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 mosca de la fruta del ojo (Drosophila melanogaster) compuesto es un buen sistema modelo para investigar la organización funcional de los fotorreceptores y las interneuronas matrices para el muestreo y procesamiento de imagen neuronal, y para la visión de los animales. El sistema cuenta con el diagrama de cableado más completa 1,2 y es amigable para las manipulaciones genéticas y supervisión de la actividad neuronal precisa (de alta relación señal-ruido y el tiempo de resolución) 3-10. El ojo de Drosophila es modular, que contiene ~ 750 estructuras de lentes con tapón aparentemente regulares llamados ommatidia, que en conjunto proporcionan un campo de visión de la panorámica que cubre casi todas las direcciones alrededor de su cabeza volar. Información primaria del ojo unidades de muestreo son sus fotorreceptores rhabdomeric 7,8,11. Cada ommatidium contiene ocho células fotorreceptoras (R1-R8), que comparten la misma lente faceta pero están alineados con siete direcciones diferentes. Mientras que los fotorreceptores exteriores R1-R6 are más sensible a la luz azul-verde, sensibilidades espectrales de las células internas R7 y R8, que están por encima de la otra y apuntan a la misma dirección, exhiben tres subtipos distintivos: pálido, amarillo y área del borde dorsal (DRA) 12- 15. <img alt="Figura 1" src="/files/ftp_upload/54142/54142fig1.jpg" /> Figura 1. Organización funcional del ojo de Drosophila. (A) Los dos primeros ganglios de la óptica, la retina y la lámina, se resaltan en gris en el interior del ojo de la mosca. Fotorreceptores de la retina R1-R6 y lámina grandes células monopolares (LMC: L1-L3) son de fácil acceso en vivo a las grabaciones de microelectrodos convencionales fuerte. El electrodo esquemática destaca la ruta normal a grabar de R1-R6 en la retina. Uno de los caminos para grabar desde LMC en la lámina es cambiar de forma paralela al electrodo de la izquierda. (B) Lamina es una matriz de órgano retinotopicallyzados cartuchos, cada uno de los cuales está lleno de neuronas que procesa la información a partir de una pequeña área específica en el espacio visual. Debido a la superposición neural, seis fotorreceptores de diferentes ommatidia vecinos envían sus axones (R1-R6) a la misma cartucho de lámina, formando sinapsis de salida histaminérgicos a L1-L3 y una célula amacrine (Am). (C) La difusión de la información neural entre terminales de los axones R1-R6 y las interneuronas visuales (incluyendo L4, L5, Lawf, C2, C3 y T1), en el interior de un cartucho de lámina es complejo. (D) R1-R6 fotorreceptor axones reciben evaluaciones sinápticas de las células monopolar L2 y L4. (B) y (C) modificado de Rivera-alba et al 2. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig1large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> El ojo de Drosophila es del tipo neuronal superposición 16. Esto significa tsombrero de las señales neurales de ocho fotorreceptores pertenecientes a siete ommatidia vecina, que se ven en el mismo punto en el espacio, se agrupan juntos en un cartucho neural en los próximos dos neuropils: la lámina y la médula. Mientras que los seis fotorreceptores exteriores del proyecto R1-R6 sus terminales de los axones neuronales en columnas a la lámina (Figura 1), las células R7 y R8 pasar por alto esta capa y hacen contactos sinápticos con su columna de médula 17-19 correspondiente. Estos cableados exactas producen el sustrato neural para el mapeo retinotopic de la mosca de la visión temprano, después de lo cual cada lámina (Figuras 1A-C) y la columna médula (cartucho) representa un único punto en el espacio. Las entradas directas de fotorreceptores R1-R6 son recibidas por las células monopolares grandes (LMC: L1, L2 y L3) y la célula amacrinas (AM) en la lámina 1,2,20. De estos, L1 y L2 son las células más grandes, que median en las principales