We would like to thank Ms. Rui Chen for the dissection and images of the Drosophila CNS shown in the figures.

1. Equipment and Materials
<ol>
	<li>Prepare the required components (listed in Table of Materials) of the electrophysiology rig to perform suction electrode recordings of the Drosophila CNS. 
	NOTE: Prior to experimentation, it is necessary to construct chambers for dissection of the Drosophila CNS and to be used for bathing the ganglia in saline during recordings. A step-by-step outline of chamber construction is provided below.</li>
	<li>Prepare the larval chamber.
	<ol>
		<li>Melt the black wax using a hot plate.</li>
		<li>Pour 2 mL of melted wax into a chamber that has a maximum volume of 2 - 2.5 mL.</li>
		<li>Let the wax cool and harden for approximately 2 h.</li>
		<li>Use a razor blade to carve a hole in the hardened wax that will hold a volume of 500 &#181;L, which has a surface area of approximately 0.5 cm2 and a depth of approximately 0.25 cm.</li>
	</ol>
	</li>
</ol>
2. Equipment and Software Configuration
NOTE: The setup of the extracellular recording is briefly described below.
<ol>
	<li>Prepare equipment for dissection and recording.
	<ol>
		<li>Position the tissue preparation, microscope, suction electrode, micromanipulators, light source inside the Faraday cage to reduce noise and eliminate extraneous electrical fields (Figure 2).</li>
		<li>Connect (preferably solder) ground and positive wires onto shielded alligator clips that will be connected to the bath and microelectrode holder, respectively. 
		NOTE: Soldered and exposed wires can be covered with aluminum foil to reduce noise.</li>
		<li>Connect the ground and positive wires to input 1 of the AC/DC differential amplifier.</li>
		<li>Connect the data acquisition system to the AC/DC differential amplifier by connecting a Bayonet Neill-Concelman (BNC) cable from input 1 of the data acquisition system to the channel 1 output of the amplifier. 
		NOTE: It is recommended to incorporate a 50/60 Hz noise eliminator between the data acquisition software and the amplifier. The use of a BNC T-connector is required.</li>
		<li>If desired, include an audio monitor into the setup by connecting input 1 of the audio monitor to input 1 of the data acquisition software.</li>
		<li>Fill the 10 mL syringe with saline and connect it to the pressure port of the electrode holder. 
		NOTE: Ensure that no air bubbles exist between the syringe, the Ag/AgCl pellet, and the electrode opening.</li>
	</ol>
	</li>
	<li>Prepare the software for dissection and recording. 
	NOTE: The outline of methods for software setup is based on the acquisition/analysis software listed in the Table of Materials which will digitize the raw electrical output and convert the data to spike frequency. However, other software can be used and multiple recording conversions can be used.
	<ol>
		<li>Open the acquisition/analysis software.</li>
		<li>Click on &#34;setup&#34; from the main toolbar and select &#34;channel settings,&#34; which will open a dialogue box.</li>
		<li>Reduce the number of total channels to three.</li>
		<li>For Channel 1, select a range of 100 mV.&#160;</li>
		<li>For Channel 2, click on the &#34;calculation&#34; tab that will open a drop-down menu. Select &#34;cyclic measurements,&#34; which will open a second dialogue box.</li>
		<li>Select channel 1 as the source. Then on the drop down menu measurement, select unit spike at events. Finally, in the detection settings area, from the drop down menu select simple threshold.&#160;</li>
		<li>To convert the electrical activity into a rate plot expressed in hertz, the 3rd channel must be set in arithmetic mode: On the input window, type in &#34;1000*smoothsec(ch2,2)&#34;. Then select as the units drop down menu hertz. 
		NOTE: Successful recordings can be performed with multiple different recording setups, but &#34;Unit spike at events and simple threshold&#34; is likely the easiest, since it allows for adjustment of the threshold above the background activity for each individual recording. Recordings under the setting of &#34;Custom-Source Input&#34; increase reproducibility of the data assuming the background activity is not different between recordings.</li>
		<li>Close dialogue boxes to return to the main screen. The y-axis should be expressed in Hz and the x-axis in time.</li>
	</ol>
	</li>
</ol>
3. Dissect and Prepare the Larval Drosophila CNS
NOTE: Methods for larval CNS dissection are clearly illustrated in Hafer and Schedl27, but these previously published methods reduce the length of the descending neurons that are important for measuring spike frequency. Here, an additional method is outlined to excise the larval CNS that maintains long, intact descending neurons.
<ol>
	<li>Saline preparation.
	<ol>
		<li>Prepare the dissection and recording saline to the following in mM28: 157 NaCl, 3 KCl, 2 CaCl2, 4 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid (HEPES); titrate pH to 7.25.</li>
	</ol>
	</li>
	<li>Dissect the larval Drosophila CNS.
	<ol>
		<li>Identify and extract a wandering third-instar Drosophila melanogaster from the culture vial and place into 200 &#181;L of saline (Figure 3A).</li>
		<li>Grasp the mouth hooks with a pair of fine forceps and grasp the abdomen of the maggot with a second pair of forceps (Figure 3B). 
		NOTE: Do not apply too much pressure to the abdomen as this can result in tearing of the thin cuticular layer of the maggot.</li>
		<li>Gently pull the mouth hooks and abdomen in different directions to separate the caudal end of the maggot from the head region and expose the viscera of the maggot (Figure 3C). 
