We would like to thank Gilad Silberberg, Michele Pignatelli, Thomas K. Berger, Luca Gambazzi, and Sonia Garcia for valuable advice on improvements for the patch-clamp procedure automation. We thank Rajnish Ranjan for valuable advice and assistance with software implementation. This work was funded in part by the EU Synapse project&#160;and partly by the Human Frontiers Science Program.

1. Equipment Preparation
<ol>
	<li>Control manipulators from a computer
	<ol>
		<li>Connect each micromanipulator controller box to a computer through serial ports (RS-232).</li>
		<li>Implement the commands for positioning, querying and adjusting settings to be sent via the serial port. Given speed and hardware compatibility issues C/C++ is recommended as the programming language.</li>
		<li>Standardize the reference system of the manipulators so that zero is the closest possible position with respect to the motors and positive movement is directed away from the motors.</li>
		<li>Position the microscope at its central position 2 mm above the specimen focal plane (coordinates [0, 0, 2000]).</li>
		<li>Store, for each manipulator, the local coordinates that allow the tip of a pipette to be observed in the center of the field of view of the microscope. This is the initial reference point for each electrode.</li>
	</ol>
	</li>
	<li>Visualize electrode positions. It is very useful to be able to track the position of each electrode during an experiment. A graphical representation is the most intuitive way of accomplishing this. To that end the reference systems of each electrode and that of the microscope must be matched. A simple way to accomplish this is:
	<ol>
		<li>Bring the tip of an electrode to the center of the field of view.</li>
		<li>Store the position on each axis of the manipulator and each axis of the microscope.</li>
		<li>Execute with the x-axis of the manipulator a relatively large movement (1 mm).</li>
		<li>Locate the tip once more moving only the microscope.</li>
		<li>Calculate the difference in the microscope position since it was last measured. These are the projections of the electrode axis movement onto the microscope axes.</li>
		<li>Repeat steps 2.3 - 2.5 for the Y and Z axes of the electrode manipulator. This allows a matrix of projections onto the microscope axes to be determined (Figure 2) also called cosine matrix: 
		<img alt="Equation 1" src="/files/ftp_upload/50630/50630eq1.jpg" /></li>
		<li>Invert this matrix to make it possible to calculate the movements, in three dimensions, required by the electrode manipulator to reach a given position in the microscope coordinate system: 
		<img alt="Equation 2" src="/files/ftp_upload/50630/50630eq2.jpg" /></li>
		<li>Using the initial reference point and the cosine matrix determine the position of each electrode in microscope coordinates.</li>
		<li>Display graphically, based on the microscope coordinates, the position of each electrode at regular intervals. We chose to use C/C++ drawing libraries (GDI+) to draw electrode positions every 40 msec.</li>
		<li>Repeat steps 1.2.1 - 1.2.7 every time manipulator angles are changed or if position changes of several millimeters are needed, for example, when the type of electrode is changed.</li>
	</ol>
	</li>
	<li>Enable storing the positions of relevant features in the tissue, such as cells or anatomical reference points, in microscope coordinates.</li>
	<li>Acquire video and overlay relevant information.
	<ol>
		<li>Mechanically align the x-axis of motion of the microscope with the horizontal axis of the microscope camera.</li>
		<li>Install on the computer a framegrabber with live video and overlay capability and a software development kit (SDK).</li>
		<li>Implement live video display operation with the SDK.</li>
		<li>Convert the microscope coordinate system to the camera reference system by the appropriate translation and scaling.</li>
		<li>Draw the relevant features in camera coordinates and overlay on the live video the resulting image at regular intervals of about 40 msec (Figure 3).</li>
	</ol>
	</li>
	<li>Control Amplifiers.
	<ol>
		<li>Use the amplifier software to control the amplifier settings from the interface.</li>
	</ol>
	</li>
	<li>Control oscilloscopes.
	<ol>
		<li>Connect the oscilloscopes to the PC using serial ports.</li>
		<li>Determine the oscilloscope scale, coupling and temporal resolution for the different steps of the patch-clamp procedure in voltage-clamp (e.g. electrode in bath, seal formation, whole-cell configuration) and current-clamp.</li>
		<li>Send the appropriate oscilloscope setting commands whenever amplifier commands are issued from the interface.</li>
	</ol>
	</li>
	<li>Control pipette pressure.
	<ol>
		<li>Assemble a pressure control system according to Figure 4.
		<ol>
			<li>Use a 12 V/5 V power supply to supply each electronic component appropriately.</li>
			<li>Connect the output of one membrane pump to the positive pressure buffer (a 100 ml container).</li>
			<li>Connect the input of one membrane pump to the negative pressure buffer (a 100 ml container).</li>
			<li>Connect the pipette holder tubing to a pressure sensor in the pressure control system and a pneumatic valve that connects to the main pressure compartment.</li>
			<li>Connect each of the pressure buffers to a valve connected to the main pressure compartment.</li>
			<li>Connect a valve between the main pressure compartment and the atmosphere.</li>
			<li>Connect a pressure sensor to the main pressure compartment.</li>
			<li>Connect a pressure sensor to each buffer.</li>
			<li>Connect the pressure control system to a data acquisition board.</li>
			<li>Connect each pressure sensor to one analog input.</li>
			<li>Connect each valve to a digital output.</li>
		</ol>
		</li>
		<li>Remove atmospheric pressure offset from all sensors by opening all valves except those connecting to the membrane pumps and subtracting the measured pressure.</li>
		<li>Implement pressure control
		<ol>
			<li>Define in the interface a control to activate or de-activate positive pressure control for each pipette.</li>
			<li>Define a minimal positive pressure for the pipettes of about 70 mbar.</li>
			<li>Periodically (every 0.5 sec) detect whenever pressure in pipettes under active pressure control drops below the set threshold.</li>
			<li>Upon threshold crossing open the positive pressure buffer to the main pressure compartment and this compartment towards the pipette in question during brief periods (20 msec) until pressure in the pipettes is above threshold. Close all other valves.</li>
			<li>De-activate further pressure control as the final approach towards a cell of interest is initiated.</li>
		</ol>
		</li>
		<li>Apply negative pressure for seal formation
		<ol>
			<li>Close all valves and open the negative pressure buffer valve towards the main pressure compartment and this compartment towards the pipette in question for the duration the experimenter requires by keeping a button pressed.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Centralize commands onto a human interface device.
