The authors thank Mick Swann, Chris Askham and Martin Gautrey for their important contributions in designing and building many electrical and mechanical components of the rigs. MJ's current research is supported by the Biotechnology and Biological Sciences Research Council (BBSRC Grant: BB/M009564/1), the State Key Laboratory of Cognitive Neuroscience and Learning open research fund (China), High-End Foreign Expert Grant (China), Jane and Aatos Erkko Foundation Fellowship (Finland), and the Leverhulme Trust grant (RPG-2012-567).

The authors have nothing to disclose

We have presented the basic key steps of how to use sharp conventional microelectrodes to record intracellular responses of R1-R6 photoreceptors and LMCs in intact fly eyes. This method has been optimized, together with bespoke hardware and software tools, over the last 18 years to provide high-quality long-lasting recordings to answer a wide range of experimental questions. By investing time and resources to construct robust and precise experimental set-ups, and to produce microelectrodes with favorable electrical properties, high-quality recordings can become the norm in any laboratory working on Drosophila visual neurophysiology. Whilst well-designed recording and light stimulation systems are important for swift execution of different experimental paradigms, there are three procedural steps that are even more critical to achieving successful recordings: (i) to make the fly preparation with minimal eye damage, (ii) to pull microelectrodes with the right electrical properties, and (iii) to drive the recording electrode into the eye without breaking its tip. Ultimately, to record meaningful data, the investigator has to understand the physical basis of electrophysiology and how to fabricate suitable microelectrodes for the targeted cell-types.
Therefore, the limitations of this technique are primarily set by the patience, experience and technical ability of the investigator. Because this technique can take a long time to master for small Drosophila cells, it is advisable for trainee electrophysiologists to first practice with larger insect eyes, such as the blowfly36 or locust35, using the same rig. Once performing high-quality intracellular recordings from the larger photoreceptors and interneurons becomes routine, it is time to move on to the Drosophila eye. Another limitation of the technique concerns cellular identification. Penetrated Drosophila cells can be loaded electrophoretically with dyes, including Lucifer yellow or neurobiotin. However, because of the small tip size of the microelectrodes, electrophoresis works less efficiently than with lower resistance electrodes, such as patch-electrodes. Furthermore, the dye-filled microelectrodes characteristically have less favorable electrical properties, making it much harder to record high-quality responses with them from Drosophila photoreceptors and LMCs.
A technical problem that occurs sometimes is unstable input signal, or a complete lack of it. This is often associated with the voltage signal being either constantly drifting or higher/lower than the amplifier's recording range. On most occasions, this behavior is caused by the recording electrode being blocked (or its tip being too fine - having too high a resistance or intramural capacitance - to properly conduct fast signal changes). Although one can try to unblock the tip by buzzing the electrode capacitance, which sometimes works, often the situation is best resolved by simply changing the recording electrode. This may further require parameter adjustments in the microelectrode puller instrument to lower the tip resistance of the new electrodes. The electrode tip can also become blocked in preparations, for which it took too much time to cover the corneal hole by petroleum jelly. Prolonged air-contact can dry up the freshly exposed retinal tissue, turning its surface layer into a glue-like substance. If this is the case, the investigator typically sees a red blob of tissue stuck on the recording electrode when pulling it out of the eye. The only solution here is to make a new preparation. Petroleum jelly may provide many benefits for electrophysiological recordings: (i) it prevents the coagulation of the hemolymph that could break the electrode tip; (ii) it coats the electrode tip reducing its intramural capacitance, which lowers the electrode's time constant, and thus has the potential to improve the temporal resolution of the recorded neural signals40,41; (iii) it keeps the electrode tip clean, facilitating penetrations; and after penetration, (iv) it may even help to seal the electrode tip to the cell membrane42. 
The signal can further be unstable or lost when the silver-chloride wire of the electrode-holder is broken or dechloridized; in which case just replace or rechloridize the old wire. The missing signal can also result from one (or both) of the electrode-holders not being securely connected to their jacks. However, it is extremely unusual that a piece of equipment would be malfunctioning. If signal is undetectable and all other possibilities have been exhausted, test that each part of the recording apparatus, including the headstage, amplifier, low-pass filters and AD/DA-converters, are connected properly and functioning normally. One way to achieve this is to replace each instrument with another from a rig that is known to operate normally. Alternatively, use a signal generator to check the performance of the electronic components one by one.
But perhaps the most common technical problem facing the electrophysiologist is that of recording noise. Broadly, recording noise is the observed electrical activity other than the direct neuronal response to a given stimulus. Because the fly preparation, when properly done, is very stable, the observed noise (beyond the natural variably of the responses) most often results from ground-loops in the recording equipment, or is picked up from nearby electrical devices. Such noise is typically 50/60 Hz mains hum and its harmonics; but sometimes composed of more complex waveforms. To work out the origin of the noise, remove the fly preparation holder from the set-up, connect the recording and reference electrodes through a drop of fly Ringer (or place them in a small Ringer's solution bath; see step 1.2.6) and record the signal in CC- or bridge-mode. If noise is observable on the recorded signal, this likely means that the noise is external to the fly preparation.
Another good test for identifying the origin of noise is to replace the electrode-holders with an electric cell model connected to the amplifier. In an ideally configured and grounded set-up, the recorded signal should now be practically noise-free, showing only stochastic bit-noise from the AD-converter (in the best case not even that!). If noise is still present, then recheck that all rig equipment is properly grounded. A convenient approach to detect ground-loops is to: (i) disconnect all the grounding wires from all the parts within the rig; (ii) ensure that, after doing this, every single part is actually isolated from ground, by means of an ohm-meter; (iii) connect the parts, one by one, to the central ground directly, not through any other part of the rig. Try also changing the equipment configurations. For example, sometimes moving the computer and monitor further away from the rig can reduce noise; yet at other times, moving the computer inside the equipment rack reduces noise. It is also worth unplugging nearby equipment to see if noise is reduced, or shield additional components. Furthermore, try unplugging or replacing different components of the recording equipment, especially BNC cables (which can have faulty ground connections). If only bit-noise is observed when using the cell model, the initial noise source is either the electrodes or the fly preparation itself. For example, it could be that the reference electrode is inadvertently touching a motor nerve or active muscle fibers inside the head capsule (or disturbing flight muscles in the thorax - if placed there). It is usually simplest to prepare a new fly for recording, taking care to minimize damage to the fly. But if the noise persists and is broadband, it is likely that the electrodes are suboptimal for the experiments; too sharp/fine (hence too noisy) or just wrong for the purpose; we have even seen quartz-electrodes acting as antennas - picking up faint broadcasting signals! Although iteration of the puller-instrument parameter settings to generate the just right microelectrodes for consistent high-quality recordings from specific cell-types can take a lot of effort, it is worth it. Once the recording electrodes are well-tailored for the experiments, they can provide long-lasting recordings of outstanding quality.
Sharp microelectrode recording techniques can be similarly applied to study neural information processing in multitude of preparations, including different processing layers in the insect eyes and brain43,44. Because the microelectrode tips can be made very fine, these typically damage the studied cells less than most patch-clamp applications. Importantly, the modern sample-and-hold microelectrode amplifiers enable good control of the tips' electrical properties40,45-47. Thus, when correctly applied, this technique can provide reliable data from both in vivo3,5,7-10,44 or in vitro48 preparations with high signal-to-noise ratio at sub-millisecond resolution. Such precision would be impossible with today's optical imaging techniques, which are noisier and slower. Moreover, the method can be used to characterize small cells' electrical membrane properties both in current- and voltage-clamp configurations5,29,33,36,40-42,49, providing valuable data for biophysical and empirical modeling approaches7,8,11,33,49-54 that link experiments to theory.

