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.

The fruit fly (Drosophila melanogaster) compound eye is a great model system to investigate the functional organization of photoreceptor and interneuron arrays for neural image sampling and processing, and for animal vision. The system has the most complete wiring diagram1,2 and is amiable to genetic manipulations and accurate neural activity monitoring (of high signal-to-noise ratio and time-resolution)3-10.
The Drosophila eye is modular, containing ~750 seemingly regular lens-capped structures called ommatidia, which together provide the fly a panoramic visual field that covers almost every direction around its head. The eye&#39;s primary information sampling units are its rhabdomeric photoreceptors7,8,11. Each ommatidium contains eight photoreceptor cells (R1-R8), which share the same facet lens but are aligned to seven different directions. Whilst the outer photoreceptors R1-R6 are most sensitive to blue-green light, spectral sensitivities of the inner cells R7 and R8, which lie on top of each other and point to the same direction, exhibit three distinctive subtypes: pale, yellow and dorsal rim area (DRA)12-15.
<img alt="Figure 1" src="/files/ftp_upload/54142/54142fig1.jpg" /> 
Figure 1. Functional Organization of the Drosophila Eye. (A) The two first optic ganglia, retina and lamina, are highlighted in gray inside the fly eye. Retina R1-R6 photoreceptors and lamina Large Monopolar Cells (LMCs: L1-L3) are readily accessible in vivo to conventional sharp microelectrode recordings. The schematic electrode highlights the normal path to record from R1-R6 in the retina. One path to record from LMCs in the lamina is to shift in parallel the electrode to left. (B) Lamina is a matrix of retinotopically organized cartridges, each of which is packed with neurons that processes information from a specific small area in the visual space. Due to neural superposition, six photoreceptors from different neighboring ommatidia send their axons (R1-R6) to the same lamina cartridge, forming histaminergic output synapses to L1-L3 and an amacrine cell (Am). (C) The spread of neural information between R1-R6 axon terminals and the visual interneurons (including L4, L5, Lawf, C2, C3 and T1), inside a lamina cartridge is complex. (D) R1-R6 photoreceptor axons receive synaptic feedbacks from L2 and L4 monopolar cells. (B) and (C) modified from Rivera-alba et al2. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The Drosophila eye is of the neural superposition type16. This means that the neural signals of eight photoreceptors belonging to seven neighboring ommatidia, which look at the same point in space, are pooled together at one neural cartridge in the next two neuropils: the lamina and medulla. While the six outer photoreceptors R1-R6 project their axon terminals to neural columns in the lamina (Figure 1), R7 and R8 cells bypass this layer and make synaptic contacts with their corresponding medulla column17-19. These exact wirings produce the neural substrate for the retinotopic mapping of fly early vision, whereupon every lamina (Figures 1A-C) and medulla column (cartridge) represents a single point in space.
Direct inputs from R1-R6 photoreceptors are received by the Large Monopolar Cells (LMCs: L1, L2 and L3) and the Amacrine Cell (Am) in the lamina1,2,20. Out of these, L1 and L2 are the largest cells, mediating major information pathways (Figure 1D), which respond to On- and Off-moving edges, and thus form the computational basis of the motion detector21,22. Behavioral experiments suggests that at intermediate contrast, the two pathways facilitate motion perception of opposite directions: back-to-front in L1 and front-to-back in L2 cells23,24. Connectivity further implies that L4 neurons may play critical role in the lateral communication between neighboring cartridges25,26. Reciprocal synapses were found between L2 and L4 cells located in the same and two adjacent cartridges. Downstream, each L2 cell and its three associated L4 cells project their axons to a common target, the Tm2 neuron in the medulla, where inputs from neighboring cartridges are believed to be integrated for processing of front-to-back motion27. Although L1 neurons receive input from same-cartridge L2s via both gap junctions and synapses, they are not directly connected to L4s and hence adjacent lamina cartridges.
Synaptic feedbacks to R1-R6 photoreceptor axons are provided only by neurons belonging to the L2/L4 circuits but not the L1 pathway1,2 (Figure 1D). Whilst same-cartridge connections are selectively from L2 to R1 and R2 and from L4 to R5, all R1-R6 photoreceptors receive synaptic feedback from L4 of either or both neighboring cartridges. Furthermore, there are strong synaptic connections from Am to R1, R2, R4 and R5, and glia cells are also synaptically connected to the network and may thus participate in neural image processing6. Finally, axonal gap-junctions, linking neighboring R1-R6 and between R6 and R7/R8 photoreceptors in the lamina, contribute to the asymmetric information representation and processing in each cartridge14,20,28.
Intracellular voltage recordings from individual photoreceptors and visual interneurons in nearly intact Drosophila provide high signal-to-noise ratio data at sub-millisecond resolution3,5,7-10,29, which is necessary for making sense of the fast neural computations between the connected neurons. This level of precision is impossible by current optical imaging techniques, which are significantly noisier and typically operate at 10 - 100 msec&#160;resolution. Furthermore, because the electrodes have very small and sharp tips, the method is not restricted to cell bodies, but can provide direct recordings from small active neural structures; such as the LMCs&#39; dendritic trees or photoreceptor axons, which cannot be accessed by much larger tips of patch-clamp electrodes. Importantly, the method is also structurally less invasive and damaging than most patch-clamp applications, and so affects less the studied cells&#39; intracellular milieu and information sampling. Thus, conventional sharp microelectrode techniques have contributed, and keep on contributing, fundamental discoveries and original insight into neural information processing at the appropriate time scale; improving our mechanistic understanding of vision3-10.
This article explains how in vivo intracellular recordings from Drosophila R1-R6 photoreceptors and LMCs are performed in the Juusola laboratory. This protocol will describe how to construct a suitable electrophysiology rig, prepare the fly, and perform the recordings. Some representative data is presented, and some common issues and potential solutions are discussed that may be encountered when using this method.

