We would like to thank Joshua Singer, Jonathan Schenk, as well as thoughtful reviewers for comments on the manuscript. We would also like to thank Ben White and Harold Zakon for discussions on the technique. Jonathan Schenk provided the data for Figure 3A. This work was supported by a Whitehall Foundation Grant and an NIH R21 to QG.

1. Prepare Flies and Identify GAL4 promoter Lines
<ol>
	<li>Select a GAL4 promoter line and cross it with flies carrying both the UAS-NaChBac transgene and an optogenetic transgene for activation. 
	NOTE: UAS-CsChrimson11 is selected due to its high conductance and the ease with which it activates NaChBac-mediated action potentials. Both UAS-NaChBac and UAS-CsChrimson are available from the Bloomington Stocks Repository.</li>
	<li>Prepare all-trans retinal as a stock solution in ethanol (35 mM) and store the solution in freezer at -20 &#176;C.</li>
	<li>Mix 28 &#181;L of this stock into approximately 5 mL of rehydrated potato flake food and add this mixture to the top of a vial of conventional Drosophila medium.</li>
	<li>Raise adult flies expressing csChrimson on food containing 0.2 mM all-trans-retinal for 1&#8211;3 days11.</li>
</ol>
2. Prepare TTX Stock
<ol>
	<li>Create a stock solution of 1 mM TTX in distilled or deionized water. Store this solution in a lab refrigerator for several months. Check the institutions policies regarding the storage of TTX as many institutions classify TTX as a hazardous or controlled substance.</li>
</ol>
3. Prepare Flies for Electrophysiology
<ol>
	<li>Collect 1&#8211;3 day old adult flies that are raised on the food with all-trans-retinal. 
	NOTE: Refer to Murthy and Turner in 2013 for a detailed description of patch clamping Drosophila neurons12,13.</li>
	<li>Put the flies in a glass scintillation vial. Place the vial onto ice for 1 min to immobilize the flies.</li>
	<li>Use coarse forceps to insert a fly into a custom-made recording chamber. The chamber consists of a 3 &#189;&#34; circle laser cut from 1/16&#34; acrylic. Cut a &#190;&#34; hole is into the center where a custom foil is attached with 2-part epoxy. The foil contains a triangular shaped hope that the fly is inserted into.</li>
	<li>Apply wax or UV curable epoxy around the animal to firmly secure it within the recording chamber.</li>
	<li>Illuminate the preparation with a gooseneck optical fiber for visualization under a stereomicroscope. Set the gooseneck fibers for oblique illumination by adjusting them so that they illuminate the fly directly from the side.</li>
	<li>Adjust the light level to lowest possible setting that still allows clear visualization of the fly. This light intensity will vary across individual experiments. 
	NOTE: This protocol can be adapted by using an in situ brain explant method14.</li>
	<li>Retrieve external recording saline. The saline contains in mM: 103 NaCl, 3 KCl, 5 N-tris(hydroxymethyl)methyl-2- aminoethane-sulfonic acid, 8 trehalose, 10 glucose, 26 NaHCO3, 1 NaH2PO4, 1.5 CaCl2, and 4 MgCl2 (adjusted to 270&#8211;275 m&#937;). The saline is bubbled with 95% O2/5% CO2 (carbogen) and has a pH adjusted to 7.315.</li>
	<li>Place 4-6 drops of extracellular recording saline over the fly preparation. At this point, increase the light level at the dissection microscope for better visibility.</li>
	<li>Sharpen a tungsten wire using electrolysis. To make the wire, prepare a saturated solution of KNO3 and apply approximately 20 V of AC current while dipping the tip of the wire into the solution 20 times for 100 ms each until the tip is sharpened. Use the sharpened tungsten wire to remove the cuticle on the head of the fly and thus exposing the brain.</li>
	<li>Further expose the brain by removing trachea and fat bodies that surround it. If accessing the antennal lobes, use a bent tungsten wire or similar tool to gently tuck antennae underneath the foil recording chamber to increase visibility. Use sharp forceps to gently tear away the glial sheathing covering the area of interest where recordings will be made.</li>
	<li>Transfer the recording chamber to the electrophysiology rig. Immediately start perfusion with external recording saline bubbled with carbogen to preserve the brain. The saline solution reservoir is placed above the microscope stage and is gravity-fed to the preparation. Use a vacuum line and a 2 L Erlenmeyer flask to collect the waste saline.</li>
</ol>
4. Obtain Whole-cell Recording
<ol>
	<li>Pull patch pipettes on a commercial pipette puller. Consult the manual for settings to achieve an appropriate pipette with a tip resistance near 7 m&#937;.</li>
	<li>Add 4 &#181;L of internal recording solution into a patch pipette. The internal saline consists of 140 potassium aspartate, 10 HEPES, 4 MgATP, 0.5 Na3GTP, 1 EGTA, and 1 KCl in mM.</li>
