Supported by the NIH grant R01 MH 099557

1. Fabrication of Recording Electrodes
<ol>
	<li>Pulling the glass electrodes
	<ol>
		<li>Use the following protocol for the microelectrode puller (see Table of Materials):
		<ol>
			<li>Line 1: Heat 510 Pull - Velocity 30 Time 250; Line 2: Heat 490 Pull - Velocity 30 Time 250. 
			NOTE: Time units correspond to 0.5 ms per unit; the other units are relative. The value of the heat should be adjusted for every filament after the ramp test is performed.</li>
		</ol>
		</li>
		<li>Use a microscope (35x magnification) to ensure that the inner diameter of the pulled electrode is in the range of 7 - 10 &#181;m (Figure 1A). Store the capillaries in tightly closed containers to prevent dust accumulation.</li>
	</ol>
	</li>
	<li>Fire polishing
	<ol>
		<li>Fire polish capillaries (80 - 90% of the maximum heat value for 1 - 2 s) using a micro-forge (see Table of Materials). Ensure the final inner diameter of the polished electrode is 5 &#181;m (Figure 1A)27.</li>
	</ol>
	</li>
	<li>Bending 
	NOTE: Two bends are made in order to position the electrode on the top of the muscle under a high magnification objective.
	<ol>
		<li>Use the apparatus shown in Figure 1B. Fix the electrode in the manipulator and position the tip over the filament, not touching it.</li>
		<li>Set heat value to 60 - 70% of the maximum and press the pedal of the micro-forge for 1 - 2 s (see Table of Materials) to heat the filament. Use an L-shaped needle to gently pull down the tip of the electrode (Figure 1B enlarged). Make the bend at approximately 90o (Figure 1C).</li>
		<li>Hold the electrode over the flame of the torch by hand using forceps and make the second bend at a distance of 7 - 10 mm from the first bend and at an angle of approximately 120o (Figure 1C).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56493/56493fig1.jpg" /> 
Figure 1. Final steps of micropipette fabrication. (A) Electrode tips after pulling and fire polishing. (B.1) The setup for tip bending. (B.2) The boxed area is shown enlarged on the right. The electrode and the filament are fixed so that the electrode tip is positioned slightly above the wire and not touching. (C.1) The recording electrode. (C.2) The boxed area is shown enlarged on the right. The first bend has an angle of approximately 90&#176;, and the distance between the first bend and the tip of the electrode is approximately 1 mm. Scale bar = 3 mm. <a href="//cloudfront.jove.com/files/ftp_upload/56493/56493fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Additional Preparatory Steps
<ol>
	<li>Prepare heamolymph-like (HL3) solution (in mM): 70 NaCl, 5 KCl, 20 MgCl2, 10 NaHCO3, 5 trehalose, 115 sucrose, 5 HEPES, and 1 mM CaCl2; adjust the pH to 7.3 - 7.4. Keep the solution in the refrigerator and make it fresh every week.</li>
	<li>Make stimulation pipets the same way as recording glass pipets, except there is no need to bend them. 
	NOTE: The preparation of stimulation electrodes is described in detail in 28 and 27. The final diameter after fire polishing should be in the range of 5 - 7 &#181;m.</li>
	<li>Insert a stimulation pipette in a microelectrode holder connected to a syringe.</li>
	<li>Manufacture dissection plates from small petri dishes (35 x 10 mm) coated with silicone rubber epoxy (see Table of Materials) as described in 28. 
	NOTE: Silicone rubber should be completely hardened before use.</li>
	<li>Pick a wandering third-instar larvae and dissect it as described in27,28,29,30. 
	NOTE: To visualize boutons, use the fly strain CD8-GFP (see Table of Materials)
	<ol>
		<li>Using forceps (see Table of Materials), pick the 3rd instar larvae from a vial.</li>
		<li>Pin the larvae, placing the first pin posterior and another pin anterior (close to mouth hooks).</li>
		<li>Add HL3 solution.</li>
		<li>Make a cut using spring scissors (see Table of Materials) all the way from the top pin to the bottom one on the dorsal side of the larvae.</li>
		<li>Pin the larvae fillet, placing 2 additional pins on the left and right sides of the larvae.</li>
		<li>Remove the guts and tracheas using forceps.</li>
		<li>Cut the nerves just outside the ventral nerve cord using spring scissors.</li>
	</ol>
	</li>
</ol>
3. Electrical recordings of EJCs
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56493/56493fig2.jpg" /> 
Figure 2. The recording setup. The sample NMJ pinned to the silicon coated petri dish (arrow) is positioned over the movable stage of the upright compound microscope equipped with epifluorescence capabilities, a high magnification objective, and two micromanipulators. The microscope is stationed on an anti-vibration table. <a href="//cloudfront.jove.com/files/ftp_upload/56493/56493fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Recording
