The authors wish to acknowledge Alexei Bogdanov for the fabrication of patch-clamp chips at the CPFC, and Hue Tran, Ping Zhao and Matthew Shiu for assistance with assembly. Naweed Syed was supported by a Canadian Institute of Health Research (CIHR) grant. Collin Luk is the recipient of NSERC and Alberta Heritage Foundation for Medical Research (AHFMR) studentships.

1. Chip fabrication
The process described in 6 results in a 3 &mu;m thick self-standing film, a low 1 Mohm access resistance and a smooth-surfaced funnel shaped aperture that facilitates an intimate seal with the cell 9, see Figure 2. The chips are singulated and glued in Plexiglas packages with the aperture facing a hole connecting the chip to a subterranean channel. The gluing is dispensed in a way that minimizes the shunt capacitance to a nominal 17 pF. Packages are fitted with two 1.5 mm diameter and 6 mm long glass tubes as fluidic ports (Figure 3).
<img alt="Figure 2" src="/files/ftp_upload/3288/3288fig2.jpg" /> 
Figure 2. Scanning electron micrograph of a focused ion beam section of an NRCC patch-clamp chip shows a smooth silicon dioxide surface and funnel shape that favors intimate cell contacts, and a shallow orifice that accounts for a low access resistance.
<img alt="Figure 2" src="/files/ftp_upload/3288/3288fig2.jpg" /> 
Figure 3. A chip is glued at the bottom of the culture vial in a Plexiglas package fitted with subterranean fluidics and glass tubes.
2. Sterilization, priming and testing
The following steps should be performed in a biology safety cabinet to avoid contamination or plugging of the aperture.
<ul type="disc">
	<li>Sterilize chips in a Harrick basic plasma cleaner (www.harrickplasma.com) with a 0.1-0.3 mbar residual air pressure for 15 min at the maximum power of 18 W. Other air plasma systems may be used, with comparable power density (24 mW/cm3) and keeping the product of power density and time constant. The plasma treatment also renders the chips hydrophilic, which facilitates priming.</li>
	<li>Fit glass tubes with sterile Silastic Laboratory silicone tubing, 1mm I.D. x 2mm O.D. (Cole Parmer Cat. # 96115-08), 3inch on one side, 1 inch on the other side.</li>
	<li>Fill fluidics through the tubes with sterile filtered standard Phosphate Buffer Solution (PBS), making sure no bubble is trapped.</li>
	<li>Clip the long silicone tube while pressurizing the PBS from the short side at 1 atm. A pool of PBS seeping through the aperture might be visible in the vial. Clip the pressurized supply of the PBS and detach the short silicone tube from that supply.</li>
	<li>Fill vial with filtered PBS using a syringe.</li>
	<li>Immerse an Ag/Ag:Cl electrode in the short tube and a counter-electrode in the vial. An impedance meter is used to confirm that the access resistance is between 300kohm and 3Mohm and the shunt capacitance is between 10pF and 25pF. A chip with a lower resistance is likely to have a leak, and a higher capacitance is considered improper for use as it may not capture the fastest dynamics of the cell&prime;s electrophysiology 6. A chip with a lower capacitance or higher resistance is considered plugged, though it could be simply that a bubble is trapped in the orifice. Some chips are re-tested after 1H and found to have the correct electrochemical impedance.</li>
	<li>Flush the fluidic channel with sterile deionized water; empty the top vial and rinse with the same, twice.</li>
	<li>Immerse the packaged chip with tubes in fresh sterile deionized water for 30min then load with other chips in a sterile container filled with sterile deionized water. This ensures that chips remain hydrophilic and protected from contamination.</li>
</ul>
3. Chips preparation in cell biology lab
The following steps are performed in a HEPA filtered laminar flow hood using aseptic techniques, all solutions used in this procedure must be filtered sterilized prior to use using a 0.22&mu;m filter.
<ul type="disc">
	<li>Open a chip container under a HEPA filtered laminar flow hood and remove chips from container face down to avoid scooping possible floating contaminants into top vial.</li>
	<li>Rinse top of vial of chip 2x with freshly filtered sterile deionized water to remove any residual debris from the chip surface.</li>
	<li>Aspirate sterile deionized water off the top vial.</li>