La mouche des fruits (Drosophila melanogaster) composé oeil est un système modèle grand pour enquêter sur l&#39;organisation fonctionnelle des photorécepteurs et interneurones tableaux pour l&#39; image neuronal échantillonnage et le traitement, et pour la vision des animaux. Le système a le schéma de câblage le plus complet 1,2 et est aimable aux manipulations génétiques et précise la surveillance de l&#39; activité neuronale (de haut rapport signal-bruit et résolution temporelle) 3-10. L&#39;œil de drosophile est modulaire, contenant ~ 750 structures de lentilles coiffées apparemment régulières appelées ommatidia, qui fournissent ensemble le champ visuel panoramique qui couvre presque toutes les directions autour de sa tête volée. L&#39; information primaire de l&#39;œil unités d&#39; échantillonnage sont ses photorécepteurs rhabdomeric 7,8,11. Chaque ommatidium contient huit cellules photoréceptrices (R1 à R8), qui partagent le même objectif de facette, mais sont alignés sur sept directions différentes. Alors que les photorécepteurs externes R1-R6 are plus sensible à la lumière bleu-vert, les sensibilités spectrales des cellules internes R7 et R8, qui se situent au - dessus de l&#39;autre et pointer vers la même direction, présentent trois sous - types distincts: pâle, zone de bord jaune et dorsale (DRA) 12- 15. <img alt="Figure 1" src="/files/ftp_upload/54142/54142fig1.jpg" /> Figure 1. Organisation fonctionnelle du Drosophila Eye. (A) Les deux premiers noyaux optiques, rétine et lamina, sont mis en évidence en gris dans l&#39;œil de mouche. Photorécepteurs Retina R1-R6 et lamina grandes cellules monopolaires (LMC: L1-L3) sont facilement accessibles in vivo à des enregistrements classiques de microélectrodes pointus. L&#39;électrode schématique montre le trajet normal à enregistrer à partir de R1 à R6 dans la rétine. Un chemin d&#39;accès à enregistrer à partir de LMC dans le limbe est de déplacer en parallèle l&#39;électrode à gauche. (B) Lamina est une matrice d&#39;organe rétinotopiquecartouches lisés, dont chacun est emballé avec les neurones qui traite des informations à partir d&#39;une petite zone spécifique dans l&#39;espace visuel. En raison de la superposition de neurones, six photorécepteurs de différents ommatidia voisins envoient leurs axones (R1-R6) à la même cartouche de lame, formant des synapses de sortie histaminergiques à L1 à L3 et d&#39;une cellule amacrine (Am). (C) La diffusion de l&#39; information neuronale entre les terminaisons axonales R1-R6 et les interneurones visuels (y compris L4, L5, Lawf, C2, C3 et T1), à l&#39; intérieur d&#39; une cartouche de lamina est complexe. (D) R1-R6 axones photoréceptrices reçoivent des évaluations synaptiques de cellules monopolaires L2 et L4. (B) et (C) modifié de Rivera-alba et al 2. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig1large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande de cette figure.</a> L&#39;œil de drosophile est du type neuronal superposition 16. Cela signifie que tchapeau les signaux neuronaux de huit photorécepteurs appartenant à sept ommatidia voisin, qui ressemblent au même point dans l&#39;espace, sont réunies ensemble à une cartouche de neurones dans les deux prochaines neuropils: le limbe et la moelle. Tandis que les six photorécepteurs externes du projet R1-R6 leurs terminaisons axonales de neurones à des colonnes dans la feuille (figure 1), les cellules R7 et R8 contourner cette couche et à établir des contacts synaptiques avec leur colonne rachidien 17-19 correspondante. Ces câblages exactes produisent le substrat neuronal pour la cartographie rétinotopique de mouche vision précoce, après quoi tous les lamina (Figures 1A-C) et de la colonne medulla (cartouche) représente un seul point dans l&#39; espace. Les apports directs de photorécepteurs R1-R6 sont reçus par les grandes cellules monopolaires (de LMC: L1, L2 et L3) et la Cellule amacrines (Am) dans la lamina 1,2,20. Parmi ceux - ci, L1 et L2 sont les plus grandes cellules, médiateurs principales voies d&#39;information (figure 1D), which répondent à l&#39; intérieur et à bords Off-déplacement, et forment ainsi la base de calcul du détecteur de mouvement 21,22. Expériences comportementales suggèrent qu&#39;au contraste intermédiaire, les deux voies facilitent la perception du mouvement des directions opposées: back-to-front en L1 et avant-arrière dans les cellules L2 23,24. Connectivité implique en outre que les neurones L4 peuvent jouer un rôle essentiel dans la communication latérale entre les cartouches voisines 25,26. synapses réciproques ont été trouvées entre les cellules L2 et L4 situées dans le même et deux cartouches adjacentes. En aval, chaque cellule L2 et ses trois cellules L4 associés projettent leurs axones vers un objectif commun, le neurone Tm2 dans le bulbe rachidien, où les entrées de cartouches voisines sont soupçonnés d&#39;être intégré pour le traitement de l&#39; avant vers l&#39; arrière le mouvement 27. Bien que les neurones L1 reçoivent