	<ol>
		<li>Place the petri dish with the preparation on the microscope stage (Figure 2). Insert the reference electrode in the bath.</li>
		<li>Fill the recording electrode with HL3 solution. Under the 10x objective (see Table of Materials), immerse the electrode into the bath and place it over muscles 6 and 7 of the abdominal segments 2, 3, or 4 using the micromanipulator (see Table of Materials) (Figure 3 A.1 and A.2).</li>
		<li>Switch the objective to 60x (see Table of Materials). Focus on the area of interest using either epi-fluorescence or DIC optics. Place the tip of the electrode on top of synaptic bouton (Figure 3 B.1-3).</li>
		<li>Press the electrode very gently onto the muscle. Excessive pressure may damage the NMJ or induce an increase in spontaneous synaptic activity. Make sure the tip of the electrode is not clogged- if it is, replace it.</li>
		<li>Switch on the amplifier, A/D board, and the computer.</li>
		<li>Choose the voltage clamp mode on the amplifier.</li>
		<li>Start acquisition software and choose the &#39;gap-free&#39; mode.</li>
		<li>Observe the appearance of mEJCs on the computer screen.</li>
		<li>Ensure that the amplitude of the mEJCs is in the range of 0.2 - 0.7 nA. 
		NOTE: Smaller EJCs indicate that the recording electrode has defects or that it is not positioned properly.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56493/56493fig3.jpg" /> 
Figure 3. Visualization of synaptic boutons. (A) A brightfield image of a hemi-segment under 10x magnification (A.1) and the enlarged boxed area showing muscles 6 and 7 (A.2, the arrow marks the recording electrode). Scale bar = 50 &#181;m. (B) Synaptic boutons are visualized in a Drosophila line with a genetically encoded neuronal marker (CD8-GFP) using epi-fluorescence imaging (B.1) or DIC optics (B.2). Images are taken with the 60x objective and the filter cube for GFP imaging (see Table of Materials). Synaptic boutons are marked with arrows, and an overlay of fluorescent and DIC images is shown in B.3. Scale bar = 10 &#181;m. <a href="//cloudfront.jove.com/files/ftp_upload/56493/56493fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>Stimulation
	<ol>
		<li>Using a micromanipulator, under a visual control, place the stimulation electrode near the axon innervating abdominal segments 2 - 4.</li>
		<li>Apply negative pressure by pulling the piston of the syringe connected to the electrode holder, so that the axon is pulled inside of the electrode (Figure 2).</li>
		<li>Turn on the stimulator. Turn the knob on the isolation unit (see Table of Materials) to set a zero current, and then gently increase it, until EJCs appear (or until the threshold is reached).</li>
		<li>Perform the stimulation in a suprathreshold regime, with the stimulation current increased approximately twice, compared to the threshold for the observation of EJCs. 
		NOTE: In our experience, such stimulation intensity is optimal to avoid action potential failures and action potential firing. For example, if EJCs appear at the stimulation current of 0.2 mA, use a current of 0.4 mA throughout the experiment.</li>
	</ol>
	</li>
	<li>Seal resistance
	<ol>
		<li>Measure the seal resistance of the recording macropatch electrode by turning the electrode resistance switch of the amplifier to &#34;seal test&#34; position. Observe that the value of seal resistance in G&#937; will be displayed in the &#34;current&#34; window.</li>
		<li>Make sure the seal resistance is in the range of 0.5 - 2 M&#8486;. Values out of this range indicate that the electrode is not properly polished or not properly positioned.</li>
		<li>Monitor the seal resistance throughout the recording, making sure it remains constant throughout the experiment.</li>
	</ol>
	</li>
</ol>
4. Analysis
<ol>
	<li>Analyze recordings employing customized software for analysis (see Table of Materials). 
	NOTE: Our lab uses in-house software Quantan31, which is customized for the detection of EJCs and mEJCs recorded focally (Figure 4). This software includes a Gaussian digital filter and allows the detection of quantal peaks in overlapping multi-quantal events (Figure 4A). Other approaches are described in17.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56493/56493fig4.jpg" /> 
Figure 4. Quantal analysis. Detection of mEJCs (A) and EJCs (B) by Quantan software. The event area is marked in green, and peaks are marked by red arrowheads. <a href="//cloudfront.jove.com/files/ftp_upload/56493/56493fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