	<li>Connect one end of the silastic tubing attached to the fluidic channel to a pressurized syringe containing sterile filtered deionized water. In our system, syringes filled with solution are pressurized by connection to a compressed air tank set to 20psi. To ensure sterility, the air is filtered (0.22 &mu;m filter) prior to entry into the syringe and our system also has an on/off valve that allows us to stop or apply pressure when needed.</li>
	<li>Using a hemostat, clamp the output end of the tubing.</li>
	<li>Open the on/off valves to pressurize the solution.</li>
	<li>To rinse the subterranen fluidic of the chip with fresh sterile deionized water, unclamp the output end of the tubing.</li>
	<li>To force the water up through the aperture, clamp the output end of the tubing with a hemostat.</li>
	<li>To avoid water drainage, clamp the input and output ends of the tubing with an hemostats and close the on/off valves.</li>
	<li>Detach the input end of the tubing from the syringe.</li>
	<li>Insert glass plugs into both ends of the tubing and remove the hemostats.</li>
	<li>Fill top vial with filter steriled deionized water.</li>
	<li>Place the lid of a 35 mm sterile dish into the base of a 100 mm sterile dish</li>
	<li>Place chip on top of the 35 mm lid and cover with 100 mm dish lid.</li>
	<li>Image chips to ensure that the membrane and aperture of the chips are free of debris and/or air bubbles. In our lab we use an upright microscope and a 20x long working distance objective.</li>
	<li>If debris and/or air bubbles are present, repeatedly flush fluidic channel and top vial. If the debris and/or air bubbles cannot be removed the chip is discarded.</li>
	<li>Once the chips are imaged, return to laminar flow hood and unplug both ends of the tubing.</li>
	<li>Attach one end of the tubing to a pressurized syringe containing physiological media.</li>
	<li>Clamp the output end of the tubing with a hemostat</li>
	<li>Open pressure valve, remove the hemostat from the output end of the fluidics and flush the fluidic channel with physiological media</li>
	<li>To rinse the subterranean fluidic with physiological media, open the on/off valves and remove the hemostat from the output end of the tubing.</li>
	<li>To force the physiological media up through the aperture, clamp the output end of the tubing with a hemostat.</li>
	<li>Clamp input end of the tubing with a hemostat and close the on/off valves.</li>
	<li>Remove the input end of the tubing from the syringe and plug both ends of the tubing with glas plugs.</li>
	<li>Remove the hemostats.</li>
	<li>Remove the water from the top vial.</li>
	<li>Fill the top vial with filter sterilized physiological media.</li>
	<li>Place the chip back into the 100 mm Petri dish.</li>
	<li>Place the base of a 35 mm dish into the 100 mm dish and fill it with filter sterilized deionized water to enrich humidity.</li>
	<li>Cover the dish until it is time for plating</li>
</ul>
4. Cell plating of snail neurons
The patch-clamp chips may be suitable for a variety of preparations. We are currently testing our chips with mammalian primary cortical neurons and have obtained preliminary results with cells cultured for 14 days 7, which indicates that our sterilization protocol is adequate and that the chips are not cytotoxic in long-term cultures. For the purpose of this protocol, snail neurons were chosen because they represent a simple but well-established model to study neuronal electrophysiology 11, and it is with those cells that we have obtained the most significant results to date 10. Detailed cell isolation and culture procedures have been described previously 12, 13.
<ul type="disc">
	<li>
	 Remove the outer shell from 2-3 month old Lymnaea stagnalis with blunt forceps and anesthetize animals in Lymnaea saline containing 10% Listerine. All subsequent steps performed aseptically within a laminar airflow hood with sterilized dissection equipment and solutions at room temperature.</li>
	<li>Replace physiological media in the fluidics with the appropriate recording solutionusing an Eppendorf pipette. In the case of Lymnaea neurons the solution contains (in mM): 50 KCl, 5 MgCl2, 5 ethylenebis(oxyethylenenitrilo)tetraacetic acid (EGTA) and 5 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES; pH 7.4; 130 mOsm)14</li>
