monitoring electrical signals from the afferent neurons in an immobilized zebrafish larva

Monitoring Electrical Signals from the Afferent Neurons in an Immobilized Zebrafish Larva

Immobilize zebrafish larvae and transfer them into a 35-millimeter petri dish using a large-tipped transfer pipette. Remove as much of the surrounding solution as possible. Immerse larvae in 10 microliters of 0.1% alpha bungarotoxin for approximately five minutes. Wash the paralyzed larva with extracellular solution for 10 minutes. Then, use a transfer pipette to move the larva from the extracellular solution bath to the silicone-bottomed recording dish.
Under a stereomicroscope, gently position the larvae with fine-tipped forceps above the center of the silicone mat, lateral side up with the body&#39;s anterior and posterior ends running left to right. Using fine-tipped forceps, insert the etched pin orthogonally to the silicone through the dorsal notochord of the larvae, directly dorsal to the anus. Insert the second pin through the notochord near the end of the tail, and insert the third pin through the notochord dorsal of the gas bladder. Insert the fourth pin through the otic vesicle while providing slight rotation as the pin inserts into the encapsulant.
As a slight rotation is applied, watch for the tissue between the cleithrum and otic vesicle to reveal the cluster of afferent somata. Place the pinned lava under the 10 times objective on a fixed stage of the DIC microscope, and orient the myoseptal clefts of the muscle blocks parallel to the left head stage vector.
Place the ground wire into the bath solution, and ensure that it is connected to the left head stage. Fill the afferent recording electrode with 30 microliters of extracellular solution, and insert it into the right head stage pipette holder. Then, lower it into the dish solution while applying positive pressure produced by a pneumatic transducer.
Locate the electrode and bring the electrode tip over the specimen. Using a micromanipulator, lower the afferent electrode tip until it is holding position above the cleithrum. Increase the magnification to 40 times immersion and locate the intersection of the posterior lateral line nerve and cleithrum. Bring the electrode tip over the afferent ganglion, and lower the pipette until the tip contacts the epithelium. Gently, maneuver the electrode so that the entire tip circumference contacts the afferent ganglion.

