We thank Dr. Fred Rieke for his kind explanation of the agar-embedded retinal slice preparation when we started using the protocol in our lab. We also thank Lori Vaskalis for the illustration of schematic overview and Drs. Veeramuthu Balakrishnan and Soyoun Cho for helpful comments on the text and video. This work was supported by an NEI-NIH RO1 grant, and was also partially supported by a Korea Research Foundation Grant funded by the Korean Government [KRF-2008-357-E00032].

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We have no competing interests to disclose.

A critical and difficult step in our protocol is the transfer of the piece of retina into the agar solution (protocol 3.4). It is necessary to carefully remove the vitreous humor and residual slice solution from the retinal piece and transfer it without distortion or bending. In order to accomplish this, we use a small spatula (21-401-25B, Fisherbrand) with a tip bent to form a 90&deg; angle, along with angled tip forceps (11251-35, Fine Science Tools), to position the retinal piece throughout the transf...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Low gelling-temperature agar</td>
 <td>Sigma</td>
 <td>A0701</td>
 <td>Agarose type VII-A</td>
 </tr>
 <tr>
 <td>Patch pipette</td>
 <td>World Precision Instruments</td>
 <td>1B150F-4</td>
 <td>Thick-walled (1.5 mm outer diameter) borosilicate glass</td>
 </tr>
 <tr>
 <td>Vertical puller</td>
 <td>Narishige</td>
 <td>PP830</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dental wax</td>
 <td>Cavex</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Spring scissors</td>
 <td>Fine Science Tools</td>
 <td>15003-08</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>45&deg; angled fine tip forceps</td>
 <td>Fine Science Tools</td>
 <td>11251-35</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Razor blade</td>
 <td>Personna</td>
 <td>&nbsp;</td>
 <td>Double-edged, cleaned with 70% ethanol and H2O</td>
 </tr>
 <tr>
 <td>Cylindrical tube</td>
 <td>Fisherbrand</td>
 <td>03-338-1B</td>
 <td>Polyethylene sample vials 2.5 ml</td>
 </tr>
 <tr>
 <td>Hyaluronidase</td>
 <td>Sigma</td>
 <td>H6254</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Vibratome slicer</td>
 <td>Leica</td>
 <td>VT1000S or VT1200S</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Upright microscope</td>
 <td>Olympus</td>
 <td>BX51WI</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>60x water-immersion objective</td>
 <td>Olympus</td>
 <td>LUMPlanFl</td>
 <td>NA 0.90</td>
 </tr>
 <tr>
 <td>CCD camera</td>
 <td>Sony</td>
 <td>XC-75</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Camera controller</td>
 <td>Hamamatsu</td>
 <td>C2400</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Monitor</td>
 <td>Sony</td>
 <td>&nbsp;</td>
 <td>13&#x201D; black and white monitor</td>
 </tr>
 <tr>
 <td>Syringe filter</td>
 <td>Nalgene</td>
 <td>&nbsp;</td>
 <td>0.2 &mu;m</td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Sutter Instrument</td>
 <td>MPC-200</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Lock-in amplifier</td>
 <td>HEKA</td>
 <td>&nbsp;</td>
 <td>EPC-9/10 amplifiers have software emulation</td>
 </tr>
 <tr>
 <td>Spinning disk laser confocal microscope</td>
 <td>Yokogawa</td>
 <td>CSU-X1</td>
 <td>Live cell imaging after patch clamp whole cell recording</td>
 </tr>
 <tr>
 <td>Slidebook software</td>
 <td>Intelligent Imaging Instruments (3i)</td>
 <td>&nbsp;</td>
 <td>Imaging data acquisition and analysis</td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>Sigma</td>
 <td>P6148</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Phosphate buffer solution</td>
 <td>GIBCO</td>
 <td>70013</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Superfrost slide</td>
 <td>Fisher Scientific</td>
 <td>&nbsp;</td>
 <td>Slide glass</td>
 </tr>
 <tr>
 <td>Anti-fading agents</td>
 <td>Biomeda corp.</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Confocal laser-scanning microscope</td>
 <td>Carl Zeiss</td>
 <td>LSM 710</td>
 <td>Imaging of fixed tissue</td>
 </tr>
 <tr>
 <td>Spatula</td>
 <td>Fisherbrand</td>
 <td>21-401-25B</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Manuel vertical slicer</td>
 <td>Narishige</td>
 <td>ST-20</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Oregon Green 488 BAPTA-1</td>
 <td>Invitrogen</td>
 <td>O-6806</td>
 <td>Ca2+ sensitive fluorescent dye</td>
 </tr>
 <tr>
 <td>Alexa Fluor 555 Hydrazide</td>
 <td>Invitrogen</td>
 <td>A-20501MP</td>
 <td>Fluorescent dye</td>
 </tr>
 </tbody>
</table>

patch-clamp capacitance measurements and ca2+ imaging at single nerve terminals in retinal slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Watch this Scientific Journal Video about Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices at JoVE.com

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

patch-clamp-capacitance-measurements-ca2-imaging-at-single-nerve

Research

JoVE Journal

Neuroscience

18.1K vistas.  Oregon Health and Science University. A continuación se describe un protocolo para la preparación de agar-incrustados trozos de retina que son adecuados para la electrofisiología y la Ca2 + imágenes. Este método permite estudiar la cinta del tipo de sinapsis en la retina utilizando microcircuitos directa patch-clamp grabaciones de una sola terminales nerviosas presinápticas.

