Soutenu par USDA-NIFA/AFRI 2010-65105-20582, NSF 0918177, et l&#39;incorporation de recherche Bedoukian.

<html> 1. Insectes <ol><li> Nombril orangeworm, Amyelois transitella (Walker) (Lepidoptera: Pyralidae) colonie dans notre laboratoire proviennent de mites recueillies auprès des amandiers à Fresno, en Californie. Notre colonie d&#39;insectes est maintenue dans l&#39;environnement des chambres (Percival Inc, IA) à 28 ± 2 ° C, 75 ± 10% d&#39;humidité relative, et sous 16h08 h (lumière: obscurité) photo régime. </li><li> Mites émergents sont collectées quotidiennement, sexué, et transféré à des boîtes en plastique (669 ml boîte à lunch, 13 x 13 cm, hauteur, 4,5 cm, Rubbermaid) fourni avec jusqu&#39;à 10 couches de serviettes papier imbibé d&#39;eau (Thirsty ultra absorbants, 27,9 x 27,9 cm ; Safeway). Couvre Box sont perforés pour permettre la circulation de l&#39;air. Cet arrangement permet ~ 100% HR. </li><li> Pour des mesures électrophysiologiques nous utilisons 2-4 jours après l&#39;émergence des papillons anciens, qui sont maintenus sous une forte humidité. </li></ol> 2. Préparation électrophysiologiques <ol><li> Mites (mâles ou femelles) sont doucement poussé dans une pipette (200 uL, USA Scientific Inc) qui est coupé par le haut pour avoir un diamètre de l&#39;extrémité de grande taille (environ 2 mm). Un papillon de nuit est doucement poussé par l&#39;autre bout avec un talon de papiers tissu humidifié jusqu&#39;au antennes et une partie de la tête dépasser de l&#39;extrémité. </li><li> Tête exposée est immobilisé en couvrant la tête avec de l&#39;argile non-séchage (Claytoon, Van Aken international, CA) en laissant une petite partie d&#39;un œil et deux antennes exposés. Cette préparation est placée sur une plate-forme de l&#39;EAG Micromanipulateur MP-12 (Syntech, Allemagne) entre les deux porte-électrodes. </li><li> Nous utilisons électrode en verre fabriqués à partir de 1,0 mm tube capillaire en borosilicate avec filament (WPI Inc, FL) et a tiré dans un modèle P-97 micropipette extracteur (Sutter Instruments, CA) pour obtenir des microélectrodes de moins d&#39;un micromètre de pointe. Les électrodes sont de retour rempli sensillum lymphatiques sonnerie (Kaissling et Thorson, 1980). Une électrode empale la surface exposée de l&#39;oeil sous le microscope (Leica MZ75) et sert de référence, tandis que l&#39;électrode d&#39;enregistrement accueille les deux antennes. Afin de faciliter le contact, distale 1-2 segments antennaires sont coupés avant d&#39;être inséré dans l&#39;électrode d&#39;enregistrement. Pour les signaux à partir des enregistrements individuels sensilles, nous avons légèrement modifier cette configuration dans laquelle l&#39;électrode d&#39;enregistrement est empalée sous le microscope (Olympus BX51WI; 800x de grossissement). Les potentiels d&#39;action sont enregistrés en insérant l&#39;électrode de verre à la base d&#39;un sensillum. Les signaux sont amplifiés par une haute impédance de pré-amplificateur (Syntech, Allemagne) et introduit dans une UB-IDAC être analysées hors ligne avec AUTOSPIKE logiciel (Syntech, Allemagne). Signaux AC sont filtrés passe-bande entre 100 -10 000 Hz et les potentiels d&#39;action sont extraits