	<ol>
		<li>Connect a commercially available wireless gamepad to the PC.</li>
		<li>Implement readout of joystick status. For instance, use the DirectX libraries for C/C++ to perform readouts every 5 msec.</li>
		<li>Compare current status with previous status to detect which buttons have been pressed, released or kept pressed since last time step.</li>
		<li>Assign functions to each button in the gamepad. An example of this mapping is shown in Figure 5.</li>
	</ol>
	</li>
</ol>
2. Patch-clamp Procedure
<ol>
	<li>Prepare the brain slices of the region of interest.</li>
	<li>Place a brain slice of interest, with the region of interest in the center of the microscope&#39;s displacement range.</li>
	<li>Cell Selection
	<ol>
		<li>Identify cells of interest by browsing with the microscope. Store the position of the cells based on the microscope coordinate system with a right mouse-click on the live video display on top of the cell of interest.</li>
		<li>The graphical interface will display the selected cells as well as the micropipettes for a global overview and software will add a marker on the cell position that will be overlaid on the images. Additionally an image of the cell is captured for future reference in the file &#39;Cell#.jpg&#39;.</li>
	</ol>
	</li>
	<li>Attribution of cells to pipettes
	<ol>
		<li>After selecting the cells of interest, assign which pipette will record each cell. The graphical interface provides assignment controls that should be set. Visualize a preview of the final configuration by selecting the checkbox &#39;Show Final Positions&#39;.</li>
		<li>Select each pipette for which the final position preview is desired or check the &#39;Select All&#39; checkbox. Disable the current position display for better visualization if needed.</li>
	</ol>
	</li>
	<li>Prepare the Pipettes
	<ol>
		<li>Fill the pipettes (6 - 8 M&#937; are usually best if many pipettes are used) with intracellular solution and load them into their holders. Place the headstages in their fixations but do not slide them forward to avoid touching the bath with the pipette tip.</li>
		<li>Enable the positive pressure control and select all pipettes to ensure that the tips will remain clean. Gently slide each headstage in place.</li>
	</ol>
	</li>
	<li>Locating the pipette tips
	<ol>
		<li>Position the microscope to a central position with focus 3 mm above the slice by pressing button R2 while holding button &#39;A&#39;. Use the corresponding position for each manipulator as stored from the previous experiment by pressing button L2 while holding button A. 
		Note: At this point there should be either a distinctive shadow of a pipette in view or a small movement along the pipette&#39;s axis should be enough to observe it in most cases. Compensation for the slight differences in pipette shape has to be performed manually.</li>
		<li>Bring the pipette tip into focus. Place the tip on the red central dot of the video display without moving the microscope (it should still be at position [0, 0, 2000]).</li>
		<li>Inform the software that the pipette is at the center of the display by pressing button Z while holding button C. After each pipette tip is located, send that pipette backwards so that the following one can be located by pressing button L1 while holding button A.</li>
	</ol>
	</li>
	<li>Approaching the cells
	<ol>
		<li>Once all pipette tips have been precisely located and each pipette is attributed to a cell, automatically position the pipettes close to their respective cells. Simply right-click in the center of the group of cells and select the option &#39;Cluster&#39; on the popup menu. On the cluster options window that will appear, select all the pipettes that you want to position at this time and click on &#39;Go with all checked pipettes&#39;. Repeat this operation for each cluster of cells of interest.</li>
		<li>Perform the final approach manually. The position of the pipettes relative to the cells can be specified differently but usually is simply 200 &#181;m away from the cell in the axis of the pipette and 200 &#181;m above it in the vertical direction, which is enough to keep the pipette&#39;s tip outside the tissue. Wait until the positioning of the pipettes is finished and move the microscope towards a pipette by pressing button R1 while holding button C.</li>
		<li>Recalibrate each pipette position by focusing the microscope on its tip anywhere in the video display and pressing button B while holding button C. A square grid should briefly appear indicating the identified position of the pipette. If the position is not correct, position the tip on the central dot of the video display and press button Z while holding button C.</li>
	</ol>
	</li>
	<li>Establish the cell-attached configuration
	<ol>
		<li>Confirm that the cell of interest for the current pipette is correctly marked. Otherwise move the microscope to match the cell of interest with the central red dot. Mark the cell by pressing button Y while holding button C. Press button L2 while holding button C to position the pipette 200 &#181;m away from its assigned cell, the microscope will automatically move to the corresponding position.</li>
		<li>Adjust the pipette position so it matches the red dot. Adjust the pipette offset by pressing button R1 while holding button X.</li>
		<li>Activate the test pulse by pressing button L1 while holding button X. Slowly approach the pipette to its attributed cell.</li>
		<li>Upon observing the formation of a dimple on the surface of the cell&#39;s membrane apply a brief pulse of negative pressure by pressing button Y while holding button Z to allow the applied pressure to reach the cell. A holding potential of about -65mV should established at this point by pressing the button L2 while holding button X.</li>
	</ol>
	</li>
	<li>Whole-cell configuration
	<ol>
		<li>Once a Giga-seal is formed for each cell, start rupturing the membranes by applying negative pressure.</li>
	</ol>
	</li>
	<li>Perform recordings
	<ol>
		<li>Use the stimulation/acquisition system to perform your recordings. Apply pulses or trains of pulses on one individual cell at a time and observe responses on the remaining cells to map connectivity among the recorded cells.</li>
	</ol>
	</li>
	<li>Recede pipettes
	<ol>
		<li>Once recordings are finished, recede pipettes slowly from the tissue by right-clicking on the Table radio-button. Select &#39;Recede-&#62;Pipettes 500 &#181;m&#39; to have the pipettes recede a short distance along their axes. Observe the drift in potential of the cells (clearing the tips by applying some positive pressure may help).</li>
		<li>To recede pipettes all the way back, set the positioning speed to &#39;Fast&#39; and repeat the same operation but choose the &#39;All&#39; option. Remove the used pipettes by gently sliding out the headstages and unscrewing the pipettes from the holders. It is useful to avoid twisting the holders in their sockets since this might greatly interfere with the pipette tip position.</li>
	</ol>
	</li>
</ol>