	<li>Set up patch clamp amplifier for whole-cell recording. Zero the amplifier&#39;s offset and set the amplifier to deliver test pulses. Here, an AM systems Patch clamp amplifier 2,400 is used.</li>
	<li>Apply positive pressure through the pipette and approach the cell. Use a micromanipulator to direct the pipette and contact the cell. Release positive pressure. Set the holding potential of the membrane to -60 mV. The pipette should form a gigaohm seal with the cell&#39;s membrane as evidenced by a low sub-picoamp holding current. 
	NOTE: If using GFP guided cell targeting, try to minimize use of epifluorescent light to prevent ChR2 activation and potential synaptic depression. Also wait for a few minutes before applying step 5.</li>
	<li>Set the amplifier to deliver test pulses from -50 mV to -60 mV. This setting is standard on patch-clamp amplifiers. Test pulses will reveal a large capacitative transient representing the current necessary to charge the patch pipette. Remove capacitative transients by adjusting the capacity compensation knobs on the AM 2400 or similar amplifier.</li>
	<li>Apply brief pulses of negative pressure to rupture the cell membrane and obtain whole-cell configuration. 
	NOTE: A whole cell configuration is observed when there is a large increase in the capacitative transients showing the capacitance of the neuron. A small increase in the holding current (baseline) will likely also be observed. Set the amplifier to current clamp mode and adjust the cell&#39;s potential to -50 to -60 mV.</li>
</ol>
5. Test Synaptic Connectivity
<ol>
	<li>Configure the data acquisition (DAQ) system to trigger an LED driver to generate light pulses. Here, a National Instruments DAQ system is used with Matlab to deliver an analog signal to a LED driver. The LED intensity is adjusted via the analog signal to the driver. The LED is a 620-630 nm wavelength LED.</li>
	<li>Position the high-powered red LED directly underneath the preparation. Adjust the light intensity so that 0.238 mW/mm2 reaches the fly.</li>
	<li>Activate presynaptic neurons while recording from the postsynaptic cell of interest. Achieve cell activation optogenetically, pharmacogentically, or via nerve stimulation. Here, csChrimson and brief pulses of red light are applied. 
	NOTE: Synaptic connections can be monitored either in current clamp as EPSPs or voltage clamp as EPSCs. A synaptic connection should be visible before applying TTX in response to a 40 ms light pulse. If no connection is observed, it is unlikely that the neurons are synaptically connected either mono- or polysynaptically. The holding potential may be adjusted to emphasize excitatory or inhibitory connections accordingly.</li>
	<li>If a connection is observed in normal saline, add TTX to the perfusion system to achieve a final concentration of 1 &#181;M. Use a recirculating pump to capture and reuse the saline to conserve TTX. The peristaltic pump will draw the saline solution from the bath and return it to the saline reservoir.</li>
	<li>Adjust the pump&#39;s speed to provide a strong constant vacuum that does not allow that saline level in the bath to rise and fall. 
	NOTE: The application of TTX should result in the cessation of spiking activity in the neuron that is being monitored in whole-cell. This confirms that enough TTX has been added to the saline.</li>
	<li>Apply brief pulses of red light (40 ms for each pulse) again. At this point, any synaptic connection that remains is likely monosynaptic. Add pharmacological antagonists to the recirculating saline to test the chemical nature of the synaptic connections that remain in TTX. 
	Note: When combining optogenetics and fluorescent targeting of neurons, it may be important not to persistently activate the presynaptic population of neurons while trying to obtain a recording. This can lead to synaptic depression and makes it difficult to see connections. Because csChrimson has a long excitation band that extends broadly into the range of green fluorescent protein, a red fluorophore can be used to visualize neurons and channel2rhodopsin (Ch2R) can be used to excite cell populations. Doing so requires specific genetics and a custom filter cube. To generate suitable flies, make flies that express the red fluorophore (RFP or mCherry) under the LexA or Q system binary expression systems. These will need to be crossed with flies expressing a Gal4 promoter and UAS-Ch2R and UAS-NaChBac effector genes. The optimal filter cube for epifluorescence will have a narrow excitation range to excite the red fluorophore, but not excite the Ch2R. Additionally, the filter set should include a long pass emission filter to collect as much emitted light. We used a custom filter set from Chroma that included part numbers et580/25x and t600lpxr (from the 49306 set), but with an et610lp barrier/emission filter.</li>
</ol>