<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.

The sharp microelectrode recording method, as adapted here for the Drosophila eye, can be used to reliably quantify neural information sampling and processing in the retina and lamina cells, and communication between them4,5,7,8,10,33. By using it to study encoding in different wild-type stocks, mutants or genetically engineered fly strains, the method has proven its value; not only in quantifying the effects of a mutation, temperature, diet or selected expression3,4,6,9,10,14,30,34, but also in revealing mechanistic reasons for altered visual behaviors14,34. The method is also readily applicable to other insect preparations35,36, empowering neuroethological vision studies. Next we showcase a few examples of its successful applications.
<img alt="Figure 7" src="/files/ftp_upload/54142/54142fig7.jpg" /> 
Figure 7. Voltage Responses of a Fruit Fly R1-R6 Photoreceptor to a Light Pulse at 20 and 25 oC. Because the sharp microelectrode penetrations are often very stable, it is possible to record voltage responses of the same R1-R6 photoreceptor to a given light stimulus at different ambient temperatures by warming or cooling the fly. In our set-ups, the fly-holder is placed on a close-loop Peltier-element-based temperature-control system. This enables us to change the fly&#39;s head temperature in seconds. Higher temperature accelerates the voltage responses and characteristically lowers the resting potential of R1-R6 photoreceptors (as indicated by red arrows). <a href="https://www.jove.com/files/ftp_upload/54142/54142fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Studying the Effect of Temperature on Photoreceptor Output
With a well-designed and vibration-isolated recording system, the method can be used for measuring the effect of temperature on an individual cell&#39;s neural output by warming or cooling the fly. The given example shows voltage responses to a bright 10 msec long pulse, recorded in the same R1-R6 photoreceptor at 20 and 25 oC (Figure 7). As quantified before4,9, warming lowers a photoreceptor&#39;s resting potential in darkness, and accelerates its voltage responses.
<img alt="Figure 8" src="/files/ftp_upload/54142/54142fig8.jpg" /> 
Figure 8. Signaling Performance a Fruit Fly R1-R6 Photoreceptor Improves with Light Intensity.&#160;(A) Photoreceptor output to dim (below) and bright (above; 10,000-times brighter light) repeated naturalistic light intensity time series recorded by the same microelectrode in the same cell at 20 oC. Responses to the bright stimulus are larger, because they integrate more samples, elementary responses (bumps) to single photons4,5,7,8. (B) 20 consecutive one-second-long voltage responses are superimposed. Individual responses (light gray) were taken after the adaptive trends (arrow in A) had receded (dotted box in A). The corresponding response means (the signals) are the darker traces. The difference between the signal and the individual responses is the noise. (C) The cells&#39; signaling performance was quantified by the recordings&#39; Signal-to-Noise Ratios (SNR) using the standard methods4,5,7,8. Photoreceptor output has about 64 Hz broader range of reliable signaling at the bright stimulation (SNR&#39;Bright &#160;&#8805;1, up to 84 Hz ) than at the dim (SNR&#39;Dim &#160;&#8805;1, up to 20 Hz), with signal-to-noise ratio improving greatly; from&#160;SNRDimMAX&#160;= 87 to SNRBrightMAX&#160;= 1,868. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Studying Adaptation and Neural Encoding by Repetitive Stimulation
The noninvasiveness of the method, causing relatively little damage in the retina and lamina structures, makes it ideal for studying the signaling performance of individual cells to different light stimuli in their near natural physiological state in vivo. Figure 8 shows voltage responses of a R1-R6 photoreceptor to a dim and bright repeated naturalistic light intensity time series stimulus at 20 oC, whereas Figure 9 shows responses of another R1-R6 photoreceptor and a LMC to a different naturalistic stimulus at 25 oC. The pre- and postsynaptic recordings were performed separately from two different flies because simultaneous intracellular recordings by two sharp microelectrodes in the same fly, one in the retina and the other in the lamina, are too difficult to be viable30.
<img alt="Figure 9" src="/files/ftp_upload/54142/54142fig9.jpg" /> 
Figure 9. Voltage Responses of a Fruit Fly R1-R6 Photoreceptor and LMC to Repeated Naturalistic Stimulation at 25 oC. (A) R1-R6 (gray) and LMC (black) outputs recorded by different microelectrodes from different flies. (B) Fully light-adapted 20 consecutive pre- (above) and postsynaptic (below) responses to the same naturalistic stimulus pattern with individual responses, shown in light gray and the corresponding response means (the signals) as the darker traces. The difference between the signal and the individual responses is the noise. (C) The cells&#39; signaling performance was quantified by the recordings&#39; Signal-to-Noise Ratios (SNR). LMC output has about 10 Hz broader range of reliable signaling (SNR&#39;LMC &#160;&#8805;1, up to 104 Hz ) than R1-R6 output (SNR&#39;R &#160;&#8805;1, up to 94 Hz). Both signal-to-noise ratios are high (SNRLMCMAX&#160;= 142, SNRRMAX&#160;= 752), and as the recording noise was low, their differences reflect real encoding differences between the cells. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