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	<li>Perin, R., Berger, T. K., Markram, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+synaptic+organizing+principle+for+cortical+neuronal+groups.">A synaptic organizing principle for cortical neuronal groups.</a> Proc. Natl. Acad. Sci. U.S.A. 108, 5419-5424 (2011).</li><li>Berger, T. K., Silberberg, G., Perin, R., Markram, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brief+Bursts+Self-Inhibit+and+Correlate+the+Pyramidal+Network.">Brief Bursts Self-Inhibit and Correlate the Pyramidal Network.</a> PLoS Biol. 8, e1000473 (2010).</li><li>Fino, E., Yuste, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dense+inhibitory+connectivity+in+neocortex.">Dense inhibitory connectivity in neocortex.</a> Neuron. 69, 1188-1203 (2011).</li><li>Packer, A. M., Yuste, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dense,+Unspecific+Connectivity+of+Neocortical+Parvalbumin-Positive+Interneurons:+A+Canonical+Microcircuit+for+Inhibition.">Dense, Unspecific Connectivity of Neocortical Parvalbumin-Positive Interneurons: A Canonical Microcircuit for Inhibition.</a> J. Neurosci. 31, 13260-13271 (2011).</li><li>Berger, T. K., Perin, R., Silberberg, G., Markram, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Frequency-dependent+disynaptic+inhibition+in+the+pyramidal+network:+a+ubiquitous+pathway+in+the+developing+rat+neocortex.">Frequency-dependent disynaptic inhibition in the pyramidal network: a ubiquitous pathway in the developing rat neocortex.</a> J. Physiol. 587, 5411-5425 (2009).</li><li>Kodandaramaiah, S. B., Franzesi, G. T., Chow, B. Y., Boyden, E. 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The authors declare that they have no competing financial interests.