		NOTE: The CNS will be intertwined with the trachea and digestive tract. Do not cut the CNS away from any tissue to prevent cutting the descending peripheral nerves.</li>
		<li>Tease the CNS out of the digestive tract and trachea with forceps (Figure 3D).</li>
		<li>If necessary, disrupt the blood brain barrier by manually transecting the CNS posterior to the cerebral lobes with Vannas spring scissors. 
		NOTE: This should be done based on the physiochemical properties of the chemical being used. The red line in Figure 4A is the suggested transection point and the transected CNS with intact descending peripheral nerve trunks shown in Figure 4B. The transected CNS is ready for experimentation after completion of step 3.2.5.</li>
	</ol>
	</li>
</ol>
4. Extracellular Recording of Drosophila CNS.
<ol>
	<li>Prepare the CNS for recording.
	<ol>
		<li>Pull a glass pipette electrode from borosilicate glass capillaries to a resistance of 5-15 m&#937;.</li>
		<li>Insert the transected CNS into the wax chamber that contains 200 &#181;L of saline. Clamp an uncoated insect pin with the alligator clip soldered to the ground wire and insert the pin into the saline to complete the circuit.</li>
		<li>Using the micromanipulators, orient the electrode to the caudal end of the transected CNS. Eliminate recording of background noise by adjusting the threshold level in the acquisition/analysis software prior to contacting the peripheral nerve trunks.</li>
		<li>Draw any convenient peripheral nerves into the suction electrode by applying slight negative pressure on the syringe. 
		NOTE: The baseline firing frequency is correlated to the number of neurons drawn into the electrode where more neurons oftentimes result in increased resistance of the electrode.</li>
	</ol>
	</li>
	<li>Begin extracellular recording of CNS descending nerve activity.
	<ol>
		<li>Start the recording on the data acquisition software and monitor the baseline firing frequency. Allow the firing rate to equilibrate for 5 min prior to collecting baseline firing rate data.</li>
		<li>Discard the preparation and recording if the pattern of firing of control treatment is not a bursting pattern similar to that shown in Figure 5A. 
		NOTE: Altered pattern of firing suggests abnormal or unstable activity of the central pattern generator, which can alter neural function.</li>
		<li>After 5 min, add 200 &#181;L of saline + vehicle to bring the total volume of the chamber to 400 &#181;L to begin recording control firing rates.</li>
		<li>After baseline has been established (3-5 min), withdraw 200 &#181;L of saline and add 200 &#181;L of the experimental agent solubilized in saline. 
		NOTE: Dimethyl sulfoxide (DMSO) is the recommended solvent for lipophilic compounds and should not exceed 0.1% v/v.</li>
		<li>Apply drug concentrations in a serial manner (low to high concentrations) and record each concentration for a select period of time (determined by the investigator based on the chemical properties of the drug29). Label this time point of drug application in the acquisition/analysis software by including a comment that includes the drug and the final concentration. 
		NOTE: Firing activity of the CNS will decline by approximately 10-20% after 30-50 min after the dissection and therefore, appropriate untreated controls should be performed, and drug treatments should not exceed this time period.</li>
	</ol>
	</li>
	<li>Analyze the data. 
	NOTE: Multiple analyses can be performed on the collected data, such as changes in action potential bursts29, spike waveform amplitude and time course, and determining mean spike frequencies after drug treatment26,30,31,32. The most common is determining the influence a drug has to the mean spike frequencies over a specified period of time, which is described below.
	<ol>
		<li>Tabulate mean spike frequencies by selecting the entire region of interest (i.e., the drug treatment time period) and determine mean spike frequencies automatically through the &#34;Datapad&#34; located on the main toolbar of the acquisition/analysis software.</li>
		<li>Determine the mean spike frequency of the CNS after drug treatment to the mean spike frequency of the vehicle control (baseline). Calculate the percent change of firing rate after drug treatment by the formula: (Treated Frequency/Baseline Frequency) &#215; 100.</li>
		<li>Use the mean spike frequencies or percent firing of control for each concentration to construct a concentration response curve with standard graphing software.</li>
		<li>Perform statistical analysis (e.g., unpaired t-test) to determine significance between time points, concentrations, or drug treatments. 
		NOTE: There are two primary methods for generating and analyzing concentration-response curves. The first method is one concentration per individual preparation. Here, the spike rate of the single concentration is normalized to baseline spike rate for each preparation. The benefits are reduced run down of the preparation due to shortened recording time, while the pitfalls are increased dissection time because this method will consist of 5-7 individual CNS preparations per concentration, and larger error bars on the averaged data set. The second method is multiple concentrations per individual preparation. Here, the spike rate for each of the 3-5 concentrations are normalized to the same baseline spike rate. The benefits are less dissection time and less variability between replicates for drug concentrations, while the pitfalls are the requirement of a fast-acting drug and unknown impact of previous drug concentration treatments on subsequent chemical treatments.</li>
	</ol>
	</li>
</ol>

