Name: Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels (Video) | JoVE Journal
Uploaded: 2018-02-14T14:00:00.000+00:00
Duration: 9 min 55 s
Description: Learn patch-clamp electrophysiology to identify monosynaptic connections in Drosophila using tetrodotoxin-resistant sodium channels. This method reveals synaptic transmission properties and chemical nature of neuronal connections.

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5 minutes

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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</ol>

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

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...

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

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

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.

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

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.

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

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.

Electrophysiological Recording in the Drosophila Embryo

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.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

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.

Testing Monosynaptic Connections Using Tetrodotoxin and Tetrodotoxin-Resistant Sodium Channels

This video demonstrates the technique for recording synaptic currents from a synaptic bouton at the Drosophila larva's neuromuscular junction. This technique enables monitoring the activity of a single synaptic bouton, which is important for the study of mutant fly lines with enhanced spontaneous or asynchronous synaptic activity.

Recording Synaptic Currents at the Drosophila Larval Neuromuscular Junction

Here we describe a simple and widely accessible method to injure segmental nerves in Drosophila larvae to visualize and quantify neurodegeneration of motor neurons at the neuromuscular junction (NMJ) of third instar larvae.

A Simple Neuronal Mechanical Injury Methodology to Study Drosophila Motor Neuron Degeneration

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

We describe a method for retrograde tracing of the Drosophila embryonic motor neurons using lipophilic fluorescent dyes.

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

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.

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

This protocol assesses functional synaptic activity between defined neuronal partners in vivo in the central nervous system of Drosophila melanogaster larvae using genetically encoded tools. CsChrimson-mediated optogenetic stimulation of presynaptic cIVda sensory neurons induces calcium-dependent photoconversion of CaMPARI in postsynaptic Basin-4 interneurons.

Functional Characterization of Individual Pre- and Postsynaptic Partners In the Drosophila Larval Central Nervous System Using CaMPARI

Learn patch-clamp electrophysiology to identify monosynaptic connections in Drosophila using tetrodotoxin-resistant sodium channels. This method reveals synaptic transmission properties and chemical nature of neuronal connections.

Examining Monosynaptic Connections in Drosophila Using Tetrodotoxin Resistant Sodium Channels

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