Drosophila represents an advantageous model organism to study synaptic transmission. Several recording configurations have been used at the larval NMJ, including intracellular recordings of synaptic potentials, recordings of synaptic currents with two electrode voltage clamp33,34, and focal macropatch recordings of synaptic currents described here. The latter technique allows the precise quantification of synaptic transmission at visualized boutons.
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Drosophila is an excellent model system to study the molecular mechanisms controlling synaptic transmission. The neuromuscular system in Drosophila is glutamatergic, and therefore the Drosophila neuromuscular junction (NMJ) can be used to study the conserved features of glutamatergic release. Since Jan and Jan&#39;s study1, the third instar larvae has been broadly used to study evoked and spontaneous synaptic transmission by monitoring excitatory junction potentials (EJPs) or currents (EJCs). EJPs are commonly recorded intracellularly with a sharp glass micro-electrode, and they reflect the activity of the entire NMJ, including all the boutons making synapses at the given muscle fiber.
In contrast, the activity of a limited number of the sites of release can be recorded focally by positioning a micropipette tip near neuronal terminals or synaptic varicosities. This technique was originally employed by Katz and Miledi2, and focal extracellular recordings have been successfully employed at several NMJ preparations, including frog3,4,5, mouse6,7,8, crustacean9,10,11,12,13,14,15,16, and Drosophila17,18,19,20,21,22,23. This approach was further developed by Dudel, who optimized macropatch recoding electrodes24,25. In Dudel&#39;s implementation, this technique closely matched the loose-patch-clamp method26.
The Drosophila larval NMJ has clearly defined synaptic boutons, and transgenic lines with genetically encoded neuronal fluorescent tags (see Table of Materials) are readily available. These advantages enabled us to record EJCs and mEJCs from a selected synaptic bouton20,21,22. Here, we describe this technique in detail.