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</ol>

The authors have nothing to disclose.

The details provided in the associated video and text have provided key steps in order to record the activity and spike discharge frequency of the Drosophila CNS ex vivo. The dissection efficacy is the most critical aspect of the method because short or few descending neurons will reduce the baseline firing rate that will result in large variances between replicates. However, once the dissection technique has been mastered, the data collected with this assay are highly reproducible and amendable for a wide variety of disciplines. One modification to the described method is the inclusion of an automated profusion system that will prevent the need to manually pipette the saline and chemical solutions into the CNS chamber. Inclusion of the profusion system will reduce disturbances of the CNS during the recordings, which occasionally occur during the application of drugs with manual pipettes.
This electrophysiological method exploits the utility of the preparation for incorporation into drug/insecticide discovery research. Furthermore, this preparation is amenable to the classroom for demonstration of fundamental concepts in neurophysiology. The method requires relatively modest financial investment and minimal preparation time while providing a robust and stable recording that can be employed to illustrate the influence of drugs to the function of fly CNS. For instance, the financial burden of the recording rig is an initial cost of approximately $10,000 USD with minor subsequent costs (e.g., saline salts, fly colony maintenance, etc.). Further, the time needed from CNS preparation to first recording is approximately 10 min. Although Drosophila are easy to maintain in culture within a laboratory or classroom, it should be noted that this suction electrode assay can also be performed using the CNS of the housefly, Musca domestica, and probably other muscoid fly larvae, as well. The pest status of Musca domestica to livestock suggests this assay could be of significant utility to research programs aiming to characterize the neuronal sensitivity of flies from different populations or to determine the potency of newly developed neurotoxicants for eventual control of infestations.
In addition to characterizing the potency of drugs, this preparation can be used to characterize the influence of genetic mutations and manipulation on the activity of the Drosophila nervous system. Previous work has shown that CNS-specific knockdown of the gene encoding the inward rectifier potassium (Kir) channel increased the baseline CNS firing frequency by approximately 2-fold when compared to control flies31. These data were combined with pharmacological data to ultimately speculate the physiological role Kir channels serve to Drosophila nervous system function.
Although this technique represents a powerful assay to test diverse toxicological and physiological hypotheses, limitations to the assay do exist and must be considered. For instance, the data generated through suction electrode recordings rarely provide conclusive evidence for the exact mode of action of a drug or precise physiological role for an ion channel and must be studied in tandem with more cell-based measurements, such as patch clamp electrophysiology. An additional limitation to this method is that the various receptors and ion channel sub-types do not influence the CNS spike rate in an equal manner. Therefore, it is possible that a modulation of a specific receptor or ion-channel subtype may not significantly influence the spike rate of the excised CNS, but may indeed have a critical effect on total nervous system function and/or integration of signals.
Although limitations exist, the data collected through suction electrode recordings do provide quality proof-of-concept data and allow researchers to generate hypotheses regarding mechanisms of action that can be validated through voltage-clamp electrophysiology, biochemical analyses, or through additional pharmacological testing. For example, the latter was performed to test the hypothesis that the insect repellent, N,N-Diethyl-3-methylbenzamide (DEET), was inhibiting the octopaminergic system in insect CNS in an effort to describe the mechanism of toxicity to mosquitoes and flies26. DEET was shown to be a neuroexcitant of housefly CNS activity and also altered the evoked excitatory postsynaptic potential at the neuromuscular junction, which are cholinergic and glutamatergic systems, respectively26. These data suggested that DEET was not likely to induce toxicity through cholinergic inhibition, as was previously suggested34. Recording the discharge frequency of the fly CNS after exposure to phentolamine, a known octopamine antagonist, showed a complete inhibition of the DEET-mediated neuroexcitation, but not that of propoxur, which provided significant evidence that DEET was more likely affecting the octopaminergic system than AChE26. These published data sets highlight the variety of hypotheses and experimental conditions that can be investigated with this method.