vías de información (Figura 1D), WHIch responden a dentro y fuera de los bordes en movimiento, y por lo tanto constituyen la base de cálculo del detector de movimiento 21,22. Experimentos sobre el comportamiento sugiere que, en contraste intermedia, las dos vías facilitan la percepción del movimiento de direcciones opuestas: back-to-front en L1 y de adelante hacia atrás en células L2 23,24. Conectividad implica, además, que las neuronas L4 pueden jugar un papel crítico en la comunicación lateral entre los cartuchos vecinos 25,26. sinapsis recíprocas se encontraron entre las células L2 y L4, situados en la misma y dos cartuchos adyacentes. Aguas abajo, cada célula L2 y sus tres células L4 asociados proyectan sus axones a un objetivo común, la neurona Tm2 en la médula, donde se cree que las aportaciones de los cartuchos de vecinos para ser integrado para el procesamiento de movimiento de delante a atrás 27. Aunque las neuronas L1 reciben el aporte de personas del mismo cartucho de L2 a través de las dos uniones comunicantes y las sinapsis, que no están conectados directamente a los cartuchos de lámina L4s y por lo tanto adyacentes. <pclass = &quot;jove_content&quot;&gt; evaluaciones Synaptic para fotorreceptor axones R1-R6 son proporcionados únicamente por las neuronas pertenecientes a los circuitos / L4 L2 pero no la vía de 1,2 L1 (Figura 1D). Mientras que las conexiones del mismo cartucho son selectivamente de L2 a R1 y R2 y de L4 a R5, todos los fotorreceptores R1-R6 reciben retroalimentación sináptica de L4 de uno o ambos cartuchos de vecinos. Además, existen fuertes conexiones sinápticas de AM a R1, R2, R4 y R5, y células gliales también están conectados sinápticamente a la red y por lo tanto pueden participar en el procesamiento de imágenes neural 6. Por último, axonal gap-uniones, que une la vecina R1-R6 y R6 y R7 entre / R8 fotorreceptores en la lámina, contribuyen a la representación de la información asimétrica y el procesamiento de cada cartucho 14,20,28. Grabaciones de tensión intracelulares de fotorreceptores individuales y las interneuronas visuales en casi intacto Drosophila proporcionan una alta r-señal-ruidoación de datos con una resolución sub-milisegundo 3,5,7-10,29, que es necesaria para dar sentido a los cálculos neuronales rápidas entre las neuronas conectadas. Este nivel de precisión no es posible mediante técnicas de imagen ópticos actuales, que son significativamente más ruidoso y por lo general operan a las 10 - una resolución de 100 ms. Por otra parte, debido a que los electrodos tienen puntas muy pequeñas y afiladas, el método no está restringido a los cuerpos de las células, pero puede proporcionar grabaciones directas de las estructuras nerviosas activas pequeñas; tales como árboles dendríticas los LMC &#39;o axones fotorreceptoras, las cuales no se puede acceder por consejos mucho más grandes de los electrodos de patch-clamp. Es importante destacar que el método también es estructuralmente menos invasivo y perjudicial que la mayoría de las aplicaciones de patch-clamp, y así afecta a menos de muestreo medio y la información intracelular de las células estudiadas. Por lo tanto, las técnicas de microelectrodos afilados convencionales han contribuido, y siguen contribuyendo, descubrimientos fundamentales y la visión original en Infor neuronalprocesamiento de la escala en el momento adecuado; mejorar nuestra comprensión mecanicista de la visión 3-10. En este artículo se explica cómo en registros intracelulares in vivo de Drosophila fotorreceptores y LMC R1-R6 se llevan a cabo en el laboratorio Juusola. Este protocolo describe cómo construir una plataforma de electrofisiología adecuado, preparar la marcha, y realizar las grabaciones. Se presenta algunos datos representativos, y algunos problemas comunes y las posibles soluciones se discuten que se pueden encontrar cuando se utiliza este método.