<table><tbody><tr><td>Sutter P-97</td><td>Sutter instrument</td><td>P-97</td><td>Microelectrode puller</td></tr><tr><td>Narishige MF-830</td><td>Narishige</td><td>MF-830</td><td>Microforge</td></tr><tr><td>WPI MF200</td><td>WPI</td><td>MF200</td><td>Microforge</td></tr><tr><td>Glass capilaries</td><td>WPI</td><td>B150-86-10</td><td>Glass capilaries</td></tr><tr><td>Microtorch 1WG61</td><td>Grainer</td><td>1WG61</td><td>Microtorch</td></tr><tr><td>Sylgard 184 Silicone Elastomer Kit</td><td>Dow Corning</td><td>SYLGARD 184</td><td>Silicone for dissection plates preparation</td></tr><tr><td>Dissection pins</td><td>Amazon</td><td>B00J5PMPJA</td><td>Pins for larvae positioning</td></tr><tr><td>Tweezers</td><td>WPIINC</td><td>500342</td><td>Tweezers for placing pins, removing the guts and tracheas.</td></tr><tr><td>Scissors</td><td>WPIINC</td><td>501778</td><td>Scissors for cutting the cuticula of the larvae and nerves.</td></tr><tr><td>Olympus BX61WI</td><td>Olympus</td><td>BX61WI</td><td>Upright microscope</td></tr><tr><td>Olympus Lumplan FL N 60x</td><td>Olympus</td><td>UPLFLN 60X</td><td>Microscope objective 60X</td></tr><tr><td>Olympus UPlan FL N 10x</td><td>Olympus</td><td>Uplanfl N 10X</td><td>Microscope objective 10X</td></tr><tr><td>Narishige Micromanipulator</td><td>Narishige</td><td>MHW-3</td><td>Three-axis Water Hydraulic Micromanipulator</td></tr><tr><td>npi Electronic GmbH ELC-03XS</td><td>npi Electronic GmbH</td><td>ELC-03XS</td><td>Electrophysiological amplifier</td></tr><tr><td>A.M.P.I Master 8</td><td>A.M.P.I.</td><td>Master 8</td><td>Electrical stimulator</td></tr><tr><td>A.M.P.I Iso-Flex</td><td>A.M.P.I.</td><td>Iso-Flex</td><td>Stimulus isolator</td></tr><tr><td>TMC antivibration table</td><td>TMC</td><td>63-9090</td><td>Antivibration table</td></tr><tr><td>TMC Faraday cage</td><td>TMC</td><td>81-333-90</td><td>Faraday cage</td></tr><tr><td>Digidata 1322A</td><td>Axon Instruments</td><td>Digidata 1322A</td><td>Digidata</td></tr><tr><td>Computer</td><td>Dell</td><td>Dell Dimension 5150</td><td>Computer with Win XP OS</td></tr><tr><td>Electrode holder</td><td>WPI</td><td>MEH3SW</td><td>Electrode holder</td></tr><tr><td>Optical filter</td><td>Omega optical</td><td>XF 115-2</td><td>Filter cube for Green Fluorescent Protein (GFP) detection</td></tr><tr><td>pCLAMP 8</td><td>Axon Instruments</td><td>8.0.0.81</td><td>Software for signal recording</td></tr><tr><td>Quantan</td><td>In-house software</td><td>-</td><td>Software for signal processing</td></tr><tr><td>Canton-S (Wildtype)</td><td>Bloomington Stock Center</td><td>64349</td><td>Control fly line</td></tr><tr><td>cpx SH1</td><td>Generous Gift of J.T. Littleton</td><td>-</td><td>Complexin knock-out fly line with increased spontaneous exocytosis</td></tr><tr><td>CD8-GFP</td><td>Bloomington Stock Center</td><td>5137</td><td>Fly line with neuronal fluorescent (GFP) Tag</td></tr></tbody></table>

focal macropatch recordings of synaptic currents from the drosophila larval neuromuscular junction

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal macropatch recordings of synaptic currents from visualized boutons at the Drosophila larval NMJ. This technique requires customized fabrication of recording micropipettes, as well as a compound microscope equipped with a high magnification, long-distance water immersion objective, differential interference contrast (DIC) optics, and a fluorescent attachment. The recording electrode is positioned on the top of a selected synaptic bouton visualized with DIC optics, epi-fluorescence, or both. The advantage of this technique is that it allows monitoring the synaptic activity of a limited number of sites of release. The recording electrode has&#160;a diameter of several microns, and the release sites positioned outside of the electrode rim do&#160;not significantly affect the recorded currents. The recorded synaptic currents have fast kinetics and can be readily resolved. These advantages are especially important for the studies of mutant fly lines with enhanced spontaneous or asynchronous synaptic activity.