The overall goal of this approach is to enable researchers to quickly measure the electrogenesis of the central nervous system (CNS) in the model insect, Drosophila melanogaster. This method is reliable, quick, and cost-efficient to perform physiological and toxicological experimentation. The CNS is essential for life and therefore, the physiological pathways critical for proper neural function have been explored extensively in an effort to understand or modify neural function. Characterization of the signaling pathways within the arthropod CNS has enabled the discovery of several chemical classes of insecticides that disrupt invertebrate neural function to induce mortality while limiting off-target consequences. Thus, the ability to measure the neural activity of insects is of significant interest to the field of insect toxicology and physiology since the nervous system is the target tissue of the majority of deployed insecticides1. Yet, continued growth of fundamental and applied knowledge regarding the insect nervous system requires advanced neurophysiological techniques that are limited in feasibility, since current techniques are labor intensive and require a high expense, insect neural cell lines are limited, and/or there is limited access to the central synapses of most arthropods. Currently, characterization of most insect neural proteins requires the target to be cloned and heterologously expressed for subsequent drug discovery and electrophysiological recordings, as was described for insect inward rectifier potassium channels2, insect ryanodine receptor3, mosquito voltage-sensitive K+ channels4, and others. To mitigate the requirement for heterologous expression and the potential for low functional expression, Bloomquist and colleagues aimed to induce a neuronal phenotype in cultured Spodoptera frugiperda (Sf21) cells as a novel method for insecticide discovery5,6. These methods provide a valid approach for the development of new chemistry, yet they oftentimes create an insurmountable bottleneck for the characterization of pharmacological agents, identifying mechanisms of insecticide resistance, and characterization of fundamental physiological principles. Here, we describe an ex vivo method that enables the recording of electrical activity from a model insect that has malleable genetics7,8,9 and known expression patterns of neural complexes10,11,12 to enable the characterization of resistance mechanisms at the level of the nerve, the mode of action of newly developed drugs, and other toxicological studies.
The fruit fly, D. melanogaster, is a common model organism for defining insect neural systems or insecticide mechanism of action and has been established as a well-suited model organism for the study of toxicological13, pharmacological14,15, neurophysiological16, and pathophysiological17,18,19,20 processes of vertebrates. D.melanogaster is a holometabolous insect that performs complete metamorphosis, including a larval and pupal stage before reaching the reproductive adult stage. Throughout the developmental process, the nervous system undergoes significant remodeling at different life stages, but the larval CNS will be the focus of this methodology. The fully developed larval CNS is anatomically simple with thoracic and abdominal segments that are fused and form the ventral ganglion, which represents an array of repeated and almost identical neuromeric units21,22. Descending motor nerves originate from the caudal end of the subesophageal ganglia and descend to innervate body wall muscles and visceral organs of the larvae. Figure 1 describes the gross anatomy of the larval Drosophila CNS.