	<li>Pin the snails into a Sylgard dish filled with Lymnaea saline and remove entire brain as previously described 13</li>
	<li>Treat the isolated brains in defined media (DM) with trypsin enzyme (0.2% Solution, Sigma, Catalog # T-9201) for 18 minutes followed by treatment in DM and trypsin inhibitor (Sigma, Soybean, Catalog # T-9003) for 15 minutes.</li>
	<li>Pin the brains into a small Sylgard dish filled with high osmolarity defined media (HODM - DM with glucose added; 750 &mu;L of 1M glucose solution to 20 mL DM).</li>
	<li>Using fine forceps and dissection scissors, remove the outer and inner sheath of the ganglia of interest, exposing the neurons.</li>
	<li>Using a fire polished glass pipette filled with HODM and attached to a microsyringe, apply gentle suction near the cell of interest (Left pedal dorsal 1) until the neuron detaches from the brain and is suspended in the pipette.</li>
	<li>The glass pipette is then moved and dipped into individual neurochips where gentle expulsion from the microsyringe allows the neurons to be gently pushed out of the pipette and placed on top of the individual patch holes on the chips.</li>
	<li>Allow cells to sit undisturbed for a minimum of 2h at room temperature to promote their attachment to the chip surface surrounding the aperture.</li>
</ul>
5. Electrophysiological recordings
To connect the chips to the amplifier (in our case a Multiclamp 700B amplifier, Molecular Devices, Foster City, CA, USA)
<ul type="disc">
	<li>Replace the solution in the subterranean fluidic and in the chip vial with the appropriate recording solutions. In the case of Lymnaea neurons , the upper culture chambers are filled with Lymnaea saline (in mM: 51.3 NaCl, 1.7 KCl, 4 CaCl2, and 1.5 MgCl2) buffered to pH 7.9 with HEPES.14</li>
	<li>For stability, glue the chip onto a glass slide and place it under a microscope.</li>
	<li>To connect the chip to the amplifier (in our case a Multiclamp 700B amplifier, Molecular Devices, Foster City, CA, USA), cut one end of the tubing and insert a silver wire which is connected to the head-stage.</li>
	<li>Place the reference electrode in the top vial of the chip.</li>
	<li>The chip is now ready for recording.</li>
	<li>Apply a 5 mV step with the amplifier to measure total resistance (Rt) and determine the configuration of the neurons: cell-attached (Rt &gt; 1 G&Omega;); whole-cell (Rt = 50-100 M&Omega; plus capacitive transients, indicative of membrane rupture over the aperture); or no seal (Rt &lt; 5 M&Omega;). In our last set of experiments 10, 58% percent of cells had high-resistance seals and, of those, 80% of cells showed excitable responses.</li>
</ul>
6. Representative Results
<ul type="disc">
	<li>Apply depolarizing current pulses (20 pA increment) to the neuron (in this case LPeD1).</li>
	<li>Hold the neuron at Vm close to its resting potential (~ - 60 mV).</li>
	<li>Observe the responses to these depolarizing pulses. Overshooting action potentials should be observed if the neuron is viable.</li>
	<li>Figure 4 shows a representative result which is discussed in details in 14, 15.</li>
</ul>
<img alt="Figure 4" src="/files/ftp_upload/3288/3288fig4.jpg" /> 
Figure 4. Voltage responses (top) of a LPeD1 neuron to graded series of intracellular current pulses (below). The current pulses were applied at Vm = - 60mV.
Troubleshooting
<ul type="disc">
	<li>Before plating, image chips to determine if membrane and aperture are free of debris and suitable for plating. We use an upright microscope and a 20x long working distance objective. In our last set of experiments 1, 67% percent of chips were successfully primed in that manner.</li>
	<li>If debris and/or air bubbles are present discard chip and try another one.</li>
	<li>Various surface treatments (plasma) or coatings (PDL, PEI etc.) may be tried, but success may be highly dependent upon cell type used.</li>
	<li>The composition of the fluidic ("pipette") solution in critical for the formation of the giga-seal and the whole-cell. Adjust solution, paying attention to osmolarity, pH, and ionic makeup.</li>
	<li>For longer term culture, begin with media in microfluidic channel, then switch to pipette solution prior to recording via gentle gravity fed perfusion (~0.5 ml/min).</li>
</ul>