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The authors have nothing to disclose.

TERPS analysis compliments current techniques used for circuit cracking by enabling the detection of synapses between identified neurons. Specifically, the approach reveals monosynaptic connections by broadly silencing action potentials with TTX while restoring excitability in a select population of neurons with the TTX-insensitive sodium channel NaChBac. Synaptic release is elicited by optogenetic stimulation while postsynaptic events are monitored with whole-cell recordings. TERPS distinguishes itself from other approa...

A major goal of neuroscience is to map the connections between neurons to understand how information flows through circuits. Numerous approaches and resources have become available to test for functional connectivity between neurons in a range of model systems1,2. To generate the most accurate wiring diagrams using electrophysiology, it is important to resolve whether observed connections between two cells are direct and monosynaptic versus indirect and polysynaptic. A gold standard for making this distinction in mammalian neurons is to measure the latency between single action potentials in the presynaptic cells and the onset of excitatory postsynaptic currents (EPSCs) in the second neuron. Monosynaptic connections should have short latencies of a few milliseconds and low variability3. This approach can be complicated in invertebrate neurons because synapses may occur in their dendrites far from the somatic recording site, causing delays in detection due to long electrotonic distances between postsynaptic conductances and the recording electrode. Such delays can introduce ambiguity regarding poly- versus monosynaptic contributions. Additionally, small synaptic events may decay before reaching the somatic recordings site, and driving stronger presynaptic activity, is likely to recruit polysynaptic events.
Various techniques have been developed to test for monosynaptic connections in invertebrates. One approach uses high divalent cation solutions (Hi-Di) containing excess Mg++ and Ca++. This solution blocks polysynaptic connections by reducing release probability and increasing action potential threshold to favor detection of monosynaptic input4,5. Determining the precise ratio of Mg++ to Ca++ required, however, is not trivial and polysynaptic contributions may persist for even modest stimulation6. An alternative approach called GFP Reconstitution Across Synaptic Partners (GRASP) takes advantage of the proximity between pre- and postsynaptic membranes found at synapses to infer a monosynaptic connection 7. Here, a component of green fluorescent protein (GFP) is expressed in one neuron, and the complementary fragment of the molecule is expressed in a putative postsynaptic partner. The presence of fluorescence indicates that the two neurons are in close enough proximity to permit reconstitution of the GFP molecule and implies the existence of a synapse. GRASP can report synapses falsely, though, if two cells have closely apposed membranes, as in a nerve or fascicle. Variants of GRASP eliminate such false positives by tethering the presynaptic fragment of GFP directly to synaptobrevin, thus allowing reconstitution only at active synapses8. While GRASP and its variants have been instrumental in determining functional connectivity in the Drosophila CNS, some connections may not be made visible by GRASP if the distances between pre- and postsynaptic partners is relatively large. This is particularly pertinent in the assessment of volume transmission associated with neuromodulation9 or GABAergic inhibition10.
Here, a complementary and novel technique is demonstrated for testing direct monosynaptic connections in the Drosophila CNS. This method, called Tetrodotoxin Engineered Resistance for Probing Synapses (TERPS), works by co-expression of the tetrodotoxin (TTX)-resistant sodium channel, NaChBac, and an optogenetic activator in a presynaptic cell while recording from putative postsynaptic partners9. In the presence of TTX, all action potential-mediated activity is suppressed in cells other than those expressing NaChBac. The NaChBac channel selectively restores excitability in the targeted presynaptic cell(s), permitting light-evoked synaptic transmission. This method permits strong activation of the presynaptic neurons, such that connections can be resolved postsynaptically by somatic recording while reducing the probability of recruiting polysynaptic circuits. Importantly, this technique permits the study of volume transmission and reveals the spread of transmitter released from a single neuron throughout a circuit. In addition, TERPS can reveal the chemical nature of the connection through conventional pharmacology. TERPS is suitable for use in any model system that allows the transgenic expression of TTX-insensitive sodium channels.