An immediate question usually arises concerning the rate of success of the procedure we described. For high success rates preparation is essential. Pipettes must have tip openings that are adequate for the cells beings recorded. Filtering the intracellular solution to avoid clogged pipettes is also important. Extremely clean, freshly pulled pipettes are another requirement. A binomial distribution is the simplest model that can be used to understand how these issues affect the final yield. It is reasonable to expe...

The capacity to record and stimulate multiple sites with micrometer precision is extremely useful for experimentally achieving a better understanding of neuronal systems. Many techniques have been developed to this end but none allow the submillivolt resolution achieved by the patch-clamp technique, essential for studying subthreshold activity and individual postsynaptic potentials. Here we cover the development of a twelve-electrode computer-assisted patch-clamp system aimed at simultaneously recording and stimulating a large number of individual cells with sufficient precision for the study of neuronal connectivity. While many other applications can be conceived for such a system, it lends itself particularly well to the study of synaptic connectivity given that the number of possible connections within a group of neurons grows proportionally to the square of the number of neurons in question. Therefore, while a system with three electrodes allows testing the occurrence of up to six connections and most often recording a single one, recording twelve neurons allows testing the occurrence of up to 132 connections and frequently observing over one dozen (Figure 1). The observation of dozens of connections simultaneously makes it possible to analyze the organization of small networks and infer statistical properties of the network structure that cannot be probed otherwise1. Moreover, precise stimulation of numerous cells also allows the quantification of recruitment of postsynaptic cells2.

<table><tbody><tr><td>Microscope</td><td>Olympus</td><td>BX51WI</td><td>40X Immersion Objective</td></tr><tr><td>Manipulators</td><td>Luigs Neumann</td><td>SM-5</td><td>Serial protocol used</td></tr><tr><td>Amplifiers</td><td>Axon Instruments</td><td>MultiClamp 700B</td><td>SDK used</td></tr><tr><td>Camera</td><td>Till Photonics</td><td>VS 55</td><td>BNC analog output</td></tr><tr><td>Framegrabber</td><td>Data Translation</td><td>DT3120</td><td>SDK used</td></tr><tr><td>Oscilloscopes</td><td>Tektronix</td><td>TDS 2014</td><td>Serial communication</td></tr><tr><td>Data acquisition</td><td>InstruTECH</td><td>ITC 1600</td><td></td></tr><tr><td>Data acquisition</td><td>National Instruments</td><td>PCI-6221</td><td>Library used (.dll)</td></tr><tr><td>Pressure valve</td><td>SMC</td><td>SMC070C-6BG-32</td><td></td></tr><tr><td>Pressure sensor</td><td>Honeywell</td><td>24PCDFA6G</td><td></td></tr><tr><td>Membrane pump</td><td>Schego</td><td>Optimal</td><td></td></tr></tbody></table>

a computer-assisted multi-electrode patch-clamp system

The patch-clamp technique is today the most well-established method for recording electrical activity from individual neurons or their subcellular compartments. Nevertheless, achieving stable recordings, even from individual cells, remains a time-consuming procedure of considerable complexity. Automation of many steps in conjunction with efficient information display can greatly assist experimentalists in performing a larger number of recordings with greater reliability and in less time. In order to achieve large-scale recordings we concluded the most efficient approach is not to fully automatize the process but to simplify the experimental steps and reduce the chances of human error while efficiently incorporating the experimenter's experience and visual feedback. With these goals in mind we developed a computer-assisted system which centralizes all the controls necessary for a multi-electrode patch-clamp experiment in a single interface, a commercially available wireless gamepad, while displaying experiment related information and guidance cues on the computer screen. Here we describe the different components of the system which allowed us to reduce the time required for achieving the recording configuration and substantially increase the chances of successfully recording large numbers of neurons simultaneously.