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Preparation of materials for electrophysiological recordings
<ol>
	<li>Make a silicone elastomer-bottomed recording dish.
	<ol>
		<li>Dispense a thin layer of self-mixing silicone elastomer components (e.g., Sylgard) into a cover glass-bottomed tissue culture dish until it levels with the rim of a shallow well. Approximately 0.5 mL is sufficient.</li>
		<li>Cover and cure the dish for a minimum of 48 h at room temperature.</li>
	</ol>
	</li>
	<li>Make dissection pins.
	<ol>
		<li>Using a DC power supply, provide a negative charge (5 V) to a 100 mL beaker of etchant (3M KOH) and attach a tungsten wire (0.002 inch; 50.8 &#181;m diameter) to the positively charged output. 
		&#8203;CAUTION: Negative and positive wires should not contact one another during this procedure as you may run the risk of producing sparks, which could pose a potential fire hazard.</li>
		<li>Etch the tungsten wire by quickly and repeatedly dipping the tip into the etchant bath until the tip narrows to a sharp point. Under a stereomicroscope, cut the wire approximately 1 mm from the tip with a straight-edge razor blade. Repeat three more times, and then insert pins into the cured recording dish using fine forceps.</li>
	</ol>
	</li>
	<li>Prepare recording electrodes.
	<ol>
		<li>Using a horizontal micropipette puller with box filament, pull a borosilicate glass capillary tube (inner diameter: 0.86 mm, outer diameter: 1.50 mm) into electrodes with a 30 &#181;m diameter tip with slight taper (Figure 1 Ai) that will be used to record afferent neurons from the posterior lateral line.</li>
		<li>Pull an additional borosilicate glass capillary tube into a pair of electrodes with smaller tip diameters (1-5 &#181;m). Holding one electrode in each hand, gently run the tips across one another to break them to a ~30&#176; angle. Using a microforge, polish the beveled tip until smooth.</li>
	</ol>
	</li>
	<li>Generate the protocol in electrophysiology recording programs.
	<ol>
		<li>Ensure the right head stage is connected to the Channel 1 input in the back of the microelectrode current and voltage clamp amplifier for the afferent neuron recordings. 
		&#8203;NOTE: Computer specifications for the current and voltage clamp amplifier minimally require a 1 Ghz or better processor, Windows XP Pro or Mac OS X 10.46.6, a CD-ROM drive with 512 MB RAM, 500 MB hard drive space, and 2 USB ports.</li>
		<li>Open the computer-controlled amplifier software.</li>
		<li>Set Channel 1 to current-clamp mode by clicking the IC button.</li>
		<li>Input the following parameter under the I-Clamp 1 tab. Primary Output: 100x AC Membrane Potential (100,000 mV/mV), Gain: 1,000, Bessel: 1 kHz, AC: 300 Hz, Scope: Bypass. Secondary Output: 100x AC Membrane Potential (100 mV/mV), Gain: 1, Lowpass Filter: 10 Hz.</li>
		<li>Save the channel parameter as Ch1_Aff.</li>
		<li>Install and open the patch clamp electrophysiology software.</li>
		<li>Click on Configure and select Lab Bench to set up the analog signals by adding Ch1_Aff to Digitizer Channels (e.g., Analog IN #0 and Analog IN #1) connected by BNC coaxial cable to the amplifier&#39;s corresponding Channel 1 Scaled Outputs. Click on OK. 
		&#8203;NOTE: The minimum computer requirements for the digitizer are: a 1 Ghz or better processor, Windows XP Pro or Mac OS X 10.46.6, CD-ROM drive 512 MB RAM, 500 MB hard drive space, and 2 USB ports.</li>
		<li>Click on Acquire and select New Protocol.</li>
		<li>In the Mode/Rate tab, select Gap Free; under Acquisition Mode, set Trial Length to either Use available disk space (i.e., record until stopped) or a desired set Duration (hh:mm:ss), and then set the Sampling rate per signal (Hz) to 20,000 to maximize the resolution.</li>
		<li>Under the Inputs tab, select the Analog IN Channels that were previously configured (in step 1.4.6) and select Ch1_Aff for the corresponding channel.</li>
		<li>Under the Outputs tab, select Cmd 0 and Cmd 1 for Channel #0 and Channel #1, respectively.
		<ol>
			<li>Connect Channel 1 Command on the amplifier to Analog Output 0 on the digitizer via BNC coaxial cable.</li>
		</ol>
		</li>
		<li>The remaining tabs can remain under default settings. Click on OK and save the protocol.</li>
	</ol>
	</li>
</ol>
2. Solution preparation
<ol>
	<li>Prepare Hank&#39;s solution: 137 mM NaCl, 5.4 mM KCl, 0.25 mM Na2HPO4, 0.44 mM KH2PO4, 1.3 mM CaCl2, 1.0 mM MgSO4, 4.2 mM NaHCO4; pH 7.3. Dilute to 10% Hank&#39;s solution by adding the appropriate volume of deionized water to the stock.</li>
	<li>Prepare the extracellular solution: 134 mM NaCl, 2.9 mM KCl, 1.2 mM MgCl2, 2.1 mM CaCl2, 10 mM glucose, and 10 mM HEPES buffer, pH 7.8 adjusted with NaOH. Vacuum filter the extracellular solution through a filter with a 0.22 &#181;m pore size.</li>
	<li>Prepare &#945; -bungarotoxin: Dissolve 1 mg of lyophilized &#945;-bungarotoxin in 10 mL extracellular solution to produce 0.1% dilution.</li>
	<li>Prepare Euthanasia solution: 50% (mg/L) buffered pharmaceutical-grade MS-222 in 10% Hank&#39;s solution. 
	CAUTION: &#945;-bungarotoxin is a potent neurotoxin that paralyzes muscles by blocking cholinergic receptors. Gloves are required and eye protection is recommended while handling the paralytic.</li>
</ol>
3. Preparation of larvae for electrophysiological recordings
<ol>
	<li>Immobilize zebrafish larvae.
	<ol>
		<li>Use larvae from the laboratory-bred population of zebrafish (Danio rerio; 4-7 days post fertilization) and house in embryo solution (10% Hank&#39;s solution) at 27 &#176;C.</li>
		<li>Using a large-tipped transfer pipette, transfer the larva from the housing into a small Petri dish (35 mm) and remove as much of the surrounding solution as possible. 
		NOTE: Removing the embryo solution prevents dilution of the paralytic and increases its efficacy. The corner of a task wipe can be used to wick away the remaining embryo solution, and it is critical not to contact the larvae or leave them exposed to air.</li>
