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.

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。 <img alt="图1" src="/files/ftp_upload/54142/54142fig1.jpg" /> 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）。 <a href="https://www.jove.com/files/ftp_upload/54142/54142fig1large.jpg" target="_blank">请点击此处查看该图的放大版本。</a> 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”&gt;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&#39;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。

<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.

锋利微电极记录方法中，作为此处适于果蝇眼睛，可以用于可靠地量化它们4,5,7,8,10,33之间在视网膜和薄层细胞神经信息采样和处理，以及通信。通过使用它来研究不同的野生型股票，突变或基因工程菌蝇编码，该方法已证明了其价值;不仅在量化的突变，温度，饮食或选定表达3,4,6,9,10,14,30,34的影响，而且也揭示了改变的视觉行为14,34机械的原因。该方法也容易适用于其他昆虫的准备工作35,36，赋予神经行为学研究的视野。接下来，我们展示的其成功的应用的几个例子。 <img alt="图7" src="/files/ftp_upload/54142/54142fig7.jpg" /> 图7. 电压一个果蝇R1-R6感光的反应，在20的光脉冲和25℃。由于尖锐的微电极穿透通常非常稳定，所以可以在同一R1-R6感光体电压响应在记录到一个给定的光刺激不同的环境温度下通过加热或冷却该飞。在我们的调校，蝇保持器被置于一个闭环基于珀尔帖元件的温度控制系统。这使我们能够改变飞的头温度为秒。较高的温度加速电压响应和典型降低R1-R6光感受器的静息电位（用红色箭头所示）。 <a href="https://www.jove.com/files/ftp_upload/54142/54142fig7large.jpg" target="_blank">请点击此处查看该图的放大版本。</a> 研究温度对感光输出的影响 25℃（ 图7）记录在同一个R1-R6感光电压响应。由于4,9之前量化，气候变暖降低黑暗感光的静息电位，并加速其电压响应。 <img alt="图8" src="/files/ftp_upload/54142/54142fig8.jpg" /> 图8. 信令性能果蝇R1-R6感光随光照强度提高 （A）感光输出变暗（下图）和光明（以上; 10,000次亮的光）通过重复同样的微电极记录的自然光强度的时间序列在20℃。在同一小区回应明亮的刺激较大，因为它们集成更多的样本，基本反应（碰撞），单光子4,5,7,8。 （B）连续20个一秒长的电压响应叠加。个体反应（浅灰色）后的自适应趋势采取（以A箭头）已消退（在一个虚线框）。相应的响应的装置（的信号）是较暗的痕迹。信号和单独的响应之间的差异是噪声。 （C）将细胞使用标准方法4,5,7,8信号-噪声比（SNR）的&#39;信令性能通过录音量化“。感光体输出具有在明亮的刺激“比在昏暗的（ 明亮 ≥1，高达84赫兹（SNR约64赫兹范围更广可靠的信令的SNR），” 昏暗 ≥1，高达20赫兹），其中signal-信噪比改善大大;从SNR 昏暗 MAX = 87 SNR 亮 MAX = 1868。 <a href="https://www.jove.com/files/ftp_upload/54142/54142fig8large.jpg" target="_blank">请点击此处查看该图的放大版本。</a> 通过反复刺激学习适应和神经编码 的方法中，使在视网膜和薄层结构相对较少损害的无创性，使得它非常适合研究各个细胞在体内其邻近自然生理状态不同的光刺激的信令性能。 图8示出R1-的电压响应R6光感受器昏暗和明亮的反复自然光强度的时间序列刺激在20℃，而图9 </strong&gt;的信息显示，在25 摄氏度的另一个R1-R6光感受器和LMC到不同的自然刺激的反应前和突触后录音被因为同时细胞内记录在同一飞两头尖微电极，一个在视网膜和其它在椎板，太难可行30从两个不同的苍蝇分开执行。 <img alt="图9" src="/files/ftp_upload/54142/54142fig9.jpg" /> 要反复刺激自然主义在25 摄氏度 （A）R1-R6（灰色）和LMC（黑色） 果蝇R1-R6光感受器和LMC 图 9. 电压输出的响应由来自不同苍蝇不同的微电极记录下来。 （B）全光适应连续20前（上图），并与个体反应一样自然的刺激模式突触后（下）的反应，在光显示灰色的D中的相应的响应的装置（的信号）作为暗痕迹。信号和单独的响应之间的差异是噪声。 （C）的细胞信号-噪声比（SNR）的&#39;信令性能通过录音量化“。 LMC输出具有“比R1-R6输出（SNR（LMC≥1，高达104赫兹- [R≥1，高达94赫兹）约10赫兹范围更广的可靠信号的信噪比 ）”。这两个信号与噪声的比值高（SNR LMC MAX = 142， 信噪比读取 最大 = 752），并作为记录噪音低，他们之间的分歧反映了细胞之间的真正的编码区别。 <a href="https://www.jove.com/files/ftp_upload/54142/54142fig9large.jpg" target="_blank">请点击此处查看大图这个数字。</a> 刺激发病后，录音典型LY秀快适应了5-6秒内消退主要趋势。从那时起，在细胞产生的每个1秒长刺激呈现（各虚线框包围的这些20）高度一致的反应。当这些叠加（ 图8B和图9B）的响应的可重复性变得明显。