Following the methods described above we succeeded in performing whole-cell recording of up to twelve neurons simultaneously, nearly doubling the largest number of neurons simultaneously patch-clamped thus far. Examples of networks of direct synaptic connections between Pyramidal Neurons recorded in Layer V of the somatosensory cortex of rats are shown in Figure 6.
The determination of connection probability profiles as a function of inter-somatic distance for a given cell-typ...

Watch this Scientific Journal Video about A Computer-assisted Multi-electrode Patch-clamp System at JoVE.com

A Computer-assisted Multi-electrode Patch-clamp System

Multi-electrode patch-clamp recordings constitute a complex task. Here we show how, by automating of many of the experimental steps, it is possible to accelerate the process leading to qualitative improvement in performance and number of recordings.

The patch-clamp technique is today the most  well-established method for recording electrical activity from individual  neurons or their subcellular ...

a-computer-assisted-multi-electrode-patch-clamp-system

Research

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Neuroscience

13.7K Views.  Ecole Polytechnique Federale de Lausanne. Multi-electrode patch-clamp recordings constitute a complex task. Here we show how, by automating of many of the experimental steps, it is possible to accelerate the process leading to qualitative improvement in performance and number of recordings.

Video: A Computer-assisted Multi-electrode Patch-clamp System

Multi-electrode Array Recordings of Human Epileptic Postoperative Cortical Tissue

Epilepsy, affecting about 1% of the population, comprises a group of neurological disorders characterized by the periodic occurrence of seizures, which disrupt normal brain function. Despite treatment with currently available antiepileptic drugs targeting neuronal functions, one third of patients with epilepsy are pharmacoresistant. In this condition, surgical resection of the brain area generating seizures remains the only alternative treatment. Studying human epileptic tissues has contributed to understand new epileptogenic mechanisms during the last 10 years. Indeed, these tissues generate spontaneous interictal epileptic discharges as well as pharmacologically-induced ictal events which can be recorded with classical electrophysiology techniques. Remarkably, multi-electrode arrays (MEAs), which are microfabricated devices embedding an array of spatially arranged microelectrodes, provide the unique opportunity to simultaneously stimulate and record field potentials, as well as action potentials of multiple neurons from different areas of the tissue. Thus MEAs recordings offer an excellent approach to study the spatio-temporal patterns of spontaneous interictal and evoked seizure-like events and the mechanisms underlying seizure onset and propagation. Here we describe how to prepare human cortical slices from surgically resected tissue and to record with MEAs interictal and ictal-like events ex vivo.

We here describe how to perform multi-electrode array recordings of human epileptic cortical tissue. Epileptic tissue resection, slice preparation and multi-electrode array recordings of interictal and ictal events are demonstrated in detail.

Epilepsy, affecting about 1% of the population, comprises a group of neurological disorders characterized by the periodic occurrence of seizures, which ...

Medicine

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Biology

Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a challenging and low-throughput technique due to its complexity and high reliance on user operation and control. This manuscript demonstrates an image-guided automatic patch clamp system for in vitro whole-cell patch clamp experiments in acute brain slices. Our system implements a computer vision-based algorithm to detect fluorescently labeled cells and to target them for fully automatic patching using a micromanipulator and internal pipette pressure control. The entire process is highly automated, with minimal requirements for human intervention. Real-time experimental information, including electrical resistance and internal pipette pressure, are documented electronically for future analysis and for optimization to different cell types. Although our system is described in the context of acute brain slice recordings, it can also be applied to the automated image-guided patch clamp of dissociated neurons, organotypic slice cultures, and other non-neuronal cell types.

This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for nerve fibers in the central and peripheral nervous systems. This method is particularly applicable to dorsal root ganglia (DRG), which are primary sensory neurons that exhibit a pseudo-unipolar structure of nervous processes. The patterns and features of the action potentials passed along axons are recordable in these neurons. The present study uses in vivo single-fiber recordings to observe the conduction failure of sciatic nerves in complete Freund&#8217;s adjuvant (CFA)-treated rats. As the underlying mechanism cannot be studied using in vivo single-fiber recordings, patch-clamp-recordings of DRG neurons are performed on preparations of intact DRG with the attached sciatic nerve. These recordings reveal a positive correlation between conduction failure and the rising slope of the after-hyperpolarization potential (AHP) of DRG neurons in CFA-treated animals. The protocol for in vivo single fiber-recordings allows the classification of nerve fibers via the measurement of conduction velocity and monitoring of abnormal conditions in nerve fibers in certain diseases. Intact DRG with attached peripheral nerve allows observation of the activity of DRG neurons in most physiological conditions. Conclusively, single-fiber recording combined with electrophysiological recording of intact DRGs is an effective method to examine the role of conduction failure during the analgesic process.

