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>

<ol>
	<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. S., Forest, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+whole-cell+patch-clamp+electrophysiology+of+neurons+in+vivo.">Automated whole-cell patch-clamp electrophysiology of neurons in vivo.</a> Nat. Methods. 9, 585-587 (2012).</li><li>Anastassiou, C. A., Perin, R., Markram, H., Koch, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ephaptic+coupling+of+cortical+neurons.">Ephaptic coupling of cortical neurons.</a> Nat. Neurosci. 14, 217-223 (2011).</li><li>Prakash, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+optogenetic+toolbox+for+fast+inhibition,+excitation+and+bistable+modulation.">Two-photon optogenetic toolbox for fast inhibition, excitation and bistable modulation.</a> Nat. Methods. 9, 1171-1179 (2012).</li><li>Papagiakoumou, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanless+two-photon+excitation+of+channelrhodopsin-2.">Scanless two-photon excitation of channelrhodopsin-2.</a> Nat Methods. 7, 848-854 (2010).</li><li>Ko, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+specificity+of+local+synaptic+connections+in+neocortical+networks.">Functional specificity of local synaptic connections in neocortical networks.</a> Nature. 473, 87-91 (2011).</li><li>Wickersham, I. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monosynaptic+Restriction+of+Transsynaptic+Tracing+from+Single,+Genetically+Targeted+Neurons.">Monosynaptic Restriction of Transsynaptic Tracing from Single, Genetically Targeted Neurons.</a> Neuron. 53, 639-647 (2007).</li><li>Liang, C. W., Mohammadi, M., Santos, M. D., Tang, C. -. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterned+Photostimulation+with+Digital+Micromirror+Devices+to+Investigate+Dendritic+Integration+Across+Branch+Points.">Patterned Photostimulation with Digital Micromirror Devices to Investigate Dendritic Integration Across Branch Points.</a> J. Vis. Exp. (49), e2003 (2011).</li><li>Nikolenko, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SLM+Microscopy:+Scanless+Two-Photon+Imaging+and+Photostimulation+with+Spatial+Light+Modulators.">SLM Microscopy: Scanless Two-Photon Imaging and Photostimulation with Spatial Light Modulators.</a> Front Neural Circuits. 2, (2008).</li></ol>

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 border="1">
	<tbody>
		<tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td>BX51WI</td>
			<td>40X Immersion Objective</td>
		</tr>
		<tr>
			<td>Manipulators</td>
			<td>Luigs &#38; 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...

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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 ...

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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

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 ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

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.

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 ...

F1FO ATPase Vesicle Preparation and Technique for Performing Patch Clamp Recordings of Submitochondrial Vesicle Membranes

Mitochondria are involved in many important cellular functions including metabolism, survival1, development and, calcium signaling2. Two of the most important mitochondrial functions are related to the efficient production of ATP, the energy currency of the cell, by oxidative phosphorylation, and the mediation of signals for programmed cell death3.
The enzyme primarily responsible for the production of ATP is the F1FO-ATP synthase, also called ATP synthase4-5. In recent years, the role of mitochondria in apoptotic and necrotic cell death has received considerable attention. In apoptotic cell death, BCL-2 family proteins such as Bax enter the mitochondrial outer membrane, oligomerize and permeabilize the outer membrane, releasing pro-apoptotic factors into the cytosol6. In classic necrotic cell death, such as that produced by ischemia or excitotoxicity in neurons, a large, poorly regulated increase in matrix calcium contributes to the opening of an inner membrane pore, the mitochondrial permeability transition pore or mPTP. This depolarizes the inner membrane and causes osmotic shifts, contributing to outer membrane rupture, release of pro-apoptotic factors, and metabolic dysfunction. Many proteins including Bcl-xL7 interact with F1FO ATP synthase, modulating its function. Bcl-xL interacts directly with the beta subunit of F1FO ATP synthase, and this interaction decreases a leak conductance within the F1FOATPasecomplex, increasing the net transport of H+ by F1FO during F1FO ATPase activity8 and thereby increasing mitochondrial efficiency. To study the activity and modulation of the ATP synthase, we isolated from rodent brain submitochondrial vesicles (SMVs) containing F1FO ATPase. The SMVs retain the structural and functional integrity of the F1FO ATPase as shown in Alavian et al. Here, we describe a method that we have used successfully for the isolation of SMVs from rat brain and we delineate the patch clamp technique to analyze channel activity (ion leak conductance) of the SMVs.

F1FO ATPase Vesicle Preparation and Technique for Performing Patch Clamp Recordings of Submitochondrial Vesicle Membranes

A method to isolate submitochondrial vesicles enriched in F1FO ATP synthase complexes from rat brain is described. These vesicles allow the study of the activity of F1FO ATPase complex and its modulation using the technique of patch clamp recording.

Mitochondria are involved in many important cellular  functions including metabolism, survival1, development and, calcium signaling2. Two of the most ...

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 ...

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 ...

Quantifying Fish Swimming Behavior in Response to Acute Exposure of Aqueous Copper Using Computer Assisted Video and Digital Image Analysis

Behavioral responses of aquatic organisms to environmental contaminants can be precursors of other effects such as survival, growth, or reproduction. However, these responses may be subtle, and measurement can be challenging. Using juvenile white sturgeon (Acipenser transmontanus) with copper exposures, this paper illustrates techniques used for quantifying behavioral responses using computer assisted video and digital image analysis. In previous studies severe impairments in swimming behavior were observed among early life stage white sturgeon during acute and chronic exposures to copper. Sturgeon behavior was rapidly impaired and to the extent that survival in the field would be jeopardized, as fish would be swept downstream, or readily captured by predators. The objectives of this investigation were to illustrate protocols to quantify swimming activity during a series of acute copper exposures to determine time to effect during early lifestage development, and to understand the significance of these responses relative to survival of these vulnerable early lifestage fish. With mortality being on a time continuum, determining when copper first affects swimming ability helps us to understand the implications for population level effects. The techniques used are readily adaptable to experimental designs with other organisms and stressors.

Measuring the impacts of environmental contaminants on fish behavior is often subjective and challenging particularly when dealing with sublethal endpoints. We describe methods including video technology to quantify swimming behavior of early life stage white sturgeon (Acipenser transmontanus) during and after 96 hr acute exposures to various concentrations of copper.

Behavioral responses of aquatic organisms to environmental contaminants can be precursors of other effects such as survival, growth, or reproduction.  ...

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 ...