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.
果蝇( 果蝇 )复眼是一个伟大的模型系统来研究神经图像采集和处理感光和中间神经元阵列的功能组织,以及对动物的视力。该系统具有最完整的接线图1,2和是可亲到遗传操作和准确的神经活动监测3-10(高信噪比和时间分辨率)。
果蝇眼睛是模块化的,含有〜750看似普通镜头覆盖的结构,称为小眼,它们共同提供悬挂该占地约它的头几乎每一个方向的全景视野。眼睛的主要信息抽样单位是其rhabdomeric感光7,8,11。每个小眼包含八个感光细胞(R1-R8),它们共享相同的小面透镜,但被对准至七个不同的方向。虽然外光感受器,R1-R6芳E要蓝绿光最为敏感,内细胞R7和R8,它趴在对方并指向同一方向的顶部光谱灵敏度,表现出三个不同的亚型:面色苍白,黄色,背缘区(DRA)12 15。
图 1. 果蝇 眼睛 的功能性组织 。 (A)的两个第一视神经节,视网膜和椎板,以灰色突出显示的蝇眼内。视网膜R1-R6感光细胞和叶片大单极细胞(低收入国家:L1-L3) 在体内常规尖锐的微电极记录容易接近。示意图电极突出,从R1-R6在视网膜记录正常轨道。从低收入国家在椎板记录之一路径是平行的电极转移到左。 (B)叶片是retinotopically器官的矩阵源化墨盒,其中的每一个填充有从在视觉空间中的特定的小区域处理信息的神经元。由于神经叠加,从不同的相邻小眼感光器6送他们的轴突(R1-R6)相同的椎板墨盒,形成组胺输出突触L1-L3和无长突细胞(AM)。 (C)的R1-R6轴突和视觉的interneurons(包括L4,L5,Lawf,C2,C3和T1),内筒层是复杂的神经之间的信息的传播。 (D)R1-R6感光轴突接收来自L2和L4单极细胞突触的反馈。从里维拉阿尔巴等2修改(B)和(C)。 请点击此处查看该图的放大版本。
果蝇眼睛是神经叠加16型。该装置T椎板和髓质:帽子八感光属于7相邻小眼,看起来在空间中的相同点的神经信号,以在接下来的两个髓神经1墨盒汇集在一起。而六个外光感受器,R1-R6项目其轴突末梢神经列在薄片( 图1)中,R7和R8细胞绕过此层,并与它们的相应髓质柱17-19突触联系。这些精确的连线生产用于飞行的早期视觉的视皮层映射神经基板,于是每一个叶片( 图1A-C)和髓质柱(盒)代表在空间中的一个点。
和无长突细胞(AM)在椎板1,2,20:从R1-R6感光体直接输入由大单极细胞(L1,L2和L3低收入国家)接收。出的这些,L1和L2是最大的细胞,介导主要信息通路( 图1D),WHI通道响应于接通和关断移动边,并由此形成所述运动检测器21,22的计算基础。行为实验表明,在中间的对比,这两种途径促进相反方向的运动知觉:后端到前在L1和前到后在L2细胞23,24。连接进一步表示:L4神经元可能在相邻的墨盒25,26之间的横向沟通起到至关重要的作用。位于同一L2和L4细胞和两个相邻盒之间发现倒数突触。下游,每个L 2小区和它的三个相关联的L4细胞投射它们的轴突到公共目标,在髓质,其中来自相邻墨盒输入信Tm2的神经元被集成为前到后运动27的处理。虽然L1的神经元通过两个间隙连接和突触接收来自同一盒L2S输入,它们不直接连接到L4S因此相邻薄层墨盒。
<p类=“jove_content”>被属于在L2 / L4电路的神经元只提供突触反馈到R1-R6感光轴突,但不用L1通路1,2( 图1D)。虽然同一墨盒连接从L2至R1和R2以及从L4到R 5是选择性地,所有的R 1〜R 6的光感受器接收来自任L4或两个相邻的墨盒一样的反馈。此外,还有从上午到R1,R2,R4和R5强突触连接,和神经胶质细胞也突触连接到网络,因此可参与神经图像处理6。最后,轴突间隙路口,连接在相邻的层R1-R6和R6和R7 / R8之间的光感受器,有助于信息不对称表示和处理每个墨盒14,20,28。从在几乎完整的果蝇个体感光细胞和视觉的interneurons细胞内电压记录提供高的信号-噪音r在亚毫秒分辨率3,5,7-10,29,这是必要用于使连接的神经元之间的快速的神经计算的感ATIO数据。的精度这一级是由目前的光成像技术,这是显著喧闹并且通常在10操作是不可能的 – 100毫秒的分辨率。此外,因为电极具有非常小的和尖锐的提示,该方法不限于细胞体,但可提供从小活性神经结构直接记录;如低收入国家'树突树木或感光体轴突,这是不能由大得多膜片钳电极的前端访问。重要的是,该方法也是在结构上比大多数膜片钳应用侵入性较少,并毁坏等影响少的研究细胞的细胞内环境和信息抽样。因此,传统尖锐的微电极技术作出了贡献,并继续做出贡献,根本发现和原来的洞察神经Infor公司在适当的时间尺度息处理;提高我们的愿景3-10的机械理解。
本文介绍了如何从 果蝇 R1-R6感光细胞和低收入国家体细胞内记录在Juusola实验室中进行。该协议将介绍如何构建一个合适的电装备,准备飞行,并进行录音。一些有代表性的数据被呈现,并且一些共同的问题和潜在的解决方案进行了讨论,可能使用该方法时会遇到。
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) |