		<li>Immerse larvae in 10 &#181;L of 0.1% &#945;-bungarotoxin for approximately 5 min. 
		NOTE: The time necessary to immobilize larvae varies between preparations. A healthy preparation depends on closely monitoring sustained fast blood flow and decreased motor responses. Brief overexposure to the paralytic can lead to a slow decline in the preparation&#39;s overall health, even after a thorough washout. It is best to apply a wash before the larva is completely immobilized while it still shows signs of subtle muscle vibrations.</li>
		<li>Wash the paralyzed larva with the extracellular solution and bathe for 10 min. 
		NOTE: The washout allows for the larva to transition from subtle muscle vibrations to complete paralysis. Also, &#945;-bungarotoxin is an antagonist of the nicotinic acetylcholine receptor (nAChR) &#945;9-subunit, which is a critical component of the endogenous feedback circuit this protocol observes; however, this effect has been shown to be reversible in Xenopus and zebrafish hair cells after a 10 min washout.</li>
	</ol>
	</li>
	<li>Pin fish in the recording dish. 
	NOTE: Anesthesia (e.g., MS-222, Tricaine) is not required while preparing larval zebrafish (4-7 dpf) as it interferes with animal health. In fact, larval zebrafish are exempt from certain vertebrate protocols.
	<ol>
		<li>Using a transfer pipette, move the larva from the extracellular solution bath to the silicone-bottomed recording dish. Fill the remainder of the dish with the extracellular solution.</li>
		<li>Under a stereomicroscope, gently position the larvae with fine-tipped forceps above the center of the silicone mat, lateral side up, with the body&#39;s anterior and posterior ends running left to right, respectively. Then grasp an etched pin from the silicone mat using fine-tipped forceps and insert the pin, orthogonally to the silicone, through the dorsal notochord of the larvae directly dorsal to the anus. Insert the second pin through the notochord near the end of the tail and insert the third pin through the notochord dorsal of the gas bladder (Figure 1B). 
		NOTE: While inserting pins, it is important to target the center of the notochord width to prevent impinging blood flow or damaging surrounding musculature. The notochord is dorsal to the posterior lateral line nerve, so with proper pinning, no damage is expected to the lateral line sensory neurons. Insert after first contact so as to not disturb the surrounding tissue. Ideally, the diameter of the pin is less than half the width of the notochord to ensure clean insertion. If the pins exceed this width, repeat step 1.2 until the desired pin width is attained. If the blood flow slows after pinning, repeat from step 1.3 onward with a new specimen.</li>
		<li>Insert the fourth pin through the otic vesicle while providing slight rotation towards the anterior as the pin inserts into the encapsulant (Figure 1B). As a slight rotation is applied, watch for the tissue between the cleithrum and otic vesicle to reveal the cluster of afferent somata. 
		&#8203;NOTE: Angled insertion of the fourth pin is to ensure the exposure of the posterior lateral line afferent ganglion, which is otherwise obstructed by the large otic vesicle.</li>
	</ol>
	</li>
</ol>
4. Afferent neuron recording
<ol>
	<li>Fill the afferent recording electrode with 30 &#181;L of the extracellular solution, insert it into the right head stage pipette holder (Figure 1B, C), and lower it into the dish solution while applying positive pressure (1-2 mm Hg) produced by a pneumatic transducer.</li>
	<li>Locate and loosely attach to posterior lateral line afferent ganglion.
	<ol>
		<li>Using a micromanipulator, lower the afferent electrode tip until it is holding position above the cleithrum.</li>
		<li>Increase the magnification to 40x immersion and locate the intersection of the posterior lateral line nerve and cleithrum. Follow the lateral line nerve anterior from the cleithrum to where the fibers innervate the posterior lateral line afferent ganglion, distinguishable by the discrete cluster of soma (Figure 1F).</li>
		<li>Bring the electrode tip over the afferent ganglion and lower the pipette until the tip contacts the epithelium. Gently, maneuver the electrode so that the entire tip circumference contacts the afferent ganglion.</li>
		<li>Apply negative pressure (20-50 mm Hg) with the pneumatic transducer and hold. 
		NOTE: The negative pressure applied to the afferent ganglion is gentle. Increasing negative pressure can improve signal-to-noise, but afferent neuron health declines under sustained aggressive suction, which decreases the probability of a successful recording.</li>
	</ol>
	</li>
	<li>Record afferent neuron activity.
	<ol>
		<li>The left head stage should be connected to the amplifier, which relays the amplified signal into a digitizer that outputs the said signal into the patch clamp electrophysiology software to be monitored on an adjacent computer.</li>
		<li>In pClamp10, click on the Play button on the toolbar to monitor afferent neurons.</li>
		<li>Ensure that the whole cell, loose patch recording of afferent neurons is achieved once spikes occur spontaneously, roughly every 100-200 ms (Figure 1E).</li>
		<li>Gradually, increase the afferent neuron recording electrode pressure back to atmospheric (0 mm Hg) and hold for the remainder of the recording.</li>
	</ol>
	</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 mL beaker</td>
			<td>PYREX</td>
			<td>1000</td>
			<td>Resceptacle for etchant</td>
		</tr>
		<tr>
			<td>10x water immersion objective</td>
			<td>Olympus</td>
			<td>UMPLFLN10xW</td>
			<td>Low magnification for positioning larvae and recording electrode</td>
		</tr>
		<tr>
			<td>40x water immersion objective</td>
			<td>Olympus</td>
			<td>LUMPLFLN40XW</td>
			<td>Higher magnification for position electrode tip and establishing patch-clamp</td>
		</tr>
		<tr>
