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.
초파리 (Drosophila의 melanogaster의) 화합물의 눈은 신경 이미지 샘플링 및 처리를위한 광 수용체 및 interneuron 배열의 기능적 조직을 조사 할 수있는 좋은 모델 시스템이며, 동물 비전. 시스템은 가장 완벽한 배선도 1,2-을 가지며 유전자 조작 및 3-10 (높은 신호 – 대 – 잡음 비율 및 시간 해상도) 정확한 신경 활동 모니터링에 호감이다.
초파리의 눈은 함께가 머리 주위 거의 모든 방향을 커버하는 탁 트인 시야 비행을 제공 ommatidia, 개안라고 ~ 750 겉으로는 일반 렌즈 덮인 구조를 포함하는 모듈이다. 단위는 rhabdomeric 광 수용체 7,8,11 있습니다 샘플링 눈의 주요 정보를 제공합니다. 각 개안은 같은면 렌즈를 공유하지만, 일곱 가지 방향으로 정렬됩니다 팔 광 수용체 세포 (R1-R8)를 포함한다. 외부 광 수용체 R1-R6 아칸소 동안전자 파란색 – 녹색 빛에 가장 민감, 같은 방향으로 서로 지점의 상단에 거짓말 내부 세포 R7과 R8의 스펙트럼 감도, 전시 세 가지 독특한 하위 유형이 : 창백한 노란색과 지느러미 가장자리 영역 (DRA) 12 15.
그림 1. 초파리 눈의 기능 조직. (A) 두 최초의 광학 신경, 망막과 얇은 판은 플라이 눈 안쪽 회색으로 강조 표시됩니다. 망막 R1-R6의 광 수용체와 얇은 대형 모노 폴라 셀 (중저 소득 : L1-L3)는 기존의 날카로운 미세 전극 녹음에 생체 내에서 쉽게 액세스 할 수 있습니다. 개략적 전극 망막 R1-R6에서 기록하는 정상 경로를 강조한다. 라미에 중저 소득 국가에서 기록하는 하나의 경로는 왼쪽으로 평행 전극으로 이동하는 것입니다. (B)는 얇은 판 retinotopically 기관의 행렬시각적 공간에서 특정 작은 영역에서 정보를 처리하는 신경 세포들이 즐비 각각화된 카트리지. 신경 중첩으로 인해, 다른 이웃 ommatidia, 개안로부터 6 광 수용체는 L1-L3와 무 축삭 세포 (암)에 히스타민 출력 시냅스를 형성, 동일한 라미 카트리지에 자신의 축삭 (R1-R6)를 보낼 수 있습니다. (C) R1-R6 엑손 단자와 라미 카트리지 복잡한 내부 (L4, L5, Lawf, C2, C3 및 T1 포함) 사이의 interneurons 시각 신경 정보의 확산. (D) R1-R6의 광 수용체의 축삭은 L2 및 L4 극성 세포에서 시냅스 피드백을받을 수 있습니다. 리베라 – 알바 등의 알 2에서 수정 (B)와 (C). 이 그림의 더 큰 버전을 보려면 여기를 클릭하십시오.
초파리의 눈은 신경 중첩 형 (16)이다. 이 t를 의미한다라미와 수질 : 모자는 공간에서 같은 지점에서 볼 ommatidia, 개안 이웃 일곱에 속하는 여덟 광 수용체의 신경 신호는 다음 두 neuropils 한 신경 카트리지에서 함께 풀링됩니다. 여섯 외부 광 수용체 라미에 신경 컬럼에 자신의 축삭 단자 R1-R6 프로젝트 (그림 1), R7 및 R8 세포는이 계층을 무시하고 자신이 수질 열 17-19 대응 시냅스 접촉을하는 동안. 이러한 정확한 배선은 모든 얇은 판 (도 1A-C) 그러자, 비행 초기 비전의 retinotopic 매핑에 대한 신경 기판을 생산하고 수질 열 (카트리지) 공간에서 하나의 지점을 나타냅니다.
