td><td>Rubbermaid</td><td></td><td></td></tr><tr><td>Thirsty Ultra Absorbent, 27.9 x 27.9 cm</td><td>Safeway</td><td></td><td></td></tr><tr><td>Non-drying clay </td><td>Claytoon , Van Aken International, CA</td><td>18150</td><td></td></tr><tr><td>Environmental chamber</td><td></td><td>I-30BLL model</td><td></td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Electrophysiological Measurements from a Moth Olfactory System at JoVE.com

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.

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

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13.7K Views.  University of California, Davis. 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. 

Video: Electrophysiological Measurements from a Moth Olfactory System

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 ...

Identification of Olfactory Volatiles using Gas Chromatography-Multi-unit Recordings (GCMR) in the Insect Antennal Lobe

All organisms inhabit a world full of sensory stimuli that determine their behavioral and physiological response to their environment. Olfaction is especially important in insects, which use their olfactory systems to respond to, and discriminate amongst, complex odor stimuli. These odors elicit behaviors that mediate processes such as reproduction and habitat selection1-3. Additionally, chemical sensing by insects mediates behaviors that are highly significant for agriculture and human health, including pollination4-6, herbivory of food crops7, and transmission of disease8,9. Identification of olfactory signals and their role in insect behavior is thus important for understanding both ecological processes and human food resources and well-being.
	To date, the identification of volatiles that drive insect behavior has been difficult and often tedious. Current techniques include gas chromatography-coupled electroantennogram recording (GC-EAG), and gas chromatography-coupled single sensillum recordings (GC-SSR)10-12. These techniques proved to be vital in the identification of bioactive compounds. We have developed a method that uses gas chromatography coupled to multi-channel electrophysiological recordings (termed 'GCMR') from neurons in the antennal lobe (AL; the insect's primary olfactory center)13,14. This state-of-the-art technique allows us to probe how odor information is represented in the insect brain. Moreover, because neural responses to odors at this level of olfactory processing are highly sensitive owing to the degree of convergence of the antenna's receptor neurons into AL neurons, AL recordings will allow the detection of active constituents of natural odors efficiently and with high sensitivity. Here we describe GCMR and give an example of its use.
	Several general steps are involved in the detection of bioactive volatiles and insect response. Volatiles first need to be collected from sources of interest (in this example we use flowers from the genus Mimulus (Phyrmaceae)) and characterized as needed using standard GC-MS techniques14-16. Insects are prepared for study using minimal dissection, after which a recording electrode is inserted into the antennal lobe and multi-channel neural recording begins. Post-processing of the neural data then reveals which particular odorants cause significant neural responses by the insect nervous system.
	Although the example we present here is specific to pollination studies, GCMR can be expanded to a wide range of study organisms and volatile sources. For instance, this method can be used in the identification of odorants attracting or repelling vector insects and crop pests. Moreover, GCMR can also be used to identify attractants for beneficial insects, such as pollinators. The technique may be expanded to non-insect subjects as well.

Identification of Olfactory Volatiles using Gas Chromatography-Multi-unit Recordings GCMR in the Insect Antennal Lobe

Olfactory cues mediate many different behaviors in insects, and are often complex mixtures comprised of tens to hundreds of volatile compounds. Using gas chromatography with multi-channel recording in the insect antennal lobe, we describe a method for the identification of bioactive compounds.

All  organisms inhabit a world full of sensory stimuli that determine their  behavioral and physiological response to their environment. Olfaction is ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

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

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

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 ...

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila express gustatory receptors1,2, ion channels3-6, and ionotropic receptors7 that are involved in the detection of volatile and non-volatile sensory cues. These neurons directly contact tastants such as food, noxious substances and pheromones and therefore influence many complex behaviors such as feeding, egg-laying and mating. Electrode recordings and calcium imaging have been widely used in insects to quantify the neuronal responses evoked by these tastants. However, electrophysiology requires specialized equipment and obtaining measurements from a single taste sensillum can be technically challenging depending on the cell-type, size, and position. In addition, single neuron resolution in Drosophila can be difficult to achieve since taste sensilla house more than one type of chemosensory neuron. The live calcium imaging method described here allows responses of single gustatory neurons in live flies to be measured. This method is especially suitable for imaging neuronal responses to lipid pheromones and other ligand types that have low solubility in water-based solvents.

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

The forelegs and proboscis of Drosophila contain a rich repertoire of gustatory sensory neurons. Here, we present a method using calcium imaging to measure physiological responses from sensory neurons in the foreleg and proboscis of live flies upon exogenous application of a gustatory pheromone.

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila ...

Immunology and Infection

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, oviposition, and mating. Myriad chemicals of varying volatilities have been identified as natural odorants that activate insect Olfactory Receptor Neurons (ORNs). However, studying the olfactory responses to low-volatility odorants has been hampered by an inability to effectively present such stimuli using conventional odor-delivery methods. Here, we describe a procedure that permits the effective presentation of low-volatility odorants for in vivo Single-Sensillum Recording (SSR). By minimizing the distance between the odor source and the target tissue, this method allows for the application of biologically salient but hitherto inaccessible odorants, including palmitoleic acid, a stimulatory pheromone with a demonstrated effect on ORNs involved in courtship and mating behavior1. Our procedure thus affords a new avenue to assay a host of low-volatility odorants for the study of insect olfaction and pheromone communication.

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

The overall goal of this protocol is to demonstrate how to present odorants of low volatility for single-sensillum recording from Drosophila olfactory receptor neurons that respond to long-chain cuticular pheromones.

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, ...

Single Sensillum Recordings for Locust Palp Sensilla Basiconica

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially for the detection of non-volatile chemical cues through sensilla chaetica (previously named terminal sensilla or crested sensilla). There is now increasing evidence that these palps also have an olfactory function. An odorant receptor (LmigOR2) and an odorant-binding protein (LmigOBP1) have been localized in the neurons and accessory cells, respectively, in the sensilla basiconica of the palps. Single sensillum recording (SSR) is used for recording the responses of odorant receptor neurons, which is an effective method for screening active ligands on specific odorant receptors. SSR is used in functional studies of odorant receptors in palp sensilla. The structure of the sensilla basiconica located on the dome of the palps differs somewhat from the structure of those on the antennae. Therefore, when performing an SSR elicited by odorants, some specific advice may be helpful for obtaining optimum results. In this paper, a detailed and highly effective protocol for an SSR from insect palp sensilla basiconica is introduced.

This paper describes a detailed and highly effective protocol for single sensillum recordings from the sensilla basiconica on the palps of insect mouthparts.

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially ...

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

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

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

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

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

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.

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 ...

Electrophysiological Measurements from a Moth Olfactory System

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Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Identification of Olfactory Volatiles using Gas Chromatography-Multi-unit Recordings (GCMR) in the Insect Antennal Lobe

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

Using Insect Electroantennogram Sensors on Autonomous Robots for Olfactory Searches

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

Single Sensillum Recordings for Locust Palp Sensilla Basiconica

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

A Step-by-Step Guide to Mosquito Electroantennography