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.
Die Fruchtfliege (Drosophila melanogaster) -Verbindung Auge ist ein großes Modellsystem die funktionelle Organisation des Photorezeptors und Euron – Arrays für die neuronale Bild Probenahme und Verarbeitung sowie für die Tier Vision zu untersuchen. Das System verfügt über das umfassendste Schaltplan 1,2 und ist liebenswürdig zu genetischen Manipulationen und präzise neuronale Aktivitätsüberwachung (von hohen Signal-zu-Rausch – Verhältnis und Zeitauflösung) 10.03.
Das Drosophila Auge ist modular aufgebaut, mit ~ 750 scheinbar normalen Linse bedeckten Strukturen Ommatidien genannt, die zusammen das eine Panorama – Sichtfeld fliegen sorgen , dass fast jede Richtung um seinen Kopf bedeckt. Das Auge des primären Informationsstichprobeneinheiten sind seine rhabdomer Photorezeptoren 7,8,11. Jede Ommatidium enthält acht Sehzellen (R1-R8), welche die gleiche Facettenlinse teilen, sind jedoch sieben verschiedene Richtungen ausgerichtet sind. Während die äußeren Photorezeptoren R1-R6 are am empfindlichsten auf blau-grünes Licht, spektralen Empfindlichkeiten der inneren Zellen R7 und R8, die in die gleiche Richtung liegen übereinander und zeigen, zeigen drei verschiedene Subtypen: blass, gelb und dorsalen Randbereich (DRA) 12- 15.
Abbildung 1. Funktionelle Organisation des Drosophila Auge. (A) Die beiden ersten Opticusganglien, Retina und Lamina, sind grau im Inneren des Fliegenaugen hervorgehoben. Retina R1-R6 und Photorezeptoren Lamina Große monopolare Zellen (LMCs: L1-L3) sind leicht zugänglich in vivo zu herkömmlichen scharfen Mikroelektroden – Aufnahmen. Die schematische Elektrode hebt den normalen Weg von R1-R6 in der Netzhaut aufzuzeichnen. Ein Weg aus LMCs in der Lamina zu erfassen ist parallel der Elektrode zu verschieben nach links. (B) Lamina ist eine Matrix von retinotop organized Patronen, von denen jede mit Neuronen gepackt, die Informationen von einem bestimmten kleinen Bereich in dem Bildraum verarbeitet. Aufgrund neuralen Überlagerung von sechs Photorezeptoren aus verschiedenen benachbarten Ommatidien senden ihre Axone (R1-R6) auf den gleichen Plättchen Patrone histaminerge Ausgangs Synapsen L1-L3 und einer amacrine Zelle (Am) bilden. (C) Die Ausbreitung der neuronalen Informationen zwischen R1-R6 Axonterminalen und die visuellen Inter (einschließlich L4, L5, Lawf, C2, C3 und T1), innerhalb einer Lamina Patrone komplex ist. (D) R1-R6 Photorezeptor Axone erhalten synaptischen Feedbacks von L2 und L4 monopolare Zellen. (B) und (C) modifiziert von Rivera-alba et al 2. Bitte hier klicken , um eine größere Version dieser Figur zu sehen.
Das Drosophila Auge des neuronalen Überlagerung Typ 16. Dies bedeutet, tHut die neuronalen Signale von acht Photorezeptoren zu sieben gehör Ommatidien Nachbar, die im Raum an der gleichen Stelle suchen, werden zusammen in den nächsten zwei Neuropile an einer neuralen Patrone gepoolt: die Lamina und Medulla. Während die sechs äußeren Photorezeptoren R1-R6 Projekt ihre Axonterminalen zu neuronalen Spalten in der Lamina (Abbildung 1), R7 und R8 Zellen umgehen diese Schicht und synaptische Kontakte mit ihren Medulla Spalte 17-19 entspricht. Diese exakten Verdrahtungen erzeugen , das neuronale Substrat für die retinotopic Abbildung von Fly frühe Vision, wonach jede Lamelle (1A-C) und der Medulla Säule (Patrone) stellt einen einzigen Punkt im Raum.