Focal macropatch recordings enable monitoring synaptic activity from selected synaptic boutons (Figure 5). When the electrode is positioned on the top of a synaptic bouton (Figure 5A, site 1), the recorded mEJCs (Figure 5C, site 1) have a amplitudes significantly exceeding the noise level and sharp rising phases (at a sub-millisecond range). When the recording electrode is moved away from the synapti...

Watch this Scientific Journal Video about Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction at JoVE.com

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Synaptic currents can be recorded focally from visualized synaptic boutons at the Drosophila third instar larvae neuromuscular junction. This technique enables monitoring the activity of a single synaptic bouton.

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal ...

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Universidad de Cartagena UDEC

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Neuroscience

6.0K Views.  Wayne State University. Synaptic currents can be recorded focally from visualized synaptic boutons at the Drosophila third instar larvae neuromuscular junction. This technique enables monitoring the activity of a single synaptic bouton.

Video: Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Paired Patch Clamp Recordings from Motor-neuron and Target Skeletal Muscle in Zebrafish

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle. This is a direct consequence of the accessibility to both cell types and ability to visually distinguish the single segmental CaP motor-neuron on the basis of morphology and location. This video demonstrates the microscopic methods used to identify a CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type. Identification of the CaP motor-neuron type is confirmed by either dye filling or by the biophysical features such as action potential waveform and cell input resistance. Motor-neuron recordings routinely last for one hour permitting long-term recordings from multiple different target muscle cells. Control over the motor-neuron firing pattern enables measurements of the frequency-dependence of synaptic transmission at the neuromuscular junction. Owing to a large quantal size and the low noise provided by whole cell voltage clamp, all of the unitary events can be resolved in muscle. This feature permits study of basic synaptic properties such as release properties, vesicle recycling, as well as synaptic depression and facilitation. The advantages offered by this in vivo preparation eclipse previous neuromuscular model systems studied wherein the motor-neurons are usually stimulated by extracellular electrodes and the muscles are too large for whole cell patch clamp. The zebrafish preparation is amenable to combining electrophysiological analysis with a wide range of approaches including transgenic lines, morpholino knockdown, pharmacological intervention and in vivo imaging. These approaches, coupled with the growing number of neuromuscular disease models provided by mutant lines of zebrafish, open the door for new understanding of human neuromuscular disorders.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target skeletal muscle. This video demonstrates the microscopic methods used to identify a segmental CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle.  ...

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The Drosophila larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila is well known for the ease of powerful genetic manipulations and the larval nervous system has proven particularly useful in studying not only normal function but also perturbations that accompany some neurological disease (Lloyd and Taylor, 2010). Many key synaptic molecules found in Drosophila are also found in mammals and like most CNS excitatory synapses in mammals, the Drosophila NMJ is glutamatergic and demonstrates activity-dependent remodeling (Kohet al. , 2000). Additionally, Drosophila neurons can be individually identified because their innervation patterns are stereotyped and repetitive making it possible to study identified synaptic terminals, such as those between motor neurons and the body-wall muscle fibers that they innervate (Keshishian and Kim, 2004). The existence of evolutionarily conserved synapse components along with the ease of genetic and physical manipulation make the Drosophila model ideal for investigating the mechanisms underlying synaptic function (Budnik, 1996).
The active zones at synaptic terminals are of particular interest because these are the sites of neurotransmitter release. NC82 is a monoclonal antibody that recognizes the Drosophila protein Bruchpilot (Brp), a CAST1/ERC family member that is an important component of the active zone (Waghet al. , 2006). Brp was shown to directly shape the active zone T-bar and is responsible for effectively clustering Ca2+ channels beneath the T-bar density (Fouquetet al. , 2009). Mutants of Brp have reduced Ca2+ channel density, depressed evoked vesicle release, and altered short-term plasticity (Kittelet al. , 2006). Alterations to active zones have been observed in Drosophila disease models. For example, immunofluorescence using the NC82 antibody showed that the active zone density was decreased in models of amyotrophic lateral sclerosis and Pitt-Hopkins syndrome (Ratnaparkhiet al. , 2008; Zweieret al. , 2009). Thus, evaluation of active zones, or other synaptic proteins, in Drosophila larvae models of disease may provide a valuable initial clue to the presence of a synaptic defect.
Preparing whole-mount dissected Drosophila larvae for immunofluorescence analysis of the NMJ requires some skill, but can be accomplished by most scientists with a little practice. Presented is a method that provides for multiple larvae to be dissected and immunostained in the same dissection dish, limiting environmental differences between each genotype and providing sufficient animals for confidence in reproducibility and statistical analysis.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The neuromuscular junction (NMJ) of Drosophila melanogaster is an important model system for studying normal synaptic function as well as perturbations to synaptic function found in certain neurological diseases. We present a protocol for dissection of the Drosophila larval motor system and immunostaining for active zone proteins within the NMJ.