<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.

El método de grabación de microelectrodos agudo, tal como fue adaptada aquí por el ojo de Drosophila, se puede utilizar para cuantificar de manera fiable el muestreo y el procesamiento neuronal de información en las células de la retina y la lámina, y la comunicación entre ellos 4,5,7,8,10,33. Al usarlo para estudiar la codificación en diferentes poblaciones de tipo salvaje, mutantes o cepas de moscas de la ingeniería genética, el método ha demostrado su valor; no sólo en la cuantificación de los efectos de una mutación, la temperatura, la dieta o de expresión seleccionado 3,4,6,9,10,14,30,34, sino también en la revelación de razones mecanicistas para alterados comportamientos visuales 14,34. El método también es fácilmente aplicable a otras preparaciones de insectos 35,36, potenciando los estudios de visión neuroethological. A continuación, mostramos algunos ejemplos de sus aplicaciones exitosas. <img alt="Figura 7" src="/files/ftp_upload/54142/54142fig7.jpg" /> Figura 7. VoltajeRespuestas de una mosca de la fruta R1-R6 fotorreceptor a un pulso de luz a los 20 y 25 ° C. Debido a las penetraciones de microelectrodos afilados son a menudo muy estable, es posible registrar las respuestas de tensión de la misma fotorreceptor R1-R6 a un estímulo de luz dada en diferentes temperaturas ambiente por el calentamiento o enfriamiento de la mosca. En nuestros montajes, la mosca titular se coloca en un sistema de control de temperatura de cerca de bucle basado elemento Peltier. Esto nos permite cambiar la temperatura de la cabeza de la marcha en cuestión de segundos. Una temperatura más alta acelera las respuestas de tensión y de forma característica reduce el potencial de reposo de los fotorreceptores R1-R6 (como se indica por las flechas rojas). <a href="https://www.jove.com/files/ftp_upload/54142/54142fig7large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> Estudiar el efecto de la temperatura sobre la salida de fotorreceptores <p class="jove_content" fo:kEEP-together.within-page = &quot;1&quot;&gt; Con un buen diseño y un sistema de grabación de vibración aislada, el método puede ser usado para medir el efecto de la temperatura en la salida de los nervios de una célula individual por el calentamiento o enfriamiento de la mosca. El ejemplo que muestra las respuestas de voltaje a un brillante pulso de 10 ms de largo, registrados en el mismo fotorreceptor R1-R6 a los 20 y 25 ° C (Figura 7). Cuantificada antes de 4,9, el calentamiento reduce el potencial de reposo de un fotorreceptor en la oscuridad, y acelera sus respuestas tensión. <img alt="Figura 8" src="/files/ftp_upload/54142/54142fig8.jpg" /> Figura 8. Rendimiento de Señalización de una mosca de la fruta R1-R6 fotorreceptor mejora con la intensidad de la luz (A) de salida de fotorreceptores se atenúen (abajo) y brillante. (Encima; 10.000 veces una luz más brillante) repiten series temporales intensidad de la luz naturalista registrado por la misma microelectrodoen la misma célula a 20 o C. Las respuestas al estímulo luminoso son más grandes, ya que integran más muestras, respuestas elementales (protuberancias) a fotones individuales 4,5,7,8. (B) 20 respuestas de voltaje consecutivos de un segundo de larga se superponen. Se tomaron las respuestas individuales (gris claro) después de que las tendencias adaptativas (flecha en A) había retrocedido (recuadro de puntos en A). Los medios de respuesta correspondientes (las señales) son las huellas más oscuros. La diferencia entre la señal y las respuestas individuales es el ruido. (C) Las células de rendimiento de señalización se cuantificó por las grabaciones de &#39;relaciones de señal a ruido (SNR) mediante los métodos estándar 4,5,7,8. Salida de los fotorreceptores tiene alrededor de 64 Hz más amplia gama de señalización fiable en la estimulación luminosa ( &#39;≥1 brillante, hasta 84 Hz que en el dim (SNR SNR)&#39; Dim ≥1, hasta 20 Hz), con signal-a-ruido mejora en gran medida; de SNR Dim MAX = 87 a SNR brillante MAX = 1,868. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig8large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> El estudio de adaptación y codificación neuronal repetitiva por estimulación La invasividad del método, haciendo relativamente poco daño en las estructuras de la retina y la lámina, lo hace ideal para el estudio del rendimiento de señalización de las células individuales a diferentes estímulos de luz en su estado fisiológico natural cerca in vivo. La figura 8 muestra las respuestas de tensión de un R1- R6 fotorreceptor a un estímulo serie oscuro y brillante repetida naturalista tiempo intensidad de la luz, a 20 ° C, mientras que la figura 9 </strong&gt; muestra las respuestas de otro fotorreceptor R1-R6 y un LMC a un estímulo naturalista diferente a 25 ° C. Las grabaciones pre y postsinápticos se realizaron por separado a partir de dos diferentes moscas porque registros intracelulares simultáneos de dos microelectrodos afilados de la misma mosca, una en la retina y el otro en la lámina, son demasiado difíciles de ser viable 30. <img alt="Figura 9" src="/files/ftp_upload/54142/54142fig9.jpg" /> Figura 9. Respuestas de tensión de una mosca de la fruta R1-R6 fotorreceptores y LMC a las repetidas naturalista estimulación a 25 ° C (A) R1-R6 (gris) y LMC (negro) salidas registradas por diferentes microelectrodos de diferentes moscas. (B) 20 pre-luz totalmente adaptada consecutiva (arriba) y las respuestas postsinápticas (a continuación) para el mismo patrón de estímulo naturalista con las respuestas individuales, se muestra en gris claro unad los correspondientes medios de respuesta (las señales) como las huellas oscuras. La diferencia entre la señal y las respuestas individuales es el ruido. (C) Las células de rendimiento de señalización se cuantificó por las grabaciones de &#39;relaciones de señal a ruido (SNR). LMC tiene salida de aproximadamente 10 Hz más amplia gama de señalización fiable (SNR &#39;LMC ≥1, hasta 104 Hz) de salida R1-R6 (SNR&#39; R ≥1, hasta 94 Hz). Ambas relaciones señal-ruido son altos (SNR LMC MAX = 142, SNR R MAX = 752), y como el ruido de la grabación fue baja, sus diferencias reflejan diferencias de codificación reales entre las células. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig9large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> Después del inicio del estímulo, las grabaciones típicaLy muestre la adaptación rápida tendencias que se desaparecen en gran parte dentro de 5-6 seg. A partir de entonces, las células producen respuestas muy consistentes a cada 1 seg presentación larga de estímulo (cada recuadro de puntos encierra 20 de estos). La repetibilidad de las respuestas se hace evidente cuando estos se superponen (Figura 8B y en la Figura 9B). Las respuestas individuales son las huellas grises delgadas, y su significan el grueso trazo más oscuro. La respuesta media se toma como la señal neural, mientras que el ruido neural es la diferencia entre la media y cada 4,5,9,37,38 respuesta individual. Las respectivas relaciones de señal a ruido en el dominio de frecuencia (Figura 8C y la Figura 9C) se obtuvieron por Fourier-transformación de los fragmentos de datos de señal y ruido en espectros de potencia, y dividiendo el espectro de potencia media de la señal con el correspondiente espectro de potencia de ruido media 4, 5,9,37,38. Característicamente, las máximas relaciones señal a ruido of las salidas neuronales registrados a la estimulación naturalista son altas (100 - 1000), y en las preparaciones más estables con ruido de grabación muy baja puede alcanzar valores &gt;&gt; 1000 (por ejemplo, la Figura 8C.). Nótese también que el calentamiento se expande el ancho de banda de las células de señalización fiable 4 (SNR &#39;brillante ≥ 1); por ejemplo, la diferencia relativa entre los dos R1-R6S en las figuras 8 y 9, respectivamente, es de 10 Hz (84 a 20 ° C y 94 Hz a 25 o C). Uno puede estimar más velocidad de transferencia de información a partir de su relación señal-ruido de cada célula mediante el uso de la ecuación de Shannon 32, o mediante el cálculo de la diferencia entre la entropía y la entropía de ruido tasas de las respuestas &#39;a través del método de extrapolación de triple 39. Más detalles acerca de los análisis teóricos de la información, así como su uso y limitacioness específicamente con este método se dan en las publicaciones anteriores 7,8,39. <img alt="Figura 10" src="/files/ftp_upload/54142/54142fig10.jpg" /> Figura 10. Respuestas de tensión de una mosca del asesino R1-R6 fotorreceptores y LMC a las repetidas naturalista estimulación a 19 ° C (A) R1-R6 (gris) y LMC (negro) resultados registrados por el mismo microelectrodo de la misma mosca; postsináptico primera y más tarde presináptica, como el electrodo se avanzó en el ojo. (B) 20 pre consecutiva (arriba) y postsinápticos (abajo) respuesta (trazas de color gris claro) para el mismo patrón de estímulo naturalista fueron capturados después de la adaptación inicial (recuadro de puntos en A). Sus medios son las señales (los trazos más oscuros en la parte superior), mientras que sus respectivas diferencias con respecto a las respuestas individuales dan el ruido. (C) El correspondiente SignaRatios-l-ruido (SNR) se calcularon como en las figuras 8C y 9C. LMC tiene salida sobre un rango más amplio de 100 Hz de señalización fiable (SNR &#39;LMC MAX ≥1, hasta 234 Hz) de salida R1-R6 (SNR&#39; R MAX ≥1, hasta 134 Hz). Ambas relaciones señal-ruido son altos (SNR LMC MAX = 137, SNR R MAX = 627), y como la misma microelectrodo se utilizó en las grabaciones, sus diferencias reflejan diferencias reales en las salidas neurales pre- y postsinápticos. Estos resultados implican que el sistema de grabación tenía bajo nivel de ruido, y su influencia en los análisis fue marginal. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig10large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> Estudios Vision Neuroethological </p&gt; El método también se puede utilizar para registrar las respuestas pre y postsinápticos de tensión de los ojos compuestos de diferentes especies de insectos 7,8,35,36 (Figura 10), lo que permite estudios comparativos neuroethological de procesamiento de la información visual. Para el sistema de grabación presentada, la única adaptación requerida es de preparación de los nuevos titulares, cada uno con una abertura de dimensiones adecuadas para las especies estudiadas. Estas grabaciones son ejemplares de un asesino de mosca hembra (Coenosia attenuata). Ellos muestran respuestas de tensión intracelulares de un fotorreceptor R1-R6 y LMC a la estimulación repetitiva luz idéntica, tal como se utiliza para los homólogos de Drosophila en la Figura 9, pero a 19 o C. En este caso, tanto los datos pre y postsinápticos se registraron de la misma mosca; uno después del otro, con el mismo electrodo de registro (lleno de 3 M KCl) primero avanzando a través de la lámina lateral antes de entraring la retina frontal. En comparación con los datos de Drosophila a 25 ° C, los datos Coenosia - incluso a la temperatura más fría - muestra las respuestas con dinámicas más rápidas; ampliar la gama de señalización fiable (relación señal a ruido &gt;&gt; 1) sobre un rango de frecuencias más amplio. Tales adaptaciones funcionales en la codificación neuronal de estímulos naturalistas están en consonancia con el estilo de vida depredador Coenosia &#39;s 36, que requieren alta precisión la información espacio-temporal para lograr comportamientos de caza aéreas rápidas.