The Drosophila blood-brain barrier (BBB) develops at the end of embryogenesis and is formed by subperineurial glial cells (SPG)21. The SPG cells form numerous filopodia-like processes that spread out to establish a contiguous, very flat, endothelial-like sheet that covers the entire Drosophila CNS23. The Drosophila BBB has similarities to the vertebrate BBB, which includes preserving the homeostasis of the neural microenvironment by controlling the entry of nutrients and xenobiotics into the CNS21. This is a prerequisite for reliable neural transmission and function, yet the protection of the CNS by the BBB restricts the permeation of synthetic drugs, most peptides, and other xenobiotics24,25, which introduces potential problems when characterizing potencies of small-molecule modulators. The method uses a simple transection to disrupt this barrier and provide ready pharmacological access to the central synapses. 
The greatest strength of the described methodology is the simplicity, reproducibility, and relatively high-throughput capacity inherent to this system. The protocol is relatively easy to master, the setup requires little space, and only an initial financial input is necessary which is reduced to reagents and consumables. Further, the described method is completely amendable to record the central descending nerve activity of the house fly, Musca domestica26.

<table><tbody><tr><td>&lt;em&gt;Drosophila melanogaster&lt;/em&gt; (strain OR)</td><td>Bloomington Drosophila Stock Center</td><td>2376</td><td></td></tr><tr><td>Vibration isolation table</td><td>Kinetic Systems</td><td>9200 series</td><td></td></tr><tr><td>Faraday Cage</td><td>Kinetic Systems</td><td>N/A</td><td></td></tr><tr><td>Dissecting Microscope on a Boom</td><td>Nikon</td><td>SMZ800N</td><td>Multiple scopes can be used; boom stand is critical</td></tr><tr><td>AC/DC differential amplifier</td><td>ADInstruments</td><td>AM3000H</td><td>The model 1700 can be used instead of the model 3000</td></tr><tr><td>audio monitor</td><td>ADInstruments</td><td>AM3300</td><td></td></tr><tr><td>Hum Bug Noise Eliminator</td><td>A-M Systems</td><td>726300</td><td></td></tr><tr><td>Data Acquisition System (PowerLab)</td><td>ADInstruments</td><td>PL3504</td><td>Multiple PowerLab models can be used.</td></tr><tr><td>Lab Chart Pro Software</td><td>ADInstruments</td><td>N/A - Online Download</td><td></td></tr><tr><td>Fiber Optic Lights</td><td>Edmund Optics</td><td>89-740</td><td>Different light sources can be used, but fiber optics are the most adaptable</td></tr><tr><td>Micromanipulator</td><td>World Precision Instruments</td><td>M325</td><td></td></tr><tr><td>Microelectrode Holder</td><td>World Precision Instruments</td><td>MEH715</td><td>Different models are acceptable</td></tr><tr><td>BNC cables</td><td>World Precision Instruments</td><td>multiple based on size</td><td></td></tr><tr><td>Glass Capillaries</td><td>World Precision Instruments</td><td>PG52151-4</td><td></td></tr><tr><td>Microelectrode Puller</td><td>Sutter Instruments</td><td>P-1000</td><td>Also can use Narashige PC-100</td></tr><tr><td>Black Wax</td><td>Carolina Biological Supply</td><td>974228</td><td></td></tr><tr><td>Non-coated insect pins, size #2</td><td>Bioquip</td><td>1208S2</td><td></td></tr><tr><td>Fince Forceps</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Vannas Spring Scissors</td><td>Fine Science Tools</td><td>15000-03</td><td></td></tr></tbody></table>