The Drosophila  larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila  is well known for ...

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. ...

Biology

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

The Drosophila larval neuromuscular preparation has proven to be a useful tool for studying synaptic physiology1,2,3. Currently, the only means available to evoke excitatory junctional potentials (EJPs) in this preparation involves the use of suction electrodes. In both research and teaching labs, students often have difficulty maneuvering and manipulating this type of stimulating electrode. In the present work, we show how to remotely stimulate synaptic potentials at the larval NMJ without the use of suction electrodes. By expressing channelrhodopsin2 (ChR2) 4,5,6 in Drosophila motor neurons using the GAL4-UAS system 7, and making minor changes to a basic electrophysiology rig, we were able to reliably evoke EJPs with pulses of blue light. This technique could be of particular use in neurophysiology teaching labs where student rig practice time and resources are limited.

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

This procedure uses a blue light-activated algal channel and cell-specific genetic expression tools to evoke synaptic potentials with light pulses at the neuromuscular junction (NMJ) in Drosophila larvae. This technique is an inexpensive and easy-to-use alternative to suction electrode stimulation for synaptic physiology studies in research and teaching laboratories.

The Drosophila larval neuromuscular preparation has proven to be a useful tool for studying synaptic physiology1,2,3. Currently, the only means available ...

Dissecting and Recording from The C. Elegans Neuromuscular Junction

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale. This complex process requires the coordinated activity of many pre- and post-synaptic proteins to ensure appropriate synaptic connectivity, conduction of electrical signals, targeting and priming of secretory vesicles, calcium sensing, vesicle fusion, localization and function of postsynaptic receptors and finally, recycling mechanisms. As neuroscientists it is our goal to elucidate which proteins function in each of these steps and understand their mechanisms of action. Electrophysiological recordings from synapses provide a quantifiable read out of the underlying electrical events that occur during synaptic transmission. By combining this technique with the powerful array of molecular and genetic tools available to manipulate synaptic proteins in C. elegans, we can analyze the resulting functional changes in synaptic transmission. 
The C. elegans NMJs formed between motor neurons and body wall muscles control locomotion, therefore, mutants with uncoordinated locomotory phenotypes (known as unc s) often perturb synaptic transmission at these synapses 1. Since unc mutants are maintained on a rich supply of a bacterial food source, they remain viable as long as they retain some pharyngeal pumping ability to ingest food. This, together with the fact that C. elegans exist as hermaphrodites, allows them to pass on mutant progeny without the need for elaborate mating behaviors. These attributes, coupled with our recent ability to record from the worms NMJs 2,3,7 make this an excellent model organism in which to address precisely how unc mutants impact neurotransmission. 
The dissection method involves immobilizing adult worms using a cyanoacrylic glue in order to make an incision in the worm cuticle exposing the NMJs. Since C. elegans adults are only 1 mm in length the dissection is performed with the use of a dissecting microscope and requires excellent hand-eye coordination. NMJ recordings are made by whole-cell voltage clamping individual body wall muscle cells and neurotransmitter release can be evoked using a variety of stimulation protocols including electrical stimulation, light-activated channel-rhodopsin-mediated depolarization 4 and hyperosmotic saline, all of which will be briefly described.

Application of electrophysiology to accessible synapses provides a quantifiable measure of synaptic activity, useful in analyzing synaptic mutants. This article describes a dissection method used to expose the neuromuscular junctions (NMJ) of Caenorhabditis elegans (C. elegans) and briefly discusses some of the uses to which this preparation can be applied.

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale.   This ...

Preparation of Drosophila Central Neurons for in situ Patch Clamping

Short generation times and facile genetic techniques make the fruit fly Drosophila melanogaster an excellent genetic model in fundamental neuroscience research. Ion channels are the basis of all behavior since they mediate neuronal excitability. The first voltage gated ion channel cloned was the Drosophila voltage gated potassium channel Shaker1,2. Toward understanding the role of ion channels and membrane excitability for nervous system function it is useful to combine powerful genetic tools available in Drosophila with in situ patch clamp recordings. For many years such recordings have been hampered by the small size of the Drosophila CNS. Furthermore, a robust sheath made of glia and collagen constituted obstacles for patch pipette access to central neurons. Removal of this sheath is a necessary precondition for patch clamp recordings from any neuron in the adult Drosophila CNS. In recent years scientists have been able to conduct in situ patch clamp recordings from neurons in the adult brain3,4 and ventral nerve cord of embryonic5,6, larval7,8,9,10, and adult Drosophila11,12,13,14. A stable giga-seal is the main precondition for a good patch and depends on clean contact of the patch pipette with the cell membrane to avoid leak currents. Therefore, for whole cell in situ patch clamp recordings from adult Drosophila neurons must be cleaned thoroughly. In the first step, the ganglionic sheath has to be treated enzymatically and mechanically removed to make the target cells accessible. In the second step, the cell membrane has to be polished so that no layer of glia, collagen or other material may disturb giga-seal formation. This article describes how to prepare an identified central neuron in the Drosophila ventral nerve cord, the flight motoneuron 5 (MN515), for somatic whole cell patch clamp recordings. Identification and visibility of the neuron is achieved by targeted expression of GFP in MN5. We do not aim to explain the patch clamp technique itself.

Preparation of Drosophila Central Neurons for in situ Patch Clamping

In situ patch clamp recordings are used for electrophysiological characterization of neurons in intact circuitry. In the Drosophila genetic model patch clamping is difficult because the CNS is small and surrounded by a robust sheath. This article describes the procedure to remove the sheath and clean neurons for subsequent patch clamp recordings.

Short generation times and facile genetic techniques make the fruit fly Drosophila melanogaster an excellent genetic model in fundamental neuroscience ...

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 ...

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Mauthner cells (M-cells) are large reticulospinal neurons located in the hindbrain of teleost fish. They are key neurons involved in a characteristic behavior known as the C-start or escape response that occurs when the organism perceives a threat. The M-cell has been extensively studied in adult goldfish where it has been shown to receive a wide range of excitatory, inhibitory and neuromodulatory signals1. We have been examining M-cell activity in embryonic zebrafish in order to study aspects of synaptic development in a vertebrate preparation. In the late 1990s Ali and colleagues developed a preparation for patch clamp recording from M-cells in zebrafish embryos, in which the CNS was largely intact2,3,4. The objective at that time was to record synaptic activity from hindbrain neurons, spinal cord neurons and trunk skeletal muscle while maintaining functional synaptic connections within an intact brain-spinal cord preparation. This preparation is still used in our laboratory today. To examine the mechanisms underlying developmental synaptic plasticity, we record excitatory (AMPA and NMDA-mediated)5,6 and inhibitory (GABA and glycine) synaptic currents from developing M-cells. Importantly, this unique preparation allows us to return to the same cell (M-cell) from preparation to preparation to carefully examine synaptic plasticity and neuro-development in an embryonic organism. The benefits provided by this preparation include 1) intact, functional synaptic connections onto the M-cell, 2) relatively inexpensive preparations, 3) a large supply of readily available embryos 4) the ability to return to the same cell type (i.e. M-cell) in every preparation, so that synaptic development at the level of an individual cell can be examined from fish to fish, and 5) imaging of whole preparations due to the transparent nature of the embryos.