Single-fiber recording is an effective electrophysiological technique that is applicable to the central and peripheral nervous systems. Along with the preparation of intact DRG with the attached sciatic nerve, the mechanism of conduction failure is examined. Both protocols improve the understanding of the peripheral nervous system&#39;s relationship with pain.

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for ...

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.

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

Patch Clamp Electrophysiology

Neuron cell membranes are populated with ion channels that control the movement of charge into and out of the cell, thereby regulating neuron firing. One extremely useful technique for investigating the biophysical properties of these channels is called patch clamp recording. In this method, neuroscientists place a polished glass micropipette against a cell and apply suction to form a high resistance seal. This process isolates a small &#8220;patch&#8221; of membrane that contains one or more ion channels. Using an electrode within the micropipette, researchers can &#8220;clamp&#8221; or control the electrical properties of the membrane, which is important for analysis of channel activity. The electrode also allows for changes in the voltage across the membrane, or the flow of ions through the membrane, to be recorded. This video begins with an overview of the principles behind patch clamp electrophysiology, an introduction to the necessary equipment, and descriptions of the various patch configurations, including whole cell, cell-attached, perforated, inside-out, and outside-out patches. Next, the key steps of a typical whole-cell patch clamp experiment are outlined, in which a current-voltage (IV) curve is generated. Finally, applications of patch clamp recording are provided to demonstrate how the biophysical properties of ion channels, cell excitability, and neuroactive compounds are evaluated in neurophysiology labs today.

Patch Clamp Electrophysiology: Principle &amp; Applications

Neuron cell membranes are populated with ion channels that control the movement of charge into and out of the cell, thereby regulating neuron firing. One ...

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Establishing a Whole-Cell Configuration through the Patch Clamp Method

Source: Segev, A. et al., <a href="https://app.jove.com/v/54024">Whole-cell Patch-clamp Recordings in Brain Slices.</a>&#160;J. Vis. Exp.&#160;(2016)
This video demonstrates the whole-cell patch clamp technique in a mouse brain slice. This technique establishes a stable connection between micropipette and neuron membrane, which is further disrupted to establish a whole-cell configuration to study the neuron&#39;s health and physiological responses.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Amphotericin-B Mediated Perforated Patch-Clamp Analysis: An Electrophysiological Technique to Study Ionic Currents in Urinary Bladder Cells

Source: Malysz, J. et al. <a href="https://www.jove.com/video/59884" target="_blank">Preparation and Utilization of Freshly Isolated Human Detrusor Smooth Muscle Cells for Characterization of 9-Phenanthrol-Sensitive Cation Currents.</a> J. Vis. Exp. (2020)
This video demonstrates the characterization of cationic currents from transient receptor potential melastatin type 4 (TRPM4) ion channels in detrusor smooth muscle cells using amphotericin-B mediated perforated patch-clamp technique. The technique helps evaluate the biophysical and pharmacological properties of ion channels.

Source: Malysz, J. et al. Preparation and Utilization of Freshly Isolated Human Detrusor Smooth Muscle Cells for Characterization of ...

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Whole-Cell Patch Clamp Electrophysiology: A Method to Study Electrical Properties of Neurons

Source: Roy, B., et al. <a href="https://www.jove.com/v/50551/" target="_blank">Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells</a>. J. Vis. Exp. (2013).
This protocol describes a technique called whole-cell patch clamp electrophysiology, which is a method to study electrical properties of Mauthner neurons and other reticulospinal cells in zebrafish embryos.

Source: Roy, B., et al. Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells. J. Vis. Exp. (2013).
This protocol describes a technique called ...

A Computer-assisted Multi-electrode Patch-clamp System

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Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

One-channel Cell-attached Patch-clamp Recording

Patch Clamp Electrophysiology

Establishing a Whole-Cell Configuration through the Patch Clamp Method

Amphotericin-B Mediated Perforated Patch-Clamp Analysis: An Electrophysiological Technique to Study Ionic Currents in Urinary Bladder Cells

Whole-Cell Patch Clamp Electrophysiology: A Method to Study Electrical Properties of Neurons