electrophysiological recording of the central nervous system activity of third-instar drosophila melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Spontaneous activity of the descending peripheral nerves arising from the Drosophila central nervous system can be recorded using extracellular suction electrodes with consistent reproducibility. Spontaneous activity of the excised and transected Drosophila CNS produces a cyclical pattern of bursting with 1-2 s of firing with approximately 1 s of near quiescent activity. For example, the CNS is near quiescent (1-2 Hz) for 0.5-1 s, followed by a burst (100-400 Hz) for approximately 1 s, and then returns to a near quiescent state (1-2 Hz) for 0.5-1 s (Figure 5A). This firing pattern is repeated every 2-3 s. The average spike discharge frequency of Drosophila CNS ranges from approximately 20-50 Hz over a 3-5 min period, when the threshold is set just above the baseline noise. The baseline firing frequency is correlated to the number of peripheral neurons drawn into the electrode and the seal formed between the electrode orifice and the ventral ganglia. Importantly, application of the chemical solvent DMSO at a final concentration of 0.1% does not alter the spike discharge frequency of the Drosophila CNS (Figure 5A).
This electrophysiological preparation provides a method to characterize the excitatory or depressant properties of various small molecules on the spike frequency of a well-characterized insect neural system. Propoxur, a known inhibitor of insect acetylcholinesterase (AChE), is a neuroexcitant and increased the spike discharge frequency of the transected Drosophila CNS in a concentration-dependent manner (Figure 5B). On the contrary, gamma-aminobutyric acid (GABA), a known inhibitory neurotransmitter acting upon the insect GABA-mediated chloride channel, is a neurodepressant and reduced the spike discharge frequency in a concentration-dependent manner (Figure 5C). Mean spike discharge frequencies can be determined across the recorded time period for each concentration to enable the construction of a concentration response curve to determine the 50% effective concentration (EC50) to elicit a response. Here, propoxur is shown to have an EC50 of 338 nM (95% confidence interval (CI): 241-474; hillslope: 1.8; r2: 0.77) with a concentration of 1 &#181;M producing maximal excitation of the Drosophila CNS at 300% activity of control (Figure 5D)26. GABA is shown to have an IC50 of 1.1 mM (95% CI: 0.7-1.5; hillslope: 1.5; r2: 0.95) with maximal inhibition at 5 mM (Figure 5E).
Oftentimes, molecular probes of neural systems are not able to be used in physiological or toxicological assays due to their lack of proper physiochemical properties that enable penetration of the blood brain barrier. For instance, monomeric tacrine is a potent (ca. 200 nM) inhibitor of insect AChE33, yet does not alter the spike frequency of the intact Drosophila CNS (Figure 6A). However, disruption of the neural lamella and the blood brain barrier through transection posterior to the cerebral lobes resulted in a near immediate increase in the spike frequency of the Drosophila CNS after exposure to 100 &#181;M monomeric tacrine (Figure 6B). Similar data have been previously described30.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58375/58375fig1.jpg" /> 
Figure 1: Excised CNS from third-instar Drosophila melanogaster. Arrows point to various anatomical structures of the CNS that correspond to the labels. The scale bar represents 250 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58375/58375fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58375/58375fig2.jpg" /> 
Figure 2: Electrophysiology setup that is used to perform extracellular recordings. (A) Faraday cage; (B) vibration table; (C) dissecting microscope; (D) AC/DC differential amplifier; (E) audio monitor; (F) noise eliminator; (G) data acquisition system; (H) computer running lab chart pro software; (I) fiberoptic cable with external illumination source; (J) micromanipulator; (K) microelectrode holder with pressure port with glass electrode and preparation wax dish. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58375/58375fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58375/58375fig3.jpg" /> 
Figure 3: Method for excising the CNS from third-instar maggots. (A) Intact maggot submerged in 200 &#181;L of saline. The arrow indicates the mouth hooks that are used for separation of the body wall. (B) Two pairs of forceps are placed at the middle of the maggot and on the mouth hooks to begin separation of the body wall. (C) Body wall is separated by applying slight and continuous pressure to expose the viscera. (D) CNS is clearly visible (white arrows) and is occasionally intertwined with the viscera. The scale bar represents 1000 &#181;m, 750 &#181;m, 500 &#181;m, and 200 &#181;m for panels A, B, C, and D, respectively. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58375/58375fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58375/58375fig4.jpg" /> 
Figure 4: Disruption of the blood brain barrier by transecting the CNS. (A) Intact CNS with descending nerves clearly visible at the caudal end of the ventral ganglia. The red line indicates the location of transecting the CNS to disrupt the BBB. (B) A transected CNS with the caudal end of the ventral ganglia still exposing long descending neurons. The ventral ganglia can be discarded. The scale bar represents 200 &#181;m for both panels. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58375/58375fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58375/58375fig5.jpg" /> 
Figure 5: Neurophysiological recordings from the CNS of third-instar larvae of D. melanogaster. Representative nerve discharge traces before and after exposure to (A) DMSO, (B) propoxur, and (C) GABA. Initial firing frequencies in spikes/s (Hz) for each experiment are given to the left of each trace. Concentration-response curves for propoxur (D) and GABA (E) to CNS nerve discharge of D. melanogaster larvae from replicated recordings (n = 3-5 concentrations per curve, with each concentration replicated at least 5 times). Arrows represent point of drug application. Data points represent mean percent increase of baseline firing rate and error bars represent standard deviation. When error bars are absent, it is because they are smaller than the size of the symbol. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58375/58375fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58375/58375fig6.jpg" /> 
Figure 6: Increased penetration of tacrine into the nervous system after transection of the CNS. Representative recordings of (A) intact and (B) transected larval CNS exposed to monomeric tacrine, which was applied at the arrow. Initial firing frequencies in spikes/s (Hz) for each experiment are given to the left of each trace. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58375/58375fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster at JoVE.com