par ordinateur en utilisant un algorithme AUTOSPIKE qui distingue leur pic à creux amplitudes du bruit. Les réponses des neurones individuels sont calculés comme l&#39;augmentation (ou diminution) de la fréquence des potentiels d&#39;action (impulsions par seconde) sur la fréquence spontanée. </li></ol> 3. Stimuli chimiques et le protocole de stimulation <ol><li> Produits chimiques de haute pureté sont utilisés. Un des principaux constituants du mélange de phéromones femelles de la orangeworm nombril, (Z, Z) -11,13-hexadecadienal a été obtenu à partir de Bedoukian Research Inc, CT. Les produits chimiques sont dilués, p / v, avec du verre-hexane distillé pour faire le point sur les solutions dilutions pg / pl et 10 décimal sont faites. Une aliquote (10 pi) d&#39;un produit chimique de relance dissous dans l&#39;hexane est chargé sur une bande de papier filtre Whatman (8x40 mm), le solvant est évaporé en agitant pendant 10 s sous une hotte et la bande est placée dans un polypropylène 5 ml seringue (seringues BD, NJ) à partir de laquelle les différents volumes sont éjectés. Hexane seul et une seringue vide servir de témoin. </li><li> La préparation se déroule dans un flux d&#39;air humidifié délivré par le contrôleur de stimulation Syntech (CS-55 modèle; Syntech, Allemagne) à 20 ml / sec à laquelle une impulsion de stimulation de 4 mL / s est ajouté pendant 500 ms. Les signaux sont enregistrés pendant 10 s, 2 s de départ avant le début de l&#39;impulsion de stimulation. Préparation antennaire est stimulé avec 500 ms d&#39;impulsion au cours de laquelle ca. 2 ml de l&#39;air purifié à partir d&#39;une seringue en polypropylène 5 ml contenant le stimulus est ajouté sur le flux d&#39;air principal. Un écart d&#39;au moins 1 min ou plus, suivant les réponses de haut, est autorisé entre les stimulations. </li></ol> 4. Les résultats représentatifs Antennes de papillons retenue (figure 1) stimulées par composante phéromone femelle, (Z, Z) -11,13-hexadecadienal générer des réponses à la dose dépendante robustes (figure 2). Cette composante majeure de phéromone suscite également dose-dépendante les réponses excitatrices à long trochoïdes sensilles (figure 3A) tel qu&#39;il apparaît dans les traces d&#39;enregistrements unitaires (figure 3B). <img src="/files/ftp_upload/2489/2489fig1.jpg" alt="Figure 1" /> Figure 1. Live nombril papillon orangeworm sobre et exposé de l&#39;antenne load/2489/2489fig2.jpg &quot;alt =&quot; Figure 2 &quot;/&gt; Figure 2. Réponses à la dose dépendante EAG a suscité en réponse à (Z, Z) -11,13-hexadecadienal, un constituant majeur des phéromones femelles. <img src="/files/ftp_upload/2489/2489fig3.jpg" alt="Figure 3" /> Figure 3. Trichoid Sensilles chez le mâle nombril papillon orangeworm répondre à (Z, Z) -11,13-hexadecadienal de façon dose dépendante. (A) Antenne du mâle nombril orangeworm papillon est multi-segmenté et chaque segment est orné d&#39;un grand nombre de cheveux comme des structures, des sensilles. Un microscope électronique à balayage, montrant les détails dans l&#39;encart (barres d&#39;échelle sont de 200 et 50 uM, respectivement). (B) extracellulaire unitaire des enregistrements à partir d&#39;un sensillum trichoid.