<table><tbody><tr><td>UAS-csChrimson</td><td>Bloomington Drosophila Stock Center</td><td>55135</td><td>Used as a neural activator</td></tr><tr><td>UAS-NaChBac</td><td>Bloomington Drosophila Stock Center</td><td>9466</td><td>Resotores excitibility in cells in TTX</td></tr><tr><td>Tetrodotoxin</td><td>Tochris</td><td>1078</td><td>Special permission may be needed to purchase TTX as it is a controlled substance</td></tr><tr><td>all trans-Retinal</td><td>Sigma-Aldrichall trans-Retinal</td><td>R2500</td><td>Require co-factor for channelrhodopsin</td></tr><tr><td>Weldable 321 Stainless Steel&amp;nbsp;Sheet,&amp;nbsp;0.002&quot;&amp;nbsp;Thick,&amp;nbsp;10&quot;&amp;nbsp;Wide</td><td>McMaster Carr</td><td>3254K7</td><td>Used to make custom fly holder. Custom foil can be laser cut at pololu.com from the provided PDF file</td></tr><tr><td>Dissection Microscope</td><td>Zeiss</td><td>Stemi 2000-C</td><td>Used for dissection of preparation</td></tr><tr><td>Waxer</td><td>Almore</td><td>Eectra Waxer 66000</td><td>Used during dissection to secure fly in foil</td></tr><tr><td>Paraffin Wax</td><td>Joann</td><td>4917217</td><td>Used with waxer</td></tr><tr><td>Number 5 forceps</td><td>Fine Science Tools</td><td>#5CO</td><td>Used for dissection and desheathing</td></tr><tr><td>Dissection Scissors</td><td>Fine Science Tools</td><td>15001-08</td><td>Used to remove parts of the cuticle during dissection</td></tr><tr><td>Tungsten wire</td><td>A-M Systems</td><td>797500</td><td>Use with electrolysis to make sharpened needles for dissection</td></tr><tr><td>Reciculating Peristaltic Pump</td><td>Simply Pumps (Amazon)</td><td>PM200S</td><td>for recirculating TTX</td></tr><tr><td>Speed Controller for peristaltic pump</td><td>Zitrades (Amazon)</td><td>N/A</td><td>PWM Dimming Controller For LED Lights or Ribbon, 12 Volt 8 Amp,Adjustable Brightness Light Switch Dimmer Controller DC12V 8A 96W for Led Strip Light B</td></tr><tr><td>Versa-Mount&amp;nbsp;Precision Compressed Air Regulator</td><td>McMaster Carr</td><td>1804T1</td><td>For applying positive pressure during patching</td></tr><tr><td>Glass capilaries</td><td>World Precision Instruments</td><td>TW150F-3</td><td>For patch pipettes</td></tr><tr><td>Multipurpose Gauge</td><td>McMaster Carr</td><td>3846K431</td><td>Gauge for pressure regulator</td></tr><tr><td>Electrophysiology Camera</td><td>Dage MTI&amp;nbsp;</td><td>IR-1000</td><td>Any camera that works in the IR range (850 nm) will work. You do not want to use red illumination as this can activate csChrimson</td></tr><tr><td>IR LED</td><td>Thorlabs&amp;nbsp;</td><td>M850F2</td><td>For oblique illumination</td></tr><tr><td>fiber optic for IR LED</td><td>Thorlabs</td><td>M89L01</td><td>Couples to IR LED</td></tr><tr><td>Objective lens&amp;nbsp;</td><td>Olympus 40X</td><td>LUMPlanFLN</td><td>This can be used on most microscipes and works well for visualizing fly neurons.