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

electrophysiological-recording-central-nervous-system-activity-third

Nanyang Technological University

Research

JoVE Journal

Neuroscience

11.5K Views.  Louisiana State University AgCenter. Deze methode kan helpen bij het beantwoorden van belangrijke vragen op het gebied van insectentoxicologie en insectenfysiologie door het meten van de elektrogenese van het Drosophila melanogaster centrale zenuwstelsel. Dit maakt het mogelijk om een breed scala van wetenschappelijke hypothesen te testen en te helpen bij de ontdekking van nieuwe insecticide werkingswijzen. Het belangrijkste voordeel van deze techniek is dat het een eenvoudige, reproduceerbare en relatief hoge doorvoer systeem m...

Video: Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

Whole Cell Recordings from Brain of Adult Drosophila

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by describing the dissecting solution and capture of the adult females used in our studies. The procedure for removing the whole brain intact, including both optic lobes, is illustrated. Dissection of the overlying trachea is also shown.  The isolated brain is not only small but needs special care in handling at this stage to prevent damage to the neurons, many of which are close to the outer surface of the tissue. We show how a special holder we developed is used to stabilize the brain in the recording chamber. A standard electrophysiology set up is used for recording from single neurons or pairs of neurons. A fluorescent image, viewed through the recording microscope, from a GAL4 line driving GFP expression (GH146) illustrates how projection neurons (PNs) are identified in the live brain. A high power Nomarski image shows a view of a single neuron that is being targeted for whole cell recording. When the brain is successfully removed without damage, the majority of the neurons are spontaneously active, firing action potentials and/or exhibiting spontaneous synaptic input. This in situ preparation, in which whole cell recording of identified neurons in the whole brain can be combined with genetic and pharmacological manipulations, is a useful model for exploring cellular physiology and plasticity in the adult CNS.