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No conflicts of interest declared.

NRCC's patch-clamp chip interrogation platform is a potentially powerful tool for high information content pharmaceutical assays and to investigate in vitro models of disease. Its advantages compared to glass pipettes are a low access resistance, which is an advantage to probe large cells, and despite a somewhat larger capacitance will result in comparable dynamics for smaller cells. Spontaneous cell to aperture seals have been routinely obtained, and whole cell entry has been observed to be spontaneous 14. A clear difference between chips and the glass pipette method is the fact that the probe is part of the cell culture dish and is not manually brought in contact with the cell membrane. Culturing cells, possibly part of functional networks, results in more biologically relevant models as disease models, and a different mechanism for securing high cell to probe seals 16. However, by contrast with cell suspensions, aspiration cannot be used to position a cell on the probe. Snail neurons, as other large cells, are amenable to manual positioning on top of the probe. For smaller cells requiring longer culture times, we have obviated the need for any manipulation and keep a high probability of obtaining a seal by patterning adhesion polypeptides on top of the probes to place cells on top of the probes, and demonstrated placement of cells on the probe 17,18.
NRCC is also developing a polyimide microfluidic patch-clamp chip 19 with a capacitance comparable to that of glass pipette. The ultimate goal of that project is a multiple-probes patch-clamp chip which allows the simultaneous monitoring of the electrophysiological activity of several neurons engaged in network behavior at the resolution of individual ion channels 14. This method is a high resolution complementary method to multi-electrode arrays 20.

culturing and electrophysiology of cells on nrcc patch-clamp chips

Due to its exquisite sensitivity and the ability to monitor and control individual cells at the level of ion channels, patch-clamping is the gold standard of electrophysiology applied to disease models and pharmaceutical screens alike 1. The method traditionally involves gently contacting a cell with a glass pipette filled by a physiological solution in order to isolate a patch of the membrane under its apex 2. An electrode inserted in the pipette captures ion-channel activity within the membrane patch or, when ruptured, for the whole cell. In the last decade, patch-clamp chips have been proposed as an alternative 3, 4: a suspended film separates the physiological medium from the culture medium, and an aperture microfabricated in the film replaces the apex of the pipette. Patch-clamp chips have been integrated in automated systems and commercialized for high-throughput screening 5. To increase throughput, they include the fluidic delivery of cells from suspension, their positioning on the aperture by suction, and automated routines to detect cell-to-probe seals and enter into whole cell mode. We have reported on the fabrication of a silicon patch-clamp chip with optimized impedance and orifice shape that permits the high-quality recording of action potentials in cultured snail neurons 6; recently, we have also reported progress towards interrogating mammalian neurons 7. Our patch-clamp chips are fabricated at the Canadian Photonics Fabrication Centre 8, a commercial foundry, and are available in large series. We are eager to engage in collaborations with electrophysiologists to validate the use of the NRCC technology in different models. The chips are used according to the general scheme represented in Figure 1: the silicon chip is at the bottom of a Plexiglas culture vial and the back of the aperture is connected to a subterranean channel fitted with tubes at either end of the package. Cells are cultured in the vial and the cell on top of the probe is monitored by a measuring electrode inserted in the channel.The two outside fluidic ports facilitate solution exchange with minimal disturbance to the cell; this is an advantage compared to glass pipettes for intracellular perfusion.
<img alt="Figure 1" src="/files/ftp_upload/3288/3288fig1.jpg" />
Figure 1. Principle of measurement using the NRCC patch-clamp chip
We detail here the protocols to sterilize and prime the chips, load them with medium, plate them with cells, and finally use them for electrophysiological recordings.