			<td>BNC coaxial cables</td>
			<td>ThorLabs</td>
			<td>2249-C-12</td>
			<td>Connecting amplifier and digitizer channels; require 4</td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries w/ filament</td>
			<td>Warner Instruments</td>
			<td>G150F-3</td>
			<td>Inner diameter: 0.86, outer diameter: 1.50; capillary glass used to form recording electrodes</td>
		</tr>
		<tr>
			<td>Computer</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Any computer should work</td>
		</tr>
		<tr>
			<td>DC Power Supply</td>
			<td>Tenma</td>
			<td>72-420</td>
			<td>Used for electrically etching dissection pins</td>
		</tr>
		<tr>
			<td>Electrophysiology digitizer</td>
			<td>Axon Instruments, Molecular Devices</td>
			<td>Axon DigiData 1440A</td>
			<td>Enables acquisition of patch-clamp data</td>
		</tr>
		<tr>
			<td>Filament</td>
			<td>Sutter Instrument Company</td>
			<td>FB255B</td>
			<td>2.5 mm box filament used in micropipette puller</td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Fine Science Tools</td>
			<td>Dumont #5 (0.05 x 0.02 mm) Item No. 11295-10</td>
			<td>Used to manipulate larvae and insert pins</td>
		</tr>
		<tr>
			<td>Fixed stage DIC microscope</td>
			<td>Olympus</td>
			<td>BX51WI</td>
			<td>Microscope used to visualize and establish patch-clamp recordings</td>
		</tr>
		<tr>
			<td>Flexible, tapered pipette tip</td>
			<td>Fisher Scientific</td>
			<td>02-707-169</td>
			<td>Flexible tips enable insertion into recording electrode to dispense extracellular solution at the tip</td>
		</tr>
		<tr>
			<td>FluoroDish</td>
			<td>World Precision Instruments Inc.</td>
			<td>FD3510-100</td>
			<td>Cover glass bottomed dish recording dish</td>
		</tr>
		<tr>
			<td>KimWipe</td>
			<td>KimTech</td>
			<td>34155</td>
			<td>Task wipe used for wicking away excess fluid from larvae</td>
		</tr>
		<tr>
			<td>Kwik-Gard</td>
			<td>World Precision Instruments Inc.</td>
			<td>710172</td>
			<td>Self-mixing sylgard elastomer</td>
		</tr>
		<tr>
			<td>Microelectrode amplifier</td>
			<td>Axon Instruments, Molecular Devices</td>
			<td>MultiClamp 700B</td>
			<td>Patch clamp amplifier for dual channel recordings</td>
		</tr>
		<tr>
			<td>Microforge</td>
			<td>Narishige</td>
			<td>MF-830 microforge</td>
			<td>To polish recording electrode</td>
		</tr>
		<tr>
			<td>Micromanipulator control unit</td>
			<td>Siskiyou</td>
			<td>MC1000-eR/T</td>
			<td>4-axis dial coordinator for controlling micromanipulator</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instrument Company</td>
			<td>Flaming/Brown P-97</td>
			<td>For pulling capillary glass into recording electrodes</td>
		</tr>
		<tr>
			<td>Microscope control unit</td>
			<td>Siskiyou</td>
			<td>MC1000e</td>
			<td>Positions the microscope around the fixed stage and preparation</td>
		</tr>
		<tr>
			<td>Motorized micromanipulator</td>
			<td>Siskiyou</td>
			<td>MX7600</td>
			<td>Positions the headstage and attached recording electrode for patch-clamp recording</td>
		</tr>
		<tr>
			<td>MultiClamp Commander</td>
			<td>Molecular Devices</td>
			<td>2.2.2</td>
			<td>Downloadable from Axon MultiClamp 700B Commander download page</td>
		</tr>
		<tr>
			<td>Optical air table</td>
			<td>Newport Corporation</td>
			<td>VH3036W-OPT</td>
			<td>Breadboard isolation table to float microscope and minimize vibrations during recordings</td>
		</tr>
		<tr>
			<td>pCLAMP</td>
			<td>Molecular Devices</td>
			<td>10.7.0</td>
			<td>Downloadable from Axon pCLAMP 10 Electrophysiology Data Acquisition &#38; Analysis Software Download page</td>
		</tr>
		<tr>
			<td>Petri-dish</td>
			<td>Falcon</td>
			<td>35-3001</td>
			<td>Used to immerse larvae in paralytic</td>
		</tr>
		<tr>
			<td>Pipette holder</td>
			<td>Molecular Devices</td>
			<td>1-HL-U</td>
			<td>Hold recording electrode and connect to the headstage</td>
		</tr>
		<tr>
			<td>Pneumatic transducer</td>
			<td>Fluke Biomedical Instruments</td>
			<td>DPM1B</td>
			<td>For controlling recording electrode internal pressure</td>
		</tr>
		<tr>
			<td>Potassium hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>221473-25G</td>
			<td>Etchant for etching dissection pins</td>
		</tr>
		<tr>
			<td>Silicone tubing</td>
			<td>Tygon</td>
			<td>14-169-1A</td>
			<td>Tubing to connect pneumatic transducer to pipette holder</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Carl Zeiss</td>
			<td>Stemi 2000-C</td>
			<td>Used to visualize pin tips and during preparation of larvae</td>
		</tr>
		<tr>
			<td>Straight edge razor blade</td>
			<td>Canopus</td>
			<td>order from amazon.com</td>
			<td>Cuts the tungsten wire while making dissection pins</td>
		</tr>
		<tr>
			<td>syringe</td>
			<td>Becton Dickinson Compoany</td>
			<td>309602</td>
			<td>Filled with extracellular solution to inject into recording electrodes</td>
		</tr>
		<tr>
			<td>Transfer pipette</td>
			<td>Sigma-Aldrich</td>
			<td>Z135003-500EA</td>
			<td>Single use, non-sterile pipette for transfering larvae</td>
		</tr>
		<tr>
			<td>Tricaine methanesulfonate</td>
			<td>Syndel</td>
			<td>12854</td>
			<td>Pharmaceutical aneasthetic used to euthanize larvae with high dosage.</td>
		</tr>
		<tr>
			<td>Tungsten wire</td>
			<td>World Precision Instruments Inc.</td>
			<td>715500</td>
			<td>0.002 inch, 50.8 &#956;m diameter; used to make dissection pins</td>
		</tr>
		<tr>
			<td>Vacuum filtration unit</td>
			<td>Sigma-Aldrich</td>
			<td>SCGVU11RE</td>
			<td>Single use, sterile, vacuum filtration units used to sterilize extracellular solution used for electrophysiology electrode ringer</td>
		</tr>
		<tr>
			<td>Voltage-clamp current-clamp headstage</td>
			<td>Molecular Devices</td>
			<td>CV-7B</td>
			<td>Supplied with MultiClamp 700B amplifier used as left and right headstages</td>
		</tr>
		<tr>
			<td>&#945;-bungarotoxin</td>
			<td>ThermoFisher</td>
			<td>B1601</td>
			<td>For immobilizing the larvae prior to recording</td>
		</tr>
	</tbody>
</table>