<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>

Cette nouvelle méthode développée pour retenir un nombril papillon orangeworm direct pour mesurer des signaux olfactifs s&#39;est avéré être robuste et très fiable. Nous avons régulièrement recours à cette méthode pour isoler et identifier de nouveaux appâts à partir de substrats naturels comme l&#39;hôte d&#39;amandes et les pistaches.

<table border="1"><tbody><tr><td> Nom du produit </td><td> Catalogue # / Modèle </td><td> Société </td><td> Commentaire </td></tr><tr><td> Microscope </td><td> BX51WI modèle </td><td> Olympus, USA </td><td></td></tr><tr><td> Microscope stéréo </td><td> MZ75 </td><td> Leica Microsystems Inc USA </td><td></td></tr><tr><td> 1,0 mm borosilicate tubes capillaires avec des filaments </td><td> 1B100F-3 </td><td> WPI Inc, FL </td><td></td></tr><tr><td> Micropipette extracteur </td><td> P-97 </td><td> Sutter Instruments, CA </td><td></td></tr><tr><td> Stimulation du contrôleur </td><td> CS-55 modèle </td><td> Syntech, Allemagne </td><td></td></tr><tr><td> Haute impédance de pré-amplificateurs (Sonde universelle single ended) </td><td></td><td> Syntech, Allemagne </td><td></td></tr><tr><td> Amplificateur / acquisition de données système (USB-IDAC) </td><td></td><td> Syntech, Allemagne </td><td></td></tr><tr><td> Micromanipulateur EAG MP-12 </td><td></td><td> Syntech, Allemagne </td><td></td></tr><tr><td> (Z, Z) -11,13-hexadecadienal </td><td></td><td> Bedoukian Research Inc, CT. </td><td></td></tr><tr><td> Papier filtre Whatman </td><td> 1001070 </td><td> Whatman USA </td><td></td></tr><tr><td> 5 ml seringue en polypropylène </td><td> 309633 </td><td> Seringues BD, NJ </td><td></td></tr><tr><td> embout de pipette (200 uL) </td><td> 1111-0806 </td><td> USA Scientific Inc </td><td></td></tr><tr><td> 669 ml boîte à lunch, 13 x 13 cm, hauteur, 4,5 cm, </td><td></td><td> Rubbermaid </td><td></td></tr><tr><td> Assoiffé ultra absorbants, 27,9 x 27,9 cm </td><td></td><td> Safeway </td><td></td></tr><tr><td> Non desséchant d&#39;argile </td><td> 18150 </td><td> Claytoon, Van Aken international, CA </td><td></td></tr><tr><td> Chambre de l&#39;environnement </td><td> I-30BLL modèle </td><td></td><td></td></tr></tbody></table>

electrophysiological measurements from a moth olfactory system

Insecte systèmes olfactifs offrent des occasions uniques pour l&#39;enregistrement odorant réponses induites dans les formes de electroantennograms (EAG) et les enregistrements sensillum unique (SSR), qui sont résumées les réponses de tous les neurones récepteurs olfactifs (Orns) situé sur l&#39;antenne et de ceux logés dans différents sensilles, respectivement. Ces approches ont été exploitées pour obtenir une meilleure compréhension de la communication chimique des insectes. Les stimuli identifiés peuvent ensuite être utilisés soit comme attractifs ou répulsifs dans des stratégies de gestion pour les insectes ravageurs.

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

Electrophysiological Measurements from a Moth Olfactory System

Insecte systèmes olfactifs offrent des occasions uniques pour l&#39;enregistrement odorant réponses induites dans les formes de electroantennograms (EAG) et les enregistrements sensillum unique (SSR), qui sont résumées les réponses de tous les neurones récepteurs olfactifs (Orns) situé sur l&#39;antenne et de ceux logés dans différents sensilles, respectivement.

Mesures électrophysiologiques à partir d&#39;un système de Moth olfactive

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

electrophysiological-measurements-from-a-moth-olfactory-system

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

13.7K vues.  University of California, Davis. Insecte systèmes olfactifs offrent des occasions uniques pour l'enregistrement odorant réponses induites dans les formes de electroantennograms (EAG) et les enregistrements sensillum unique (SSR), qui sont résumées les réponses de tous les neurones récepteurs olfactifs (Orns) situé sur l'antenne et de ceux logés dans différents sensilles, respectivement.

Vidéo: 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).

Enregistrements Sensillum simple chez les insectes Drosophila melanogaster Et Anopheles gambiae</em

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

Recherche

Neurosciences

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.

L&#39;identification des composés volatils olfactifs chromatographie en phase gazeuse-à logements multiples enregistrements (GCMR) dans le lobe antennaire des insectes

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.

Méthodes d&#39;enregistrement multi-unités pour caractériser l&#39;activité neuronale dans le Criquet (<em&gt; Schistocerca americana</em&gt; Circuits olfactifs)

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.

Enregistrement électrophysiologique De<em&gt; Drosophila</em&gt; Labellar Goût Sensilles

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.

Utilisation d&#39;insectes électroantennogramme capteurs sur autonome Robots Recherches olfactives

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.

Mesurer les réponses physiologiques de<em&gt; Drosophila</em&gt; Neurones sensoriels à Lipid Phéromones Utilisation en direct d&#39;imagerie de calcium

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

Immunologie et 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.

Enregistrement électrophysiologique de<em&gt; Drosophila</em&gt; Sensilla Trichoid en Réponse aux Odorants de Faible Volatilité

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.

Sensille unique Recordings pour Locust Palp sensilles basiconiques

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.

Enregistrement électrophysiologique de l’activité du système nerveux Central du troisième stade de Drosophila Melanogaster

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.

Un guide étape par étape de l’électroanténnographie des moustiques

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 ...

Mesures électrophysiologiques à partir d'un système de Moth olfactive

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Un guide étape par étape de l’électroanténnographie des moustiques