We have developed an intact brain-spinal cord preparation to record and monitor electrical activity via patch clamp recording from the Mauthner neurons and other reticulospinal cells in zebrafish embryos. Thus, we are able to record excitatory and inhibitory synaptic currents, voltage-gated channel activity and action potentials from key neurons in a developing embryo.

Mauthner cells (M-cells) are large  reticulospinal neurons located in the hindbrain of teleost fish. They are key  neurons involved in a characteristic ...

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at the Drosophila NMJ enables researchers to acquire synaptic information, encompassing both quantitative data on synapse abundance and detailed insights into their morphology. However, the diffuse distribution and limited visual range of the TEM present challenges for the ultrastructural analysis. This study introduces an innovative and efficient sample preparation method that surpasses the conventional approach. The procedure begins by placing a metal mesh at the base of a flat-bottomed bottle or test tube, followed by positioning fixed larvae samples onto the mesh. An additional mesh is placed over the samples, ensuring that they are positioned between the two meshes. The fixed samples are thoroughly dehydrated and infiltrated before proceeding with the embedding procedure. Then embedding of the samples in epoxy resin is performed in a flat sheet manner, which allows for the preparation of muscles for positioning and sectioning. Benefiting from these steps, all the muscles of Drosophila larvae can be visualized under light microscopy, therey facilitating subsequent positioning and sectioning. Excess resin is removed after locating the 6th and 7th muscles of body segments A2 and A3. Serial ultra-thin sectioning of the 6th or 7th muscle is performed.

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

This report provides a new sample preparation procedure for visualizing neuromuscular junctions in Drosophila Larvae. This method is more effective in preventing the curling of the samples compared to the traditional method and is particularly useful for Drosophila neuromuscular junction ultrastructural analysis.

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at ...

Drosophila Neuromuscular Junction (NMJ) Quantification: A Method to Assess Synaptic Morphology and Function

Source: Castells-Nobau, A., et al. <a href="https://jove.unicartagenaproxy.elogim.com/video/55395/" target="_blank">Two Algorithms for High-throughput and Multi-parametric Quantification of Drosophila Neuromuscular Junction Morphology</a>. J. Vis. Exp. (2017).
The Drosophila neuromuscular junction (NMJ) is used as a model to study the morphology and function of synapses. This video describes important features that researchers quantify to characterize the NMJ. The example protocol demonstrates a software able to perform the quantification automatically.

Drosophila Kwantyfikacja połączenia nerwowo-mięśniowego (NMJ): metoda oceny morfologii i funkcji synaps

Source: Castells-Nobau, A., et al. Two Algorithms for High-throughput and Multi-parametric Quantification of Drosophila Neuromuscular Junction Morphology. ...

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

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Paired Patch Clamp Recordings from Motor-neuron and Target Skeletal Muscle in Zebrafish

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

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

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

Dissecting and Recording from The C. Elegans Neuromuscular Junction

Preparation of Drosophila Central Neurons for in situ Patch Clamping

Electrophysiological Recording in the Drosophila Embryo

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

Drosophila Kwantyfikacja połączenia nerwowo-mięśniowego (NMJ): metoda oceny morfologii i funkcji synaps