des commentaires du même cartouche L2 via les jonctions lacunaires et synapses, ils ne sont pas directement reliés aux cartouches lamina L4S et donc adjacents. <pclass = &quot;jove_content&quot;&gt; évaluations synaptiques à R1-R6 axones photoréceptrices sont fournis uniquement par les neurones appartenant aux circuits / L4 L2 mais pas la voie 1,2 L1 (figure 1D). Tandis que les connexions de même cartouche sont sélectivement L2 R1 et R2 et de R5 à L4, les photorécepteurs R1-R6 synaptiques reçoivent une rétroaction à partir L4 de l&#39;une ou les deux cartouches voisines. En outre, il y a de fortes connexions synaptiques de Am à R1, R2, R4 et R5, et les cellules gliales sont également synaptique connectés au réseau et peuvent ainsi participer au traitement de l&#39; image neuronal 6. Enfin, gap-jonctions axonales, reliant voisins R1-R6 et entre R6 et R7 / R8 photorécepteurs dans la lamina, contribuent à la représentation de l&#39; asymétrie d&#39; information et de traitement dans chaque cartouche 14,20,28. Enregistrements de tension intracellulaires de photorécepteurs individuels et des interneurones visuels dans presque intact Drosophila offrent une haute r signal-bruitdonnées atio à une résolution inférieure à la milliseconde 3,5,7-10,29, qui est nécessaire pour donner un sens aux calculs neuronaux rapides entre les neurones connectés. Ce niveau de précision est impossible par des techniques d&#39;imagerie optique actuelles, qui sont nettement plus bruyants et opèrent généralement à 10-100 résolution msec. En outre, parce que les électrodes ont de très petits et pointus conseils, la méthode ne se limite pas à des organismes cellulaires, mais peut fournir des enregistrements directs de petites structures neurales actives; tels que des arbres dendritiques des LMC ou axones photoréceptrices, qui ne peuvent être accessibles par beaucoup plus grandes pointes des électrodes de patch-clamp. Fait important, la méthode est aussi structurellement moins invasive et dommageable que la plupart des applications de patch-clamp, et affecte donc moins intracellulaire milieu et de l&#39;information d&#39;échantillonnage »les cellules étudiées. Ainsi, les techniques classiques de microélectrodes nettes ont contribué, et de continuer à contribuer, les découvertes fondamentales et un aperçu original dans infor neuronalle traitement de l&#39;infor- à l&#39;échelle de temps appropriée; améliorer notre compréhension des mécanismes de la vision 10/03. Cet article explique comment les enregistrements intracellulaires in vivo de la drosophile photorécepteurs et LMC R1-R6 sont effectuées dans le laboratoire Juusola. Ce protocole décrit comment construire une plate-forme d&#39;électrophysiologie approprié, préparer la mouche, et effectuer les enregistrements. Certaines données représentatives sont présentées, et des problèmes communs et les solutions possibles sont discutées qui peuvent être rencontrées lors de l&#39;utilisation de cette méthode.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Stereo Zoom Microscope for making the fly preparation</td>
			<td>Olympus</td>
			<td>SZX12 DFPLFL1.6x PF eyepieces: WHN30x-H/22</td>
			<td>Capable of ~150X magnification with long working distance; bespoke heavy steel table mount stand</td>
		</tr>
		<tr>
			<td rowspan="2">Stereomicroscope in the intracellular set-up</td>
			<td>&#183;&#160; Olympus</td>
			<td>Olympus SZX7; eyepieces: WHN30x-H/22</td>
			<td rowspan="2">30x eyepieces are needed for seeing the electrode tip reflections well when driving it through the small corneal hole into the eye</td>
		</tr>
		<tr>
			<td>&#183;&#160; Nikon</td>
			<td>Nikon SMZ645; eyepieces: C-W30x/7</td>
		</tr>
		<tr>
			<td rowspan="2">Anti-vibration Table</td>
			<td>&#183;&#160; Melles Griot</td>
			<td rowspan="2">With metric M6 holes on the breadboard</td>
			<td rowspan="2">Our bespoke rigs have a large hole drilled through the thick breadboard that lets in the fly preparation platform pole (houses a copper heatsink with electronics) from below</td>
		</tr>
		<tr>
			<td>&#183;&#160; Newport</td>
		</tr>
		<tr>
			<td rowspan="4">Micromanipulators</td>
			<td>&#183;&#160; Narishige</td>
			<td>&#183;&#160; Narishige NMN-21</td>
			<td rowspan="4">In our intracellular set-ups, different micromanipulator systems are used for driving the shap recording electrodes into the fly eye. All the listed manipulators are succesfully providing long-lasting stable recordings from Drosophila photoreceptors and LMCs.&#160;</td>
		</tr>
		<tr>
			<td>&#183;&#160; Huxley Bertram</td>
			<td>&#183;&#160; Huxley xyz-axis with fine manual control</td>
		</tr>
		<tr>
			<td>&#183;&#160; Sensapex</td>
			<td>&#183;&#160; Sensapex triple axis</td>
		</tr>
		<tr>
			<td>&#183;&#160; M&#228;rzh&#228;user</td>
			<td>&#183;&#160; M&#228;rzh&#228;user DC-3K with additional x-axis piezo stepper and MS 314 controller</td>
		</tr>
		<tr>
			<td>Magnetic Stands</td>
			<td>Any magnetic base with on/off switch will do</td>
			<td></td>