This video demonstrates the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons using standard whole cell technology. It includes images of GFP labeled cells and neurons viewed during recording.

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by ...

Biology

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" (or "startle") response during the adult stage is mediated by the multi-component neuronal circuit called the Giant Fiber System (GFS). The comparative large size of the neurons, their distinctive morphology and simple connectivity make the GFS an attractive model system for studying neuronal circuitry. The GFS pathway is composed of two bilaterally symmetrical Giant Fiber (GF) interneurons whose axons descend from the brain along the midline into the thoracic ganglion via the cervical connective. In the mesothoracic neuromere (T2) of the ventral ganglia the GFs form electro-chemical synapses with 1) the large medial dendrite of the ipsilateral motorneuron (TTMn) which drives the tergotrochanteral muscle (TTM), the main extensor for the mesothoracic femur/leg, and 2) the contralateral peripherally synapsing interneuron (PSI) which in turn forms chemical (cholinergic) synapses with the motorneurons (DLMns) of the dorsal longitudinal muscles (DLMs), the wing depressors. The neuronal pathway(s) to the dorsovental muscles (DVMs), the wing elevators, has not yet been worked out (the DLMs and DVMs are known jointly as indirect flight muscles - they are not attached directly to the wings, but rather move the wings indirectly by distorting the nearby thoracic cuticle) (King and Wyman, 1980; Allen et al., 2006). The di-synaptic activation of the DLMs (via PSI) causes a small but important delay in the timing of the contraction of these muscles relative to the monosynaptic activation of TTM (~0.5 ms) allowing the TTMs to first extend the femur and propel the fly off the ground. The TTMs simultaneously stretch-activate the DLMs which in turn mutually stretch-activate the DVMs for the duration of the flight. The GF pathway can be activated either indirectly by applying a sensory (e.g."air-puff" or "lights-off") stimulus, or directly by a supra-threshold electrical stimulus to the brain (described here). In both cases, an action potential reaches the TTMs and DLMs solely via the GFs, PSIs, and TTM/DLM motoneurons, although the TTMns and DLMns do have other, as yet unidentified, sensory inputs. Measuring "latency response" (the time between the stimulation and muscle depolarization) and the "following to high frequency stimulation" (the number of successful responses to a certain number of high frequency stimuli) provides a way to reproducibly and quantitatively assess the functional status of the GFS components, including both central synapses (GF-TTMn, GF-PSI, PSI-DLMn) and the chemical (glutamatergic) neuromuscular junctions (TTMn-TTM and DLMn-DLM). It has been used to identify genes involved in central synapse formation and to assess CNS function.

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

The Giant Fiber System is a simple neuronal circuit of adult Drosophila melanogaster containing the largest neurons in the fly. We describe the protocol for monitoring synaptic transmission through this pathway by recording post synaptic potentials in dorsal longitudinal (DLM) and tergotrochanteral (TTM) muscles following direct stimulation of the Giant Fiber interneurons.

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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 ...

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.

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.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

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, ...

Electrophysiological Recording of Voltage Responses of Drosophila Retinal Photoreceptors to Light Stimuli

Source: Juusola, M., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/54142">Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo.</a> J. Vis. Exp. (2016).
This video outlines the protocol for electrophysiological recording of photoreceptor activity in the eye of an immobilized Drosophila, enabling precise measurement of voltage changes in response to light stimuli.

1. Recording from R1-R6 Photoreceptors 

	Always be grounded when operating the microelectrode amplifier (for example by touching the metal surface of the ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

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