Source: Lunsford, E. T., et al. <a href="https://app.jove.com/v/62233">Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish.</a> J. Vis. Exp. (2021).
This video demonstrates the recording of electrical signals from the afferent neurons in a paralyzed zebrafish larva by positioning a micropipette near the lateral line ganglion and forming a loose seal for signal capture. The extracellular solution maintains neuronal viability during this process.

<img alt="Figure 1" src="/files/ftp_upload/22731/22731_Figure1.jpg" /> 
Figure 1: Simultaneous electrophysiological recording of posterior lateral line afferent neuron and ventral motor root activity. (A). Example of a loose-patch afferent (i) and ventral motor root (ii) recording electrodes. Scale bars represent 50 &#181;m. (B) Larval zebrafish are paralyzed and pinned in four locations (cross symbo...

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Start with an immobilized, paralyzed zebrafish larva in a recording chamber filled with an extracellular solution in an electrophysiology setup.
Fill an afferent micropipette with the extracellular solution containing ions that provide electrical conductivity for recording neuronal activity.
Assemble it into a micromanipulator containing a recording electrode connected to an amplifier for signal recording.
Apply positive air pressure to prevent the pipette from clogging.
Move the pipette tip along the lateral line nerve and position it adjacent to the afferent ganglion that contains a cluster of afferent neuronal cell bodies.
These neurons&#39; cellular processes are connected to the hair cells located along the lateral line, which sense water movements.
Then apply gentle negative pressure to form a loose seal between the micropipette and the afferent ganglion.
Record the electrical signals from the afferent neurons, which receive the sensory input from the hair cells in the immobilized larva.