			<td>For example, to manage cables inside the Faraday cage</td>
		</tr>
		<tr>
			<td>Electrode Holders</td>
			<td>Harvard Apparatus</td>
			<td>ESP/W-F10N</td>
			<td></td>
		</tr>
		<tr>
			<td>Silver Wire</td>
			<td>World Precision Instruments&#160;</td>
			<td>AGW1510</td>
			<td>0.3-0.5 mm diameter; needs to be chloridized for the electrode holders</td>
		</tr>
		<tr>
			<td>Fiber Optic Light Source</td>
			<td>Many different, including Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td rowspan="2">Fiber Optic Bundles</td>
			<td>&#183;&#160; UltraFine Technology</td>
			<td rowspan="2"></td>
			<td rowspan="2">To deliver the LED light stimulus to the Cardan arm system. We use both liquid and quartz light guides (range from UV to IR)</td>
		</tr>
		<tr>
			<td>&#183;&#160; Thorn Labs</td>
		</tr>
		<tr>
			<td>Fly Cathing Tube</td>
			<td></td>
			<td>P80-50P 50ml Cent. Tube PP., Pack of 100 Pcs</td>
			<td>Cut the conical bottom off from 50 ml Plastic Centrifuge Tube and glue a 1 ml pipette tip on it.</td>
		</tr>
		<tr>
			<td>Digital Acquisition System</td>
			<td>National Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Single-electrode current/voltage-clamp microelectrode amplifier</td>
			<td>npi SEC-10LX</td>
			<td>http://www.npielectronic.de/products/amplifiers/sec-single-electrode-clamp/sec-10lx.html</td>
			<td>Outstanding performer!</td>
		</tr>
		<tr>
			<td>Head-stage</td>
			<td>Standard (+/- 150 nA)</td>
			<td>For npi SEC-10LX</td>
			<td></td>
		</tr>
		<tr>
			<td rowspan="2">LED light sources and drivers</td>
			<td>&#183;&#160; 2-channel OptoLED (Cairn Research Ltd., UK)</td>
			<td rowspan="2"></td>
			<td rowspan="2">Many of our stimulus systems are in-house built&#160;</td>
		</tr>
		<tr>
			<td>&#183;&#160; Self-designed and constructed</td>
		</tr>
		<tr>
			<td>Acquisition and Analyses Software</td>
			<td>Many companies to choose from</td>
			<td></td>
			<td>Biosyst; custom written Matlab-based system for experimental and theoretical work in the Juusola laboratory</td>
		</tr>
		<tr>
			<td>Personal Computer or Mac</td>
			<td></td>
			<td></td>
			<td>Ensure that PC or Mac is compatible with data acquisition system and software</td>
		</tr>
		<tr>
			<td>Cardan arm system&#160;</td>
			<td>Self-designed and constructed</td>
			<td></td>
			<td>Providing accurate x,y,z-positioning of the light stimuli</td>
		</tr>
		<tr>
			<td>Peltier temperature control system&#160;</td>
			<td>Self-designed and constructed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Faraday Cage</td>
			<td>Self-constructed</td>
			<td></td>
			<td>Electromagnetic noise shielding</td>
		</tr>
		<tr>
			<td rowspan="2">Filamented Borosilicate Glass Capillaries</td>
			<td rowspan="2"></td>
			<td>Outer diameter: 1 mm</td>
			<td rowspan="2"></td>
		</tr>
		<tr>
			<td>Inner diameter: 0.5-0.7 mm</td>
		</tr>
		<tr>
			<td rowspan="2">Filamented Quartz Glass Capillaries</td>
			<td rowspan="2"></td>
			<td>Outer diameter: 1 mm</td>
			<td rowspan="2"></td>
		</tr>
		<tr>
			<td>Inner diameter: 0.5-0.7 mm</td>
		</tr>
		<tr>
			<td>Pipette Puller</td>
			<td>Sutter Instrument Company</td>
			<td>Model P-2000 laser Flaming/Brown Micropipette Puller</td>
			<td>For borosilicate reference electrodes, use the preset program #11 (patch electrodes): Heat = 350; Filament = 4; Velocity 36; Delay = 200).1.2.1). For borosilicate recording electrodes, use the preset program #12 (this typically pulls good conventional sharps&#160; for photoreceptor recordings): Heat = 355; Filament = 4; Velocity 50; Delay = 225; Pull = 150. For LMC recordings, which require electrodes with finer tips, these values need to be adjusted. For pulling quartz capillaries, P-2000 manual suggests programs for fine tipped microelectrodes. These programs&#8217; preset parameters serve as useful starting points for systematic modifications to generate electrodes with good penetration success and low recording noise.</td>
		</tr>
		<tr>
			<td>Extracellular Ringer Solution for the reference electrode</td>
			<td>Chemicals from Fisher Scientific</td>
			<td>10326390, NaCl 10010310, KCl 10147753, TES 10161800, CaCl2 10159872, MgCl2 10000430, sucrose</td>
			<td>See the recipe in the protocol section</td>
		</tr>
		<tr>
			<td>3 M KCl solution for filling the filamented recording microelectrode</td>
			<td>Salts from Fisher Scientific</td>
			<td>10010310, KCl</td>
			<td></td>
		</tr>
		<tr>
			<td>Petroleum jelly</td>
			<td>Vaselin</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Non-stainless steel razor blades</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Blade holder/breaker</td>
			<td>Fine Science Tools By Dumont</td>
			<td>10053-09</td>
			<td>9 cm</td>
		</tr>
		<tr>
			<td>Blu-tack</td>
			<td>Bostik</td>
			<td></td>
			<td>Alternatively, use molding clay</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fine Science Tools By Dumont</td>
			<td>11252-00</td>
			<td>#5SF (super-fine tips)</td>
		</tr>
	</tbody>
</table>