Watch this Scientific Journal Video about Culturing and Electrophysiology of Cells on NRCC Patch-clamp Chips at JoVE.com

Culturing and Electrophysiology of Cells on NRCC Patch-clamp Chips

We show how planar patch-clamp chips fabricated at the National Research Council of Canada are sterilized, primed, loaded with medium, plated with cells, and used for electrophysiological recordings.

Due to its exquisite sensitivity and the ability to monitor and control individual cells at the level of ion channels, patch-clamping is the gold standard ...

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15.6K Views.  National Research Council of Canada. We show how planar patch-clamp chips fabricated at the National Research Council of Canada are sterilized, primed, loaded with medium, plated with cells, and used for electrophysiological recordings.

Video: Culturing and Electrophysiology of Cells on NRCC Patch-clamp Chips

Patch Clamp Recordings in Inner Ear Hair Cells Isolated from Zebrafish

Patch clamp analyses of the voltage-gated channels in sensory hair cells isolated from a variety of species have been described previously1-4 but this video represents the first application of those techniques to hair cells from zebrafish. Here we demonstrate a method to isolate healthy, intact hair cells from all of the inner ear end-organs: saccule, lagena, utricle and semicircular canals. Further, we demonstrate the diversity in hair cell size and morphology and give an example of the kinds of patch clamp recordings that can be obtained. The advantage of the use of this zebrafish model system over others stems from the availability of zebrafish mutants that affect both hearing and balance. In combination with the use of transgenic lines and other techniques that utilize genetic analysis and manipulation, the cell isolation and electrophysiological methods introduced here should facilitate greater insight into the roles hair cells play in mediating these sensory modalities.

The purpose of this video is to demonstrate procedures for obtaining healthy, intact hair cells from the inner ear organs of adult zebrafish and then using them for patch clamp studies aimed at characterizing the biophysical properties of their voltage-gated channels.

Patch  clamp analyses of the voltage-gated channels in sensory hair cells isolated  from a variety of species have been described previously1-4 but this  ...

Whole Cell Patch Clamp for Investigating the Mechanisms of Infrared Neural Stimulation

It has been demonstrated in recent years that pulsed, infrared laser light can be used to elicit electrical responses in neural tissue, independent of any further modification of the target tissue. Infrared neural stimulation has been reported in a variety of peripheral and sensory neural tissue in vivo, with particular interest shown in stimulation of neurons in the auditory nerve. However, while INS has been shown to work in these settings, the mechanism (or mechanisms) by which infrared light causes neural excitation is currently not well understood. The protocol presented here describes a whole cell patch clamp method designed to facilitate the investigation of infrared neural stimulation in cultured primary auditory neurons. By thoroughly characterizing the response of these cells to infrared laser illumination in vitro under controlled conditions, it may be possible to gain an improved understanding of the fundamental physical and biochemical processes underlying infrared neural stimulation.

Infrared nerve stimulation has been proposed as an alternative to electrical stimulation in a range of nerve types, including those associated with the auditory system. This protocol describes a patch clamp method for studying the mechanism of infrared nerve stimulation in a culture of primary auditory neurons.

It has been  demonstrated in recent years that pulsed, infrared laser light can be used to  elicit electrical responses in neural tissue, independent of ...