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.

Método electrofisiológico para registrar las respuestas de voltaje intracelular<em&gt; Drosophila</em&gt; Los fotorreceptores y interneuronas a la luz Estímulos<em&gt; En 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 vistas.  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.

Video: 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.

Dos registros electrofisiológicos de sinápticamente respuestas evocadas neuronal y astroglial en agudas rodajas de hipocampo

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.

Un método para Sistemática Electroquímica y electrofisiológico Evaluación de Neuronales Grabación Electrodos

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.

Electrofisiológico Grabación desde<em&gt; Drosophila</em&gt; Labellar Taste Sensilla

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.

Una guía para<em&gt; En vivo</em&gt; Single-unidad de grabación de Optogenetically Identificado cortical inhibitorio interneuronas

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.

Analizando Synaptic Modulación de<em&gt; Drosophila melanogaster</em&gt; Los fotorreceptores después de la exposición prolongada a la luz

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.

Método electrofisiológico para grabaciones de abrazadera de voltaje de células enteras<em&gt; Drosophila</em&gt; Fotorreceptores

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.

Registro electrofisiológico de<em&gt; Drosophila</em&gt; Tricoide Sensilla en Respuesta a Odorantes de Baja Volatilidad

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 Proyección de imagen de calcio para la visualización de las respuestas cerebrales a la señalización endocrina en Drosophila

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.

Preparación de Drosophila melanogaster adulto para imágenes cerebrales completas durante las respuestas de comportamiento y estímulos

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.

Imágenes de calcio in vivo de las respuestas de las neuronas ganglionares geniculadas de ratón a los estímulos del gusto

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