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.
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 mouche des fruits (Drosophila melanogaster) composé oeil est un système modèle grand pour enquêter sur l'organisation fonctionnelle des photorécepteurs et interneurones tableaux pour l' 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' activité neuronale (de haut rapport signal-bruit et résolution temporelle) 3-10.
L'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' information primaire de l'œil unités d' 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'autre et pointer vers la même direction, présentent trois sous – types distincts: pâle, zone de bord jaune et dorsale (DRA) 12- 15.
Figure 1. Organisation fonctionnelle du Drosophila Eye. (A) Les deux premiers noyaux optiques, rétine et lamina, sont mis en évidence en gris dans l'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'électrode schématique montre le trajet normal à enregistrer à partir de R1 à R6 dans la rétine. Un chemin d'accès à enregistrer à partir de LMC dans le limbe est de déplacer en parallèle l'électrode à gauche. (B) Lamina est une matrice d'organe rétinotopiquecartouches lisés, dont chacun est emballé avec les neurones qui traite des informations à partir d'une petite zone spécifique dans l'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'une cellule amacrine (Am). (C) La diffusion de l' information neuronale entre les terminaisons axonales R1-R6 et les interneurones visuels (y compris L4, L5, Lawf, C2, C3 et T1), à l' intérieur d' 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. S'il vous plaît cliquer ici pour voir une version plus grande de cette figure.
L'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'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' 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'information (figure 1D), which répondent à l' intérieur et à bords Off-déplacement, et forment ainsi la base de calcul du détecteur de mouvement 21,22. Expériences comportementales suggèrent qu'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'être intégré pour le traitement de l' avant vers l' 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 = "jove_content"> 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'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' 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' asymétrie d' 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'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'information d'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'infor- à l'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'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'utilisation de cette méthode.
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.
The authors have nothing to disclose.
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).
Stereo Zoom Microscope for making the fly preparation | Olympus | SZX12 DFPLFL1.6x PF eyepieces: WHN30x-H/22 | Capable of ~150X magnification with long working distance; bespoke heavy steel table mount stand |
Stereomicroscope in the intracellular set-up | · Olympus | Olympus SZX7; eyepieces: WHN30x-H/22 | 30x eyepieces are needed for seeing the electrode tip reflections well when driving it through the small corneal hole into the eye |
· Nikon | Nikon SMZ645; eyepieces: C-W30x/7 | ||
Anti-vibration Table | · Melles Griot | With metric M6 holes on the breadboard | 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 |
· Newport | |||
Micromanipulators | · Narishige | · Narishige NMN-21 | 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. |
· Huxley Bertram | · Huxley xyz-axis with fine manual control | ||
· Sensapex | · Sensapex triple axis | ||
· Märzhäuser | · Märzhäuser DC-3K with additional x-axis piezo stepper and MS 314 controller | ||
Magnetic Stands | Any magnetic base with on/off switch will do | For example, to manage cables inside the Faraday cage | |
Electrode Holders | Harvard Apparatus | ESP/W-F10N | |
Silver Wire | World Precision Instruments | AGW1510 | 0.3-0.5 mm diameter; needs to be chloridized for the electrode holders |
Fiber Optic Light Source | Many different, including Olympus | ||
Fiber Optic Bundles | · UltraFine Technology | To deliver the LED light stimulus to the Cardan arm system. We use both liquid and quartz light guides (range from UV to IR) | |
· Thorn Labs | |||
Fly Cathing Tube | P80-50P 50ml Cent. Tube PP., Pack of 100 Pcs | Cut the conical bottom off from 50 ml Plastic Centrifuge Tube and glue a 1 ml pipette tip on it. | |
Digital Acquisition System | National Instruments | ||
Single-electrode current/voltage-clamp microelectrode amplifier | npi SEC-10LX | http://www.npielectronic.de/products/amplifiers/sec-single-electrode-clamp/sec-10lx.html | Outstanding performer! |
Head-stage | Standard (+/- 150 nA) | For npi SEC-10LX | |
LED light sources and drivers | · 2-channel OptoLED (Cairn Research Ltd., UK) | Many of our stimulus systems are in-house built | |
· Self-designed and constructed | |||
Acquisition and Analyses Software | Many companies to choose from | Biosyst; custom written Matlab-based system for experimental and theoretical work in the Juusola laboratory | |
Personal Computer or Mac | Ensure that PC or Mac is compatible with data acquisition system and software | ||
Cardan arm system | Self-designed and constructed | Providing accurate x,y,z-positioning of the light stimuli | |
Peltier temperature control system | Self-designed and constructed | ||
Faraday Cage | Self-constructed | Electromagnetic noise shielding | |
Filamented Borosilicate Glass Capillaries | Outer diameter: 1 mm | ||
Inner diameter: 0.5-0.7 mm | |||
Filamented Quartz Glass Capillaries | Outer diameter: 1 mm | ||
Inner diameter: 0.5-0.7 mm | |||
Pipette Puller | Sutter Instrument Company | Model P-2000 laser Flaming/Brown Micropipette Puller | 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 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’ preset parameters serve as useful starting points for systematic modifications to generate electrodes with good penetration success and low recording noise. |
Extracellular Ringer Solution for the reference electrode | Chemicals from Fisher Scientific | 10326390, NaCl 10010310, KCl 10147753, TES 10161800, CaCl2 10159872, MgCl2 10000430, sucrose | See the recipe in the protocol section |
3 M KCl solution for filling the filamented recording microelectrode | Salts from Fisher Scientific | 10010310, KCl | |
Petroleum jelly | Vaselin | ||
Non-stainless steel razor blades | |||
Blade holder/breaker | Fine Science Tools By Dumont | 10053-09 | 9 cm |
Blu-tack | Bostik | Alternatively, use molding clay | |
Forceps | Fine Science Tools By Dumont | 11252-00 | #5SF (super-fine tips) |