个体反应是细灰的痕迹，他们的意思是较厚的深色痕迹。平均响应被作为神经信号 ，而神经噪声的平均值和每个单独的响应4,5,9,37,38之间的差。在频域中的相应的信号对噪声比（ 图8C和图9C）被傅立叶变换的信号和噪声的数据块到功率谱，并除以对应的平均噪声功率谱4的平均信号功率谱得到5,9,37,38。特征，最大信号 - 噪声比öF中的记录，神经输出到自然的刺激是高（100 - 1000），并且在具有非常低记录噪声最稳定的制剂可达到的值&gt;&gt; 1000（ 例如 ， 图8C）。还要注意的是气候变暖扩大了细胞的信号的可靠带宽4（SNR&#39; 光明 ≥1）;例如，在图8 和9两R1-R6S之间的相对差，分别是10赫兹（84在20℃，并在25℃下94赫兹）。 一个可以通过使用香农公式32，或通过三重外推法39计算的响应“熵和噪声熵率之间的差进一步估计距其信噪比信息传输的每个细胞的速率。有关信息理论分析的更多细节，以及它们的使用和限制š专门用这种方法在以前的出版物7,8,39中给出。 <img alt="图10" src="/files/ftp_upload/54142/54142fig10.jpg" /> 杀手粉煤灰，R1-R6光感受器和LMC的 图10 的电压响应反复自然主义刺激在19℃。（A）中，R1-R6（灰色）和LMC（黑色）输出从同一飞同一微电极记录;第一突触后和突触前后，作为电极是在眼推进。 （B）连续20前（上）和突触后（下）的反应（浅灰色的痕迹），以同样的自然刺激模式进行了初步适应（在一个虚线框）后抓获。他们的手段是信号（在顶部的暗的痕迹），而他们各自的个体反应的差异给出的噪音。 （C）的相应的Signa1对噪声比（SNR），计算为在图8C和9C所示 。 LMC输出有关于“比R1-R6输出（LMC MAX≥1，高达234赫兹（SNR 100赫兹范围更广的可靠信号的信噪比 ）&#39;R MAX≥1，最高134赫兹）。两个信号-噪声比是高信噪比（SNR LMC MAX = 137， 信噪比读取 最大 = 627），并作为同一微电极是在录音使用时，它们的差异反映了前和突触后神经输出的实际差别。这些结果表明，该系统记录了低噪音，及其对分析的影响是微不足道的。 <a href="https://www.jove.com/files/ftp_upload/54142/54142fig10large.jpg" target="_blank">请点击此处查看该图的放大版本。</a> 神经行为学研究视野 </p&gt; 该方法也可以用来从不同的昆虫物种7,8,35,36的复眼（ 图10）记录前和突触后电压响应，允许可视信息处理的比较神经行为学研究。对于所提出的记录系统中，唯一需要的适应是新制备持，各与用于研究的物种适宜尺寸的开口。这些示范录音是从女杀手飞（ 秽蝇attenuata）的。它们表现出R1〜R6感光体和LMC到相同的重复光刺激的细胞内电压响应，作为用于果蝇对应于图9中 ，但在19℃。在这种情况下，无论是前和突触后的数据是从同一飞行记录;一前一后，用同样的记录电极（填充用3M KCl）中通过侧面薄层第一前进之前进入ING额视网膜。在25℃时， 秽蝇数据相比于果蝇数据-即使在冷却器温度-示出了具有更快的动力学响应;扩大可靠的信令（信噪比&gt;&gt; 1）在较宽的频率范围内的范围。在自然刺激神经 ​​编码这种功能的修改与秽蝇的掠夺性生活36，这需要高精度的时空信息，以实现快速的空中狩猎行为是一致的。

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.

为记录细胞内电压响应电法<em&gt;果蝇</em&gt;光感受器和中间对光刺激<em&gt;在体内</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 次观看.  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.

视频: 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.

双突触诱发的星形胶质细胞和神经元的反应，在急性海马脑片电生理记录

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.

对于系统电化学神经记录电极和电生理评价方法

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.

电生理记录从<em&gt;果蝇</em&gt; Labellar味感器

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.

指南<em&gt;在体内</em&gt;单单元从Optogenetically鉴定皮质抑制性录像

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.

分析突触调制<em&gt;果蝇</em&gt;曝光时间延长后，光光感受器

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.

全细胞电压钳记录的电生理方法<em&gt;果蝇</em&gt;光感受器

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.

电生理记录<em&gt;果蝇</em&gt; Trichoid Sensilla响应低挥发性气味

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.

前体钙成像用于可视化大脑对果蝇内分泌信号的反应

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.

在行为和刺激反应期间为整个大脑成像准备成人德罗索菲拉褪黑麦片加斯特

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.

在活体钙成像小鼠致病性甘里昂神经元对味觉刺激的反应

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