After the stimulus onset, the recordings typically show fast adapting trends that largely subside within 5-6 sec. From then on, the cells produce highly consistent responses to each 1 sec&#160;long stimulus presentation (each dotted box encloses 20 of these). The repeatability of the responses becomes obvious when these are superimposed (Figure 8B and Figure 9B). Individual responses are the thin gray traces, and their mean the thicker darker trace. The mean response is taken as the neural signal, whereas the neural noise is the difference between the mean and each individual response4,5,9,37,38. The respective signal-to-noise ratios in frequency domain (Figure 8C and Figure 9C) were obtained by Fourier-transforming the signal and noise data chunks into power spectra, and dividing the mean signal power spectrum with the corresponding mean noise power spectrum4,5,9,37,38. Characteristically, the maximum signal-to-noise ratios of the recorded neural outputs to naturalistic stimulation are high (100 - 1,000), and in the most stable preparations with very low recording noise can reach values &#62;&#62;1,000 (e.g.,&#160;Figure 8C). Notice also that warming expands the cells&#39; bandwidth of reliable signaling4 (SNR&#39;Bright&#8805; 1); for example, the relative difference between the two R1-R6s in Figures 8 and 9, respectively, is 10 Hz (84 at 20 oC and 94 Hz at 25 oC).
One can further estimate each cell&#39;s rate of information transfer from its signal-to-noise ratio by using the Shannon equation32, or by calculating the difference between the responses&#39; entropy and noise entropy rates through the triple extrapolation method39. More details about the information theoretical analyses, and their use and limitations specifically with this method are given in the previous publications7,8,39.
<img alt="Figure 10" src="/files/ftp_upload/54142/54142fig10.jpg" /> 
Figure 10.&#160;Voltage Responses of a Killer Fly R1-R6 Photoreceptor and LMC to Repeated Naturalistic Stimulation at 19 oC. (A) R1-R6 (gray) and LMC (black) outputs recorded by the same microelectrode from the same fly; first postsynaptically and later presynaptically, as the electrode was advanced in the eye. (B) 20 consecutive pre- (above) and postsynaptic (below) responses (light gray traces) to the same naturalistic stimulus pattern were captured after initial adaptation (dotted box in A). Their means are the signals (the darker traces on top), while their respective differences to the individual responses give the noise. (C) The corresponding Signal-to-Noise Ratios (SNR) were calculated as in Figures 8C and 9C. LMC output has about a 100 Hz broader range of reliable signaling (SNR&#39;LMCMAX&#160;&#8805;1, up to 234 Hz) than R1-R6 output (SNR&#39;RMAX &#160;&#8805;1, up to 134 Hz). Both signal-to-noise ratios are high (SNRLMCMAX&#160;= 137, SNRRMAX&#160;= 627), and as the same microelectrode was used in the recordings, their differences reflect real differences in the pre- and postsynaptic neural outputs. These results imply that the recording system had low noise, and its influence on the analyses was marginal. <a href="https://www.jove.com/files/ftp_upload/54142/54142fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Neuroethological Vision Studies
The method can also be used to record pre- and postsynaptic voltage responses from the compound eyes of different insect species7,8,35,36 (Figure 10), permitting comparative neuroethological studies of visual information processing. For the presented recording system, the only required adaptation is new preparation-holders, each with a suitably-sized opening for the studied species. These exemplary recordings are from of a female killer fly (Coenosia attenuata). They show intracellular voltage responses of a R1-R6 photoreceptor and LMC to identical repetitive light stimulation, as used for the Drosophila counterparts in Figure 9, but at 19 oC. In this case, both the pre- and postsynaptic data were recorded from the same fly; one after the other, with the same recording electrode (filled with 3 M KCl) first advancing through the lateral lamina before entering the frontal retina. In comparison to the Drosophila data at 25 oC, the Coenosia data - even at the cooler temperature - shows responses with faster dynamics; expanding the range of reliable signaling (signal-to-noise ratio &#62;&#62;1) over a broader frequency range. Such functional adaptations in neural encoding of naturalistic stimuli are consistent with Coenosia&#39;s predatory lifestyle36, which require high-precision spatiotemporal information to attain fast aerial hunting behaviors.