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

The marine gastropod mollusk Aplysia californica has a venerable history as a model of nervous system function, with particular significance in studies of learning and memory. The typical preparations for such studies are ones in which the sensory and motoneurons are left intact in a minimally dissected animal, or a technically elaborate neuronal co-culture of individual sensory and motoneurons. Less common is the isolated neuronal preparation in which small clusters of nominally homogeneous neurons are dissociated into single cells in short term culture. Such isolated cells are useful for the biophysical characterization of ion currents using patch clamp techniques, and targeted modulation of these conductances. A protocol for preparing such cultures is described. The protocol takes advantage of the easily identifiable glutamatergic sensory neurons of the pleural and buccal ganglia, and describes their dissociation and minimal maintenance in culture for several days without serum.

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

We describe the dissection of the nervous system of the marine sea hare Aplysia after anesthesia, the isolation of neurons for short term-tissue culture, and recordings of single cell ion currents via the patch clamp technique.

The marine gastropod mollusk Aplysia californica has a venerable  history as a model of nervous system function, with particular significance in  studies ...

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Mauthner cells (M-cells) are large reticulospinal neurons located in the hindbrain of teleost fish. They are key neurons involved in a characteristic behavior known as the C-start or escape response that occurs when the organism perceives a threat. The M-cell has been extensively studied in adult goldfish where it has been shown to receive a wide range of excitatory, inhibitory and neuromodulatory signals1. We have been examining M-cell activity in embryonic zebrafish in order to study aspects of synaptic development in a vertebrate preparation. In the late 1990s Ali and colleagues developed a preparation for patch clamp recording from M-cells in zebrafish embryos, in which the CNS was largely intact2,3,4. The objective at that time was to record synaptic activity from hindbrain neurons, spinal cord neurons and trunk skeletal muscle while maintaining functional synaptic connections within an intact brain-spinal cord preparation. This preparation is still used in our laboratory today. To examine the mechanisms underlying developmental synaptic plasticity, we record excitatory (AMPA and NMDA-mediated)5,6 and inhibitory (GABA and glycine) synaptic currents from developing M-cells. Importantly, this unique preparation allows us to return to the same cell (M-cell) from preparation to preparation to carefully examine synaptic plasticity and neuro-development in an embryonic organism. The benefits provided by this preparation include 1) intact, functional synaptic connections onto the M-cell, 2) relatively inexpensive preparations, 3) a large supply of readily available embryos 4) the ability to return to the same cell type (i.e. M-cell) in every preparation, so that synaptic development at the level of an individual cell can be examined from fish to fish, and 5) imaging of whole preparations due to the transparent nature of the embryos.

We have developed an intact brain-spinal cord preparation to record and monitor electrical activity via patch clamp recording from the Mauthner neurons and other reticulospinal cells in zebrafish embryos. Thus, we are able to record excitatory and inhibitory synaptic currents, voltage-gated channel activity and action potentials from key neurons in a developing embryo.

Mauthner cells (M-cells) are large  reticulospinal neurons located in the hindbrain of teleost fish. They are key  neurons involved in a characteristic ...

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.

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 ...

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Cortical networks are controlled by a small, but diverse set of inhibitory interneurons. Functional investigation of interneurons therefore requires targeted recording and rigorous identification. Described here is a combined approach involving whole-cell recordings from single or synaptically-coupled pairs of neurons with intracellular labeling, post-hoc morphological and immunocytochemical analysis.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal ...

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 ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic network, electrophysiological recordings are frequently used to study connections among individual neurons. We have optimized a flat-mount preparation for patch clamp recording of genetically marked neurons in both GCL (ganglion cell layer) and INL (inner nuclear layer) of mouse retinas. Recording INL neurons in flat-mounts is favored over slices because both vertical and lateral connections are preserved in the former configuration, allowing retinal circuits with large lateral components to be studied. We have used this procedure to compare responses of mirror-partnered neurons in retinas such as the cholinergic starburst amacrine cells (SACs).

This protocol demonstrates how to perform whole-cell patch clamp recording on retinal neurons from a flat-mount preparation.

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic ...

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.