</td></tr><tr><td>Amber LED</td><td>Thorlabs</td><td>M590L3d</td><td>For visualizing RFP and mCherry</td></tr><tr><td>Blue LED</td><td>Thorlabs</td><td>&lt;strong&gt;M470L3&lt;/strong&gt;&lt;sup&gt;d&lt;/sup&gt;</td><td>For visualizing GFP</td></tr><tr><td>GFP filter set</td><td>Chroma</td><td>&lt;strong&gt;49011&lt;/strong&gt;</td><td>For visualizing GFP or stimulating channel2rhodopsin</td></tr><tr><td>Custom mCherry Filter Set</td><td>Chroma</td><td>&amp;nbsp;et580/25x and t600lpxr (from the 49306 set) but with an et610lp barrier/emission optic</td><td>Use only if you wish to patch identified neurons with channel2rhodopsin</td></tr><tr><td>Dichroic to combine Amber and blue LED</td><td>Thorlabs</td><td>DMLP550R</td><td>Use only patch under mCherry and excite channel2rhodopsin with blue light.</td></tr><tr><td>Red LED</td><td>LEDSupply</td><td>Cree XPE 620 - 630 nm</td><td>Used to drive csChrimson</td></tr><tr><td>LED Driver</td><td>LEDSupply</td><td>3021-D-E-1000</td><td>Used to drive LEDs for optogenetic stimulation</td></tr><tr><td>Manipulator</td><td>Sutter Instruments</td><td>MP-225</td><td>Used to position pipette during recordings</td></tr><tr><td>Patchclamp Amplifier</td><td>A-M Systems</td><td>Model 2400</td><td>An equivalent amplifier is suitable</td></tr><tr><td>Bessel Filter</td><td>Warner Instruments&amp;nbsp;</td><td>LPF 202A</td><td>Auxillary filter used to filter current trace to oscilloscope during patching.</td></tr><tr><td>Data Acquisition System&amp;nbsp;</td><td>National Instruments</td><td>NI PCIe-6321&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; 781044-01</td><td>Used to record data from amplifier to computer</td></tr><tr><td>Connector Block - BNC Terminal BNC-2090A</td><td>National Instruments</td><td>779556-01</td><td>Used to connect amplifier to DAQ card.</td></tr><tr><td>Steel Foil&amp;nbsp;</td><td>McMaster Carr</td><td>3254K7</td><td>Steel foil for custom recording chamber</td></tr><tr><td>Magnets</td><td>K&amp;amp;J Magnetics</td><td>D42</td><td>To secure recording chamber to ring stand</td></tr><tr><td>1/16&quot; Cell Cast Acrylic Clear</td><td>Pololu</td><td></td><td>Used to make custom recording chamber. Acrylic can be laser cut at pololu.com from the provided PDF file</td></tr></tbody></table>