Video: Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...

Methods for Cell-attached Capacitance Measurements in Mouse Adrenal Chromaffin Cell

Neuronal transmission is an integral part of cellular communication within the brain. Depolarization of the presynaptic membrane leads to vesicle fusion known as exocytosis that mediates synaptic transmission. Subsequent retrieval of synaptic vesicles is necessary to generate new neurotransmitter-filled vesicles in a process identified as endocytosis. During exocytosis, fusing vesicle membranes will result in an increase in surface area and subsequent endocytosis results in a decrease in the surface area. Here, our lab demonstrates a basic introduction to cell-attached capacitance recordings of single endocytic events in the mouse adrenal chromaffin cell. This type of electrical recording is useful for high-resolution recordings of exocytosis and endocytosis at the single vesicle level. While this technique can detect both vesicle exocytosis and endocytosis, the focus of our lab is vesicle endocytosis. Moreover, this technique allows us to analyze the kinetics of single endocytic events. Here the methods for mouse adrenal gland tissue dissection, chromaffin cell culture, basic cell-attached techniques, and subsequent examples of individual traces measuring singular endocytic event are described.

After exocytosis, fused plasma membrane is retrieved through a process known as endocytosis. This mechanism reforms new synaptic vesicles for the next round of release. Individual endocytic events are captured and analyzed through the use of the cell-attached capacitance recordings in mouse adrenal chromaffin cells.

Neuronal transmission is an integral part of cellular communication within the brain. Depolarization of the presynaptic membrane leads to vesicle fusion ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. Retinal neurons in the network comprise of numerous subtypes. More than 10 subtypes of bipolar cells, ganglion cells, and amacrine cells have been identified by morphological studies. Multiple subtypes of retinal neurons are thought to encode distinct features of visual signaling, such as motion and color, and form multiple neural pathways. However, the functional roles of each neuron in visual signal processing are not fully understood. The patch clamp method is useful to address this fundamental question. Here, a protocol to record light-evoked synaptic responses in mouse retinal neurons using patch clamp recordings in dark-adapted conditions is provided. The mouse eyes are dark-adapted O/N, and retinal slice preparations are dissected in a dark room using infrared illumination and viewers. Infrared light does not activate mouse photoreceptors and thus preserves their light responsiveness. Patch clamp is used to record light-evoked responses in retinal neurons. A fluorescent dye is injected during recordings to characterize neuronal morphological subtypes. This procedure enables us to determine the physiological functions of each neuron in the mouse retina.

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to blindness in many retinal diseases. Calcium (Ca2+), a key second messenger in photoreceptor signaling and metabolism, has been proposed to be indirectly linked with photoreceptor degeneration in various animal models. Systematically studying these aspects of cone physiology and pathophysiology has been hampered by the difficulties of electrically recording from these small cells, in particular in the mouse where the retina is dominated by rod photoreceptors. To circumvent this issue, we established a two-photon Ca2+ imaging protocol using a transgenic mouse line that expresses the genetically encoded Ca2+ biosensor TN-XL exclusively in cones and can be crossbred with mouse models for photoreceptor degeneration. The protocol described here involves preparing vertical sections (&#8220;slices&#8221;) of retinas from mice and optical imaging of light stimulus-evoked changes in cone Ca2+ level. The protocol also allows &#8220;in-slice measurement&#8221; of absolute Ca2+ concentrations; as the recordings can be followed by calibration. This protocol enables studies into functional cone properties and is expected to contribute to the understanding of cone Ca2+ signaling as well as the potential involvement of Ca2+ in photoreceptor death and retinal degeneration.

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

We describe a protocol to monitor Ca2+ dynamics in the axon terminals of cone photoreceptors using an ex-vivo&#160;slice preparation of the mouse retina. This protocol allows comprehensive studies of cone Ca2+ signaling in an important mammalian model system, the mouse.

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to ...

High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs). Physiological data have indicated that glutamate receptors at RGCs are expressed not only in postsynaptic but also in perisynaptic or extrasynaptic membrane compartments. However, precise anatomical locations for glutamate receptors at RGC synapses have not been determined. Although a high-resolution quantitative analysis of glutamate receptors at central synapses is widely employed, this approach has had only limited success in the retina. We developed a postembedding immunogold method for analysis of membrane receptors, making it possible to estimate the number, density and variability of these receptors at retinal ribbon synapses. Here we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of retinal fixation, 2) freeze-substitution, 3) postembedding immunogold electron microscope (EM) immunocytochemistry and, 4) quantitative visualization of glutamate receptors at ribbon synapses.

The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA ...

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.

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

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.

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

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 ...