24 vistas. Monitoreo de señales eléctricas de las neuronas aferentes en una larva de pez cebra inmovilizada

Video: Monitoreo de señales eléctricas de las neuronas aferentes en una larva de pez cebra inmovilizada - Experimento

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Las grabaciones de células enteras Patch-clamp en rodajas de cerebro

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Investigación

JoVE Journal

Neurociencias

Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze motor-learning induced plasticity, we trained rats using an accelerated rotor rod task. Rats performed the task 10 times at 30-s intervals for 1 or 2 days. Performance was significantly improved on the training days compared to the first trial. We then prepared acute brain slices of the primary motor cortex (M1) in untrained and trained rats. Current-clamp analysis showed dynamic changes in resting membrane potential, spike threshold, afterhyperpolarization, and membrane resistance in layer II/III pyramidal neurons. Current injection induced many more spikes in 2-day trained rats than in untrained controls.
To analyze contextual-learning induced plasticity, we trained rats using an inhibitory avoidance (IA) task. After experiencing foot-shock in the dark side of a box, the rats learned to avoid it, staying in the lighted side. We prepared acute hippocampal slices from untrained, IA-trained, unpaired, and walk-through rats. Voltage-clamp analysis was used to sequentially record miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs) from the same CA1 neuron. We found different mean mEPSC and mIPSC amplitudes in each CA1 neuron, suggesting that each neuron had different postsynaptic strengths at its excitatory and inhibitory synapses. Moreover, compared with untrained controls, IA-trained rats had higher mEPSC and mIPSC amplitudes, with broad diversity. These results suggested that contextual learning creates postsynaptic diversity in both excitatory and inhibitory synapses at each CA1 neuron.
AMPA or GABAA receptors seemed to mediate the postsynaptic currents, since bath treatment with CNQX or bicuculline blocked the mEPSC or mIPSC events, respectively. This technique can be used to study different types of learning in other regions, such as the sensory cortex and amygdala.

The slice patch clamp technique is an effective method for analyzing learning-induced changes in the intrinsic properties and plasticity of excitatory or inhibitory synapses.

Slice Patch Clamp técnica para analizar la plasticidad inducida por el aprendizaje

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Un canal celular conectado a Patch-clamp grabación

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Monitoring Electrical Signals from the Afferent Neurons in an Immobilized Zebrafish Larva

Transcripción

Monitoring Electrical Signals from the Afferent Neurons in an Immobilized Zebrafish Larva