examining monosynaptic connections in drosophila using tetrodotoxin resistant sodium channels

Here, a new technique termed Tetrotoxin (TTX) Engineered Resistance for Probing Synapses (TERPS) is applied to test for monosynaptic connections between target neurons. The method relies on co-expression of a transgenic activator with the tetrodotoxin-resistant sodium channel, NaChBac, in a specific presynaptic neuron. Connections with putative post-synaptic partners are determined by whole-cell recordings in the presence of TTX, which blocks electrical activity in neurons that do not express NaChBac. This approach can be modified to work with any activator or calcium imaging as a reporter of connections. TERPS adds to the growing set of tools available for determining connectivity within networks. However, TERPS is unique in that it also reliably reports bulk or volume transmission and spillover transmission.

TERPS is used to distinguish between mono- and polysynaptic contributions in synaptic connections between neurons. While weak stimulation of a cell may be used to test direct connections, driving greater presynaptic activity often recruits polysynaptic connections (Figure 1A). TERPS works by co-expressing the TTX-insensitive sodium channel NaChBac and an optogenetic activator, and testing connections in the presence of TTX to eliminate polysynaptic connection...

Watch this Scientific Journal Video about Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels at JoVE.com

Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels

This article features a method to test the monosynaptic connections between neurons by employing tetrodotoxin and the tetrodotoxin-resistant sodium channel, NaChBac.

Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels

Here, a new technique termed Tetrotoxin (TTX) Engineered Resistance for Probing Synapses (TERPS) is applied to test for monosynaptic connections between ...

examining-monosynaptic-connections-drosophila-using-tetrodotoxin

Research

JoVE Journal

Neuroscience

9.9K Views.  University of Maryland. This article features a method to test the monosynaptic connections between neurons by employing tetrodotoxin and the tetrodotoxin-resistant sodium channel, NaChBac.

Video: Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels

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 ...

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

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. 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.

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

This technique exposes the Drosophila embryonic neuromusculature for immunohistochemistry or electrophysiological recording. It is useful for studying early events in neuromuscular development or performing electrophysiology in mutants that cannot hatch.

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

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.

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 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.

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

Viewing Microtubule Networks in Drosophila Neuromuscular Junctions and Muscle Cells

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network

The microtubule network is an essential component of the nervous system. Mutations in many microtubules regulatory proteins are associated with neurodevelopmental disorders and neurological diseases, such as microtubule-associated protein Tau to neurodegenerative diseases, microtubule severing protein Spastin and Katanin 60 cause hereditary spastic paraplegia and neurodevelopmental abnormalities, respectively. Detection of microtubule networks in neurons is advantageous for elucidating the pathogenesis of neurological disorders. However, the small size of neurons and the dense arrangement of axonal microtubule bundles make visualizing the microtubule networks challenging. In this study, we describe a method for dissection of the larval neuromuscular junction and muscle cells, as well as immunostaining of &#945;-tubulin and microtubule-associated protein Futsch to visualize microtubule networks in Drosophila melanogaster. The neuromuscular junction permits us to observe both pre-and post-synaptic microtubules, and the large size of muscle cells in Drosophila larva allows for clear visualization of the microtubule network. Here, by mutating and overexpressing Katanin 60 in Drosophila melanogaster,&#160;and then examining the microtubule networks in the neuromuscular junction and muscle cells, we&#160;accurately reveal the regulatory role of Katanin 60 in neurodevelopment. Therefore, combined with the powerful genetic tools of Drosophila&#160;melanogaster, this protocol greatly facilitates genetic screening and microtubule dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

Author Spotlight: Understanding Microtubule Network in Drosophila Neuromuscular Junctions 

Here, we present a detailed protocol to visualize the microtubule networks in neuromuscular junctions and muscle cells. Combined with the powerful genetic tools of Drosophila melanogaster, this protocol greatly facilitates genetic screening and microtubule-dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

The microtubule network is an essential component of the nervous system. Mutations in many microtubules regulatory proteins are associated with ...