electrophysiological method for recording intracellular voltage responses of drosophila photoreceptors and interneurons to light stimuli in vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

La méthode d&#39;enregistrement de microélectrodes forte, adaptée ici pour l&#39;œil de drosophile, peut être utilisé pour quantifier de façon fiable neuronal échantillonnage de l&#39; information et de traitement dans les cellules de la rétine et de la lamina, et la communication entre les 4,5,7,8,10,33. En l&#39;utilisant pour étudier le codage dans différentes actions de type sauvage, des mutants ou des souches de mouches génétiquement modifiées, la méthode a fait ses preuves; non seulement pour quantifier les effets d&#39;une mutation, la température, l&#39; alimentation ou l&#39; expression sélectionnée 3,4,6,9,10,14,30,34, mais aussi en révélant des raisons mécanistes pour les comportements visuels altérés 14,34. Le procédé est également facilement applicable à d&#39; autres préparations d&#39;insectes, 35,36 habilitation des études de vision neuroethological. Ensuite, nous présentons quelques exemples de ses applications réussies. <img alt="Figure 7" src="/files/ftp_upload/54142/54142fig7.jpg" /> Figure 7 TensionRéponses d&#39;un Fruit Fly R1-R6 photorécepteur à une impulsion de lumière à 20 et 25 o C. Parce que les pénétrations de microélectrodes vives sont souvent très stable, il est possible d&#39;enregistrer les réponses de tension de la même photorécepteur R1-R6 à un stimulus lumineux donné au différentes températures ambiantes par le réchauffement ou le refroidissement de la mouche. Dans nos set-ups, le porte-mouche est placé sur un système de contrôle de température à base Peltier-élément en boucle fermée. Cela nous permet de changer la température de la tête de la mouche en quelques secondes. Une température plus élevée accélère les réactions de tension et abaisse le potentiel de repos de photorécepteurs R1-R6 (comme indiqué par des flèches rouges) caractéristique. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig7large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande de cette figure.</a> L&#39; étude de l&#39;effet de la température sur photorécepteur sortie <p class="jove_content" fo:kEEP-together.within-page = &quot;1&quot;&gt; Avec un bien conçu et un système d&#39;enregistrement isolé des vibrations, la méthode peut être utilisée pour mesurer l&#39;effet de la température sur la production de neurones d&#39;une cellule individuelle par le réchauffement ou le refroidissement de la mouche. L&#39;exemple suivant montre les réponses de tension à une impulsion de 10 msec de temps lumineux, enregistré dans le même photorécepteur R1 à R6 et 20 à 25 ° C (figure 7). Comme quantifiée avant 4,9, le réchauffement réduit le potentiel de repos d&#39;un photorécepteur dans l&#39; obscurité, et accélère ses réponses de tension. <img alt="Figure 8" src="/files/ftp_upload/54142/54142fig8.jpg" /> Figure 8. Signalisation Performance d&#39; un Fruit Fly R1-R6 photorécepteur améliore avec l&#39; intensité lumineuse (A) Sortie de photorécepteur à dim (ci - dessous) et lumineux. (Ci - dessus; 10.000 fois la lumière plus brillante) répétées séries temporelles d&#39;intensité lumineuse naturaliste enregistrée par le même microélectrodesdans la même cellule à 20 ° C. Les réponses au stimulus lumineux sont plus grandes, parce qu&#39;ils intègrent plus d&#39; échantillons, des réponses élémentaires (bosses) à photons uniques 4,5,7,8. (B) 20 réponses consécutives de tension d&#39; une seconde-longues sont superposées. Les réponses individuelles (lumière gris) ont été prises après les tendances adaptatives (flèche en A) a reculé (encadré en pointillés dans A). Les moyens de réponse correspondants (les signaux) sont les traces foncées. La différence entre le signal et les réponses individuelles est le bruit. (C) Les cellules «performances de signalisation a été quantifiée par les enregistrements de signal-bruit Ratios (SNR) en utilisant les méthodes standard 4,5,7,8. Sortie photorécepteur a une portée d&#39; environ 64 Hz plus large de la signalisation fiable à la stimulation lumineuse ( &#39;de ≥1 vive, jusqu&#39;à 84 Hz qu&#39;au dim (SNR SNR)&#39; Dim ≥1, jusqu&#39;à 20 Hz), avec s-Ignal à bruit en améliorant grandement; de SNR Dim MAX = 87 à SNR brillant MAX = 1,868. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig8large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande de cette figure.</a> Etudier l&#39; adaptation et Neural Encoding par Répétitive Stimulation La non - invasivité du procédé, ce qui provoque relativement peu de dommages aux structures de la rétine et de la lame, il est idéal pour étudier les performances de la signalisation des cellules individuelles à différents stimuli lumineux dans leur état ​​physiologique naturel près in vivo. La figure 8 montre les réponses de tension d&#39;un R1- R6 photorécepteur à un naturaliste temps d&#39;intensité lumineuse série stimulus répété faible et lumineux à 20 ° C, tandis que la figure 9 </strong&gt; montre les réponses d&#39; une autre photorécepteur R1-R6 et un LMC à un stimulus naturaliste différent à 25 o C. Les enregistrements pré et post - synaptiques ont été réalisées séparément de deux mouches différentes parce que les enregistrements intracellulaires simultanés par deux tranchants microélectrodes dans la même volée, l&#39; une dans la rétine et l&#39;autre dans la lamina, sont trop difficiles pour être viable 30. <img alt="Figure 9" src="/files/ftp_upload/54142/54142fig9.jpg" /> Figure 9. Réponses de tension d&#39;un Fruit Fly R1-R6 photorécepteur et LMC à répétition Naturalistic Stimulation à 25 o C (A) R1-R6 (gris) et LMC (noir) sorties enregistrées par microélectrodes différentes de différentes mouches. (B) 20 avant entièrement la lumière adaptée consécutive (ci - dessus) et post - synaptiques (ci - dessous) les réponses au même schéma de relance naturaliste avec des réponses individuelles, représenté en gris clair und les moyens correspondants de réponse (les signaux) que les traces les plus sombres. La différence entre le signal et les réponses individuelles est le bruit. (C) Les cellules «performances de signalisation a été quantifiée par les enregistrements de signal-bruit Ratios (SNR). Sortie LMC a une portée d&#39; environ 10 Hz plus large de la signalisation fiable (SNR &#39;LMC ≥1, jusqu&#39;à 104 Hz) que la sortie R1-R6 (SNR&#39; R ≥1, jusqu&#39;à 94 Hz). Les deux rapports signal sur bruit est élevé (SNR LMC MAX = 142, SNR R MAX = 752), et comme le bruit d&#39;enregistrement était faible, leurs différences reflètent des différences réelles de codage entre les cellules. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig9large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande cette figure.</a> Après le début de relance, les enregistrements typiquesly montrent l&#39;adaptation rapide des tendances qui s&#39;atténuent largement dans 5-6 sec. A partir de là, les cellules produisent des réponses très cohérentes à chaque présentation à long stimulus 1 sec (chaque boîte en pointillés entoure 20 de ceux-ci). La reproductibilité des réponses devient évidente lorsque celles - ci sont superposées (figure 8B et figure 9B). Les réponses individuelles sont les traces grises minces, et de leur dire la trace sombre plus épaisse. La réponse moyenne est pris comme signal de neurones, alors que le bruit de neurones est la différence entre la moyenne et chaque 4,5,9,37,38 de réponse individuelle. Les rapports respectifs signal-bruit dans le domaine fréquentiel (figure 8C et la figure 9C) ont été obtenus par transformation de Fourier du signal et de bruit blocs de données dans les spectres de puissance, et en divisant le spectre de puissance moyenne du signal avec le spectre de puissance de bruit moyenne correspondante 4, 5,9,37,38. De façon caractéristique, les rapports maximaux signal-bruit of les sorties neuronaux enregistrées à la stimulation naturaliste sont élevés (100 - 1000), et dans les préparations les plus stables avec le bruit d&#39;enregistrement très bas peut atteindre des valeurs &gt;&gt; 1000 (par exemple, la figure 8C.). Notez également que le réchauffement augmente les cellules de la bande passante de données fiables de signalisation 4 (SNR Bright ≥ 1); par exemple, la différence relative entre les deux R1- les R6 sur les figures 8 et 9, respectivement, est de 10 Hz (84 à 20 ° C et 94 Hz à 25 ° C). On peut en outre estimer le taux de transfert d&#39;information de son rapport signal sur bruit de chaque cellule en utilisant l&#39;équation Shannon 32, ou en calculant la différence entre l&#39; entropie et le bruit des taux d&#39;entropie des réponses grâce à la méthode d&#39;extrapolation triple 39. Plus de détails sur les informations des analyses théoriques, et leur utilisation et limitations spécifiquement avec cette méthode sont donnés dans les publications antérieures 7,8,39. <img alt="Figure 10" src="/files/ftp_upload/54142/54142fig10.jpg" /> Figure 10. Réponses de tension d&#39;un tueur Fly R1-R6 photorécepteur et LMC à répétition Naturalistic Stimulation à 19 o C (A) R1-R6 (gris) et LMC (noir) sorties enregistrées par le même microélectrodes de la même volée; postsynaptique premier et ensuite présynaptique, que l&#39;électrode a été avancé dans l&#39;oeil. (B) 20 pré consécutifs (ci - dessus) et post - synaptiques (ci - dessous) Réponses (traces gris clair) pour le même motif de stimulus naturaliste ont été capturés après l&#39; adaptation initiale (boîte en pointillés dans A). Leurs moyens sont les signaux (les traces sombres sur le dessus), tandis que leurs différences respectives aux réponses individuelles donnent le bruit. (C) correspondant SignaLes ratios l-bruit (SNR) ont été calculées comme dans les figures 8C et 9C. Sortie LMC a environ une plage de 100 Hz plus large de signalisation fiable (SNR &#39;LMC MAX ≥1, jusqu&#39;à 234 Hz) que la sortie R1-R6 (SNR&#39; R MAX ≥1, jusqu&#39;à 134 Hz). Les deux rapports signal sur bruit est élevé (SNR LMC MAX = 137, SNR R MAX = 627), et que le même microélectrodes a été utilisé dans les enregistrements, leurs différences reflètent des différences réelles dans les sorties de neurones pré- et post - synaptiques. Ces résultats impliquent que le système d&#39;enregistrement avait peu de bruit, et son influence sur les analyses était marginal. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig10large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande de cette figure.</a> Neuroethological études Vision </p&gt; Le procédé peut également être utilisé pour enregistrer les réponses pré- et post - synaptiques tension des yeux composés de différentes espèces d&#39;insectes 7,8,35,36 (figure 10), ce qui permet des études comparatives neuroethological de traitement de l&#39; information visuelle. Pour le système d&#39;enregistrement présenté, la seule adaptation nécessaire est-détenteurs de préparation de nouvelles, chacune avec une ouverture de taille appropriée pour les espèces étudiées. Ces enregistrements sont exemplaires à partir d&#39;un tueur femelle voler (Coenosia de attenuata). Ils montrent des réponses de tension intracellulaire d&#39;un photorécepteur R1 à R6 et LMC à une stimulation lumineuse répétitif identique, que celle utilisée pour les homologues de Drosophila de la figure 9, mais à 19 ° C Dans ce cas, à la fois pré- et post-synaptiques des données ont été enregistrées à la même volée; une après l&#39;autre, avec la même électrode d&#39;enregistrement (rempli de 3 M de KCl) avançant à travers la première lame latérale avant de la saisirment la rétine frontale. En comparaison avec les données de drosophile à 25 o C, les données Coenosia - même à la température plus froide - montre les réponses avec une dynamique plus rapide; élargir la gamme de signalisation fiable (rapport signal-bruit &gt;&gt; 1) sur une gamme de fréquences plus large. De telles adaptations fonctionnelles dans le codage neuronal des stimuli naturalistes sont compatibles avec le mode de vie prédateur de Coenosia 36, qui exigent de haute précision des informations spatiotemporelle pour atteindre les comportements de chasse aériens rapides.