Direkte Eingänge von R1-R6 Photorezeptoren sind durch die großen monopolare Zellen erhalten (LMCs: L1, L2 und L3) und die Amakrinzell (Am) in der Lamina 1,2,20. Von diesen L1 und L2 sind die größten Zellen, zu vermitteln wichtige Informationswege (1D), which reagieren auf On- und Off-bewegten Kanten und bilden somit die Berechnungsgrundlage des Bewegungsmelders 21,22. Verhaltensexperimente legen nahe , dass bei mittleren dagegen die beiden Bahnen Bewegungswahrnehmung von entgegengesetzten Richtungen zu erleichtern: Back-to-Front in L1 und Front-to-Back in L2 – Zellen 23,24. Konnektivität bedeutet ferner , dass L4 – Neuronen entscheidende Rolle zwischen benachbarten Patronen 25,26 in der seitlichen Kommunikation spielen kann. Gegenseitige Synapsen wurden zwischen L2 und L4-Zellen befindet sich in der gleichen und zwei benachbarte Patronen gefunden. Downstream jede L2 – Zelle und die drei zugeordneten L4 Zellen ihre Axone mit einem gemeinsamen Ziel – Projekt, das Tm2 Neuron in der Medulla, wobei Eingänge von benachbarten Patronen geglaubt werden , integriert werden zur Verarbeitung von vorne nach hinten Bewegung 27. Obwohl L1 Neuronen über beide gap junctions und Synapsen Eingang von gleich Patrone L2s erhalten, sind sie nicht direkt mit L4s und damit benachbarte Lamina Cartridges.
<pclass = "jove_content"> Synaptische Feedbacks zu R1-R6 Photorezeptor Axone werden nur von Neuronen, die zu den L2 / L4 – Schaltungen aber nicht die L1 – Weg 1,2 (1D) zur Verfügung gestellt. Während gleichen Patronen Verbindungen selektiv von L2 auf R1 und R2 und von L4 bis R5 alle R1-R6 Photorezeptoren erhalten synaptische Rückkopplung von L4 von einem oder beiden benachbarten Patronen. Darüber hinaus gibt es starke synaptische Verbindungen von Am zu R1, R2, R4 und R5 und Glia – Zellen werden auch synaptisch mit dem Netzwerk verbunden und kann somit in neuronalen Bildverarbeitung 6 teilnehmen. Schließlich tragen axonalen spalt Kreuzungen verbindet benachbarte R1-R6 und zwischen R6 und R7 / R8 Photorezeptoren in der Lamina, die asymmetrische Informationsdarstellung und -verarbeitung in jeder Patrone 14,20,28.Intrazellulärer Spannung Aufnahmen von einzelnen Photorezeptoren und visuelle Inter in nahezu intakten Drosophila bieten hohen Signal-zu-Rausch – ratio Daten an Sub-Millisekunden – Auflösung 3,5,7-10,29, die zur Herstellung von Sinn der schnellen neuronalen Berechnungen zwischen den verbundenen Neuronen notwendig ist. Dieser Grad an Genauigkeit ist durch aktuelle optische Abbildungstechniken unmöglich, die wesentlich geräuschvoller sind und typischerweise bei 10 arbeiten – 100 msec Auflösung. Darüber hinaus, da die Elektroden sehr klein und scharfe Spitzen haben, ist das Verfahren nicht auf Zellkörper beschränkt, sondern direkte Aufnahmen von kleinen aktiven neuronalen Strukturen zur Verfügung stellen kann; wie die dendritischen Bäume oder Photorezeptor Axone 'LMCs, die von viel größeren Spitzen Patch-Clamp-Elektroden nicht zugegriffen werden kann. Wichtig ist, dass das Verfahren auch strukturell weniger invasive und schädlich als die meisten Patch-Clamp-Anwendungen, und wirkt sich so weniger die intrazelluläre Milieu und Informationen Sampling 'studierte Zellen. So haben herkömmliche scharfe Mikroelektroden-Techniken beigetragen, und halten Sie auf einen Beitrag, grundlegenden Entdeckungen und originelle Einblicke in neuronale information Verarbeitung bei der entsprechenden Zeitskala; unser mechanistisches Verständnis der Vision Verbesserung 10.03.
Dieser Artikel beschreibt , wie in vivo intrazelluläre Aufnahmen von Drosophila R1-R6 und Photorezeptoren LMCs im Juusola Labor durchgeführt werden. Dieses Protokoll wird beschrieben, wie ein geeignetes Elektro rig zu konstruieren, bereiten die Fliege, und die Aufnahmen durchführen. Einige repräsentative Daten vorgestellt, und einige häufig auftretende Probleme und mögliche Lösungen diskutiert, die auftreten können, wenn mit dieser Methode.
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) |