Single-Neuron Ablation in Drosophila for Studying Nervous System Adaptation

Laser Cell Ablation in Intact Drosophila Larvae Reveals Synaptic Competition

The protocol describes single-neuron ablation with a 2-photon laser system in the central nervous system (CNS) of intact Drosophila melanogaster larvae. Using this non-invasive method, the developing nervous system can be manipulated in a cell-specific manner. Disrupting the development of individual neurons in a network can be used to study how the nervous system can compensate for the loss of synaptic input. Individual neurons were specifically ablated in the giant fiber system of Drosophila, with a focus on two neurons: the presynaptic giant fiber (GF) and the postsynaptic tergotrochanteral motor neuron (TTMn). The GF synapses with the ipsilateral TTMn, which is crucial to the escape response. Ablating one of the GFs in the 3rd instar brain, just after the GF starts axonal growth, permanently removes the cell during the development of the CNS. The remaining GF reacts to the absent neighbor and forms an ectopic synaptic terminal to the contralateral TTMn. This atypical, bilaterally symmetric terminal innervates both TTMns, as demonstrated by dye coupling, and drives both motor neurons, as demonstrated by electrophysiological assays. In summary, the ablation of a single interneuron demonstrates synaptic competition between a bilateral pair of neurons that can compensate for the loss of one neuron and restore normal responses to the escape circuit.

Author Spotlight: Investigating the Mechanisms of Neural Circuit Assembly and Synapse Formation in Drosophila

This protocol demonstrates the laser cell ablation of individual neurons in intact Drosophila larvae. The method enables the study of the effect of reducing competition between neurons in the developing nervous system.

The protocol describes single-neuron ablation with a 2-photon laser system in the central nervous system (CNS) of intact Drosophila melanogaster larvae. ...

Testing Monosynaptic Connections Using Tetrodotoxin and Tetrodotoxin-Resistant Sodium Channels

Source: Zhang, X. et al., <a href="https://app.jove.com/v/57052">Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels.</a> J. Vis. Exp. (2018)
This video demonstrates testing monosynaptic connections using a transgenic fly. The fly has tetrodotoxin-resistant sodium channels and red light-activated channels in its presynaptic neurons, which are optogenetically excited to transmit signals to postsynaptic neurons.

Testowanie połączeń monosynaptycznych za pomocą tetrodotoksyny i kanałów sodowych opornych na tetrodotoksynę

1. Prepare Flies and Identify GAL4 promoter Lines

	Select a GAL4 promoter line and cross it with flies carrying both the UAS-NaChBac transgene and an ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

In Vivo Calcium Imaging Using Synaptically Localized Calcium Sensors in Drosophila melanogaster

Source: Hancock, C. E. et al., <a href="https://app.jove.com/v/60288">In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster.</a>&#160;J. Vis. Exp. (2019).
This video demonstrates a method for in vivo calcium imaging using synaptically localized calcium sensors in flies. It outlines the steps for preparing the flies for microscopy, conducting olfactory conditioning, and visualizing odor-induced calcium influx.

In vivo (in vivo) Obrazowanie wapnia za pomocą synaptycznie zlokalizowanych czujników wapnia u Drosophila melanogaster

1. Transgenic fruit flies,&#160;Drosophila melanogaster

	Cross female virgin and male flies (raised at 25 &#176;C in 60% relative humidity on a 12 h ...

Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels

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

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Electrophysiological Recording in the Drosophila Embryo

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

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network

Laser Cell Ablation in Intact Drosophila Larvae Reveals Synaptic Competition

Testowanie połączeń monosynaptycznych za pomocą tetrodotoksyny i kanałów sodowych opornych na tetrodotoksynę

In vivo (in vivo) Obrazowanie wapnia za pomocą synaptycznie zlokalizowanych czujników wapnia u Drosophila melanogaster