Watch this Scientific Journal Video about Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo at JoVE.com

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Méthode électrophysiologique de réponses d&#39;enregistrement intracellulaire tension de<em&gt; Drosophila</em&gt; Photorécepteurs et Interneurones à Light Stimuli<em&gt; In Vivo</em

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

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Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

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18.5K vues.  Beijing Normal University. Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Vidéo: Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Deux enregistrements électrophysiologiques des synapses-réponses évoquées astrogliales et neuronales aiguës dans des coupes d&#39;hippocampe

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

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A Method for Systematic Electrochemical and Electrophysiological Evaluation of Neural Recording Electrodes

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant characteristics.&#160;This precludes a rational selection of a particular implant as optimal for a particular application and the development of new materials based on the most critical performance parameters.&#160;This article develops a protocol for in vitro and in vivo testing of neural recording electrodes.&#160;Recommended parameters for electrochemical and electrophysiological testing are documented with the key steps and potential issues discussed.&#160;This method eliminates or reduces the impact of many systematic errors present in simpler in vivo testing paradigms, especially variations in electrode/neuron distance and between animal models.&#160;The result is a strong correlation between the critical in vitro and in vivo responses, such as impedance and signal-to-noise ratio.&#160;This protocol can easily be adapted to test other electrode materials and designs.&#160;The in vitro techniques can be expanded to any other nondestructive method to determine further important performance indicators.&#160;The principles used for the surgical approach in the auditory pathway can also be modified to other neural regions or tissue.

Different electrode coatings affect neural recording performance through changes to electrochemical, chemical and mechanical properties. Comparison of electrodes in vitro is relatively simple, however comparison of in vivo response is typically complicated by variations in electrode/neuron distance and between animals. This article provides a robust method to compare neural recording electrodes.

Une méthode pour électrochimique systématique et l&#39;évaluation électrophysiologique des neurones électrodes d&#39;enregistrement

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant ...

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 ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

Un guide pour<em&gt; In vivo</em&gt; Simple unité d&#39;enregistrement de Optogenetically identifié corticale inhibitrice Interneurones

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at the synaptic level. The Active Zone (AZ) is the region at the presynaptic membrane that mediates neurotransmitter release and is composed of a dense collection of scaffold proteins. AZs of Drosophila melanogaster (Drosophila) photoreceptors undergo molecular remodeling after prolonged exposure to natural ambient light. Thus the level of neuronal activity can rearrange the molecular composition of the AZ and contribute to the regulation of the functional output.
Starting from the light exposure set-up preparation to the immunohistochemistry, this protocol details how to quantify the number, the spatial distribution, and the delocalization level of synaptic molecules at AZs in Drosophila photoreceptors. Using image analysis software, clusters of the GFP-fused AZ component Bruchpilot were identified for each R8 photoreceptor (R8) axon terminal. Detected Bruchpilot spots were automatically assigned to individual R8 axons. To calculate the distribution of spot frequency along the axon, we implemented a customized software plugin. Each axon&#39;s start-point and end-point were manually defined and the position of each Bruchpilot spot was projected onto the connecting line between start and end-point. Besides the number of Bruchpilot clusters, we also quantified the delocalization level of Bruchpilot-GFP within the clusters. These measurements reflect in detail the spatially resolved synaptic dynamics in a single neuron under different environmental conditions to stimuli.

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

Here we show how to quantify the number and spatial distribution of synaptic active zones in Drosophila melanogaster&#160;photoreceptors, highlighted with a genetically encoded molecular marker, and their modulation after prolonged exposure to light.

Analyse Synaptic Modulation de<em&gt; Drosophila melanogaster</em&gt; Photorécepteurs après exposition à la lumière prolongée

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at ...

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, enabling the use of D. melanogaster molecular genetics to study inositol-lipid signaling and Transient Receptor Potential (TRP) channels at the single-molecule level. A handful of labs have mastered this powerful technique, which enables the analysis of the physiological responses to light under highly controlled conditions. This technique allows control over the intracellular and extracellular media; the membrane voltage; and the fast application of pharmacological compounds, such as a variety of ionic or pH indicators, to the intra- and extracellular media. With an exceptionally high signal-to-noise ratio, this method enables the measurement of dark spontaneous and light-induced unitary currents (i.e.&#160;spontaneous and quantum bumps) and macroscopic Light-induced Currents (LIC) from single D. melanogaster photoreceptors. This protocol outlines, in great detail, all the key steps necessary to perform this technique, which includes both electrophysiological and optical recordings. The fly retina dissection procedure for the attainment of intact and viable ex vivo isolated ommatidia in the bath chamber is described. The equipment needed to perform whole-cell and fluorescence imaging measurements are also detailed. Finally, the pitfalls in using this delicate preparation during extended experiments are explained.

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell recordings from Drosophila melanogaster photoreceptors enable the measurement of spontaneous dark bumps, quantum bumps, macroscopic responses to light, and current-voltage relationships under various conditions. In combination with D. melanogaster genetic manipulation tools, this method enables the study of the ubiquitous inositol-lipid signaling pathway and its target, the TRP channel.

Méthode électrophysiologique pour les enregistrements de la pince à tension intégrale à partir de<em&gt; Drosophila</em&gt; Photorécepteurs

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, ...

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, ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Ex Vivo Imagerie calcique pour visualiser le cerveau réponses à signalisation endocrinienne chez la drosophile

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

Préparation du drosophile adulte melanogaster pour l’imagerie cérébrale entière pendant les réponses au comportement et aux stimuli

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

In vivo Calcium Imaging of Mouse Geniculate Ganglion Neuron Responses to Taste Stimuli

Within the last ten years, advances in genetically encoded calcium indicators (GECIs) have promoted a revolution in in vivo functional imaging. Using calcium as a proxy for neuronal activity, these techniques provide a way to monitor the responses of individual cells within large neuronal ensembles to a variety of stimuli in real time. We, and others, have applied these techniques to image the responses of individual geniculate ganglion neurons to taste stimuli applied to the tongues of live anesthetized mice. The geniculate ganglion is comprised of the cell bodies of gustatory neurons innervating the anterior tongue and palate as well as some somatosensory neurons innervating the pinna of the ear. Imaging the taste-evoked responses of individual geniculate ganglion neurons with GCaMP has provided important information about the tuning profiles of these neurons in wild-type mice as well as a way to detect peripheral taste miswiring phenotypes in genetically manipulated mice. Here we demonstrate the surgical procedure to expose the geniculate ganglion, GCaMP fluorescence image acquisition, initial steps for data analysis, and troubleshooting. This technique can be used with transgenically encoded GCaMP, or with AAV-mediated GCaMP expression, and can be modified to image particular genetic subsets of interest (i.e., Cre-mediated GCaMP expression). Overall, in vivo calcium imaging of geniculate ganglion neurons is a powerful technique for monitoring the activity of peripheral gustatory neurons and provides complementary information to more traditional whole-nerve chorda tympani recordings or taste behavior assays.

Here we present how to expose the geniculate ganglion of a live, anesthetized laboratory mouse and how to use calcium imaging to measure the responses of ensembles of these neurons to taste stimuli, allowing for multiple trials with different stimulants. This allows for in depth comparisons of which neurons respond to which tastants.

Imagerie calcique in vivo des réponses des neurones ganglionnaires géniculés de souris aux stimuli gustatifs

Within the last ten years, advances in genetically encoded calcium indicators (GECIs) have promoted a revolution in in vivo functional imaging. Using ...