라미 1,2,20에서와 무 축삭 세포 (암) : R1-R6의 광 수용체에서 직접 입력은 대형 모노 폴라 셀 (L1, L2 및 L3 중저 소득)에 의해 수신된다. 이 중, L1 및 L2 주요 정보 경로 (그림 1D)를 매개하는 가장 큰 세포는, WHI 있습니다채널의 응답은 온 및 오프 에지를 이동함으로써, 움직임 검출부 (21, 22)의 계산 근거를 형성한다. L2 세포 23, 24의 L1에에 전면 후면 및 전면에서 후면 : 행동 실험은 중간 반면에, 두 개의 경로가 반대 방향의 모션 인식을 용이하게 제안합니다. 연결은 또한 L4 뉴런 이웃 카트리지 (25, 26) 사이의 측 방향 연통 중요한 역할을 할 수 있다는 것을 의미한다. 상호 시냅스는 L2 및 L4 동일한에있는 세포와 두 개의 인접한 카트리지 사이에 발견되었다. 다운 스트림, 각각의 L2 셀과 세 개의 관련 L4 세포가 공통의 목표에 자신의 축삭 프로젝트, 이웃 카트리지에서 입력이 생각하는 수질에있는 TM2 신경 세포는 전후 운동 (27)의 처리를 위해 통합 될 수 있습니다. L1 신경 세포가 모두 갭 접합과 시냅스를 통해 동일한 카트리지 L2S에서 입력을받을 수 있지만, 그들은 직접 L4s 따라서 인접 라미 카트리지에 연결되어 있지 않습니다.
<p클래스 = "jove_content"> R1-R6의 광 수용체의 축색 돌기에 시냅틱 피드백은 L2 / L4 회로에 속하는 뉴런 만 제공 아니라 L1 경로의 1, 2 (그림 1D). 같은 카트리지 연결이 R1과 R2에 L2로부터 L4에서 R5에 선택적으로있는 동안, 모든 R1-R6의 광 수용체 중 하나의 L4 또는 둘 모두 이웃 카트리지에서 시냅스 피드백을받을 수 있습니다. 또한, R1, R2, R4 및 R5로 오전 강한 시냅스 연결이 있고, 신경교 세포는 시냅스 네트워크에 연결되어 있으므로 신경 화상 처리부 (6)에 참가할 수있다. 마지막으로, 얇은 판에서 R1-R6 및 R6 및 R7 / R8 사이의 광 수용체 이웃 연결 축삭 갭 접합은, 각 카트리지 14,20,28에서 비대칭 정보의 표현 및 처리에 기여한다.거의 그대로 초파리에서 개별 광 수용체 및 시각의 interneurons에서 세포 내 전압 녹음은 높은 신호 대 잡음 연구를 제공연결된 뉴런 사이의 빠른 신경 계산의 의미를 만들기 위해 필요한 서브 밀리 초 해상도 3,5,7-10,29,에서 다음 페이지에 계속 데이터. 100 밀리 초 해상도 – 정밀도 수준은 상당히 잡음이 있고 일반적으로 10으로 동작 전류 결상 기술에 의해 불가능하다. 전극은 매우 작고 날카로운 팁을 가지고 있기 때문에 또한,이 방법은 세포 기관에 한정되지 않고, 작은 활성 신경 구조의 직접 기록을 제공 할 수있다; 이러한 패치 클램프 전극의 훨씬 더 큰 팁에 액세스 할 수없는 중저 소득 '수지상 나무 또는 광 수용체의 축색 돌기, 등. 중요한 것은,이 방법은 또한 대부분의 패치 클램프 애플리케이션보다 덜 침습적 구조적 손상이기 때문에 적은 연구 세포의 세포 내 환경 정보 샘플링에 영향을 미친다. 따라서, 기존의 날카로운 미세 기술이 기여하고, 신경 INFOR으로, 근본적인 발견과 원래의 통찰력을 기여 계속했다적절한 시간 규모 메이션 처리; 비전 3-10 우리의 기계론의 이해를 향상시킬 수있다.
초파리 R1-R6의 광 수용체 및 중저 소득 국가에서 생체 세포 내 녹음에 Juusola 실험실에서 수행하는 방법이 문서에서는 설명합니다. 이 프로토콜은, 적절한 전기 생리학 장비를 구성하는 방법에 대해 설명 비행을 준비하고 녹음을 수행합니다. 일부 대표 데이터를 제시하고, 몇 가지 일반적인 문제와 잠재적 인 솔루션은이 방법을 사용할 때 발생 될 수있는 논의된다.
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) |