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.

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 ...

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18.6K Views.  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.

Video: Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Recording Behavioral Responses to Reflection in Crayfish

Social behavior depends on sensory input from the visual, mechanical and olfactory systems. One important issue concerns the relative roles of each sensory modality in guiding behavior. The role of visual inputs has been examined by isolating visual stimuli from mechanical and chemosensory stimuli. In some studies (Bruski &amp; Dunham, 1987: Delgado-Morales et al., 2004) visual inputs have been removed with blindfolds or low light intensity, and effects of remaining sensory modalities have been elucidated. An alternative approach is to study the effects of visual inputs in the absence of any appropriate mechanical and chemosensory cues. This approach aims to identify the exclusive role of visual inputs. 
We have used two methods to provide visual stimuli to crayfish without providing chemical and mechanical cues. In one method, crayfish are videotaped in an aquarium where half of the walls are covered in mirrors to provide a reflective environment, and the other half are covered in a non-reflective (matte finish) plastic. This gives the crayfish a choice between reflective and non-reflective environments. The reflective environment provides visual cues in the form of reflected images of the crayfish as it moves throughout half of the tank; these visual cues are missing from the non-reflective half of the tank. An alternative method is to videotape the behavior of crayfish in an aquarium separated by a smaller chamber at each end, with a crayfish in one small chamber providing visual cues and an inert object in the opposite small chamber providing visual input from a non-moving, non-crayfish source. 
Our published results indicate that responses of crayfish to the reflective environment depend on socialization and dominance rank. Socialized crayfish spent more time in the reflective environment and exhibited certain behaviors more frequently there than in the non-reflective environment; isolated crayfish showed no such differences. Crayfish that were housed in same-sex pairs developed a social rank of either dominant or subordinate. Responses to reflection differed between dominant and subordinate crayfish (May &amp; Mercier, 2006; May &amp; Mercier, 2007). Dominant crayfish spent more time on the reflective side, entered reflective corners more frequently and spent more time in reflective corners compared to the non-reflective side. Subordinate crayfish walked in reverse more often on the reflective side than on the non-reflective side. Preliminary data suggest similar effects from visual cues provided by a crayfish in a small adjoining chamber (May et al., 2008).

We have developed two methods for studying effects of visual cues on behavior in the absence of tactile and chemical cues. One method involves videotaping responses of crayfish to reflective walls in an aquarium; the other examines effects of visual inputs provided by a live crayfish behind a transparent partition.

Social behavior depends on sensory input from the visual, mechanical and olfactory systems.  One important issue concerns the relative roles of each ...

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.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

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.

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.

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.

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.

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.

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.

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.

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.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

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Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo