We are grateful for the financial support from the Whitehall Foundation. We would like to thank Samuel T. Kissinger for the valuable comments.

1. System Setup
<ol>
	<li>Construct the pressure control unit.
	<ol>
		<li>Assemble the pressure control unit according to the circuit map (Figure 1). Solder the necessary parts onto the Printed Circuit Board (PCB) manufactured according to the electrical circuit schematics (Figure 1b). Use standard resistors, LEDs, Metal-Oxide Semiconductor Field-Eeffect Transistors (MOSFETs), capacitors, and connectors (see the Table of Materials). Solder solenoid valves onto the PCB. Connect the air pump and air pressure sensor to the PCB with electrical wire. 
		NOTE: It should take about 2 h to construct the pressure control unit with all necessary parts made available.</li>
	</ol>
	</li>
	<li>Connect the secondary data acquisition (DAQ) board.
	<ol>
		<li>Connect data outputs from the printed circuit board to the DAQ board, following Table 1. 
		NOTE: The DAQ board will be running in &#34;Single-ended mode.&#34; The port map can be found in the user manual (see the Table of Materials).</li>
		<li>Connect &#34;AIn Pr S&#34; to one of the analog input (AI) channels and &#34;R-Gr&#34; to one of the analog grounds on the secondary DAQ board.</li>
		<li>Connect the primary output from the amplifier to one of the AI channels and the ground to the analog ground of the secondary DAQ board. 
		NOTE: A standard BNC cable can be used to connect the primary output from the amplifier.</li>
		<li>Strip the other end and connect the positive signal (i.e.&#160;copper core) to the AI channel and the ground (i.e.&#160;the thin wire around the core) to the analog ground. Repeat this step for a second channel if more than one patch channel is used. 
		NOTE: The analog input to the DAQ board will be configured in later steps.</li>
		<li>Connect power to the power output of the secondary DAQ board. Use a separate 12 V power source for the pump.</li>
	</ol>
	</li>
	<li>Connect the tubing.
	<ol>
		<li>Connect the air pump and the two valves according to Table 2. Use a 3-way connector to connect the soft tubing from the valve 2 top port, the pressure sensor, and the pipette holder in the last step.</li>
		<li>Add another 3-way connector to the tubing connected to the pipette holder if two pipettes are used. Manually switch between the valves and the pipettes in use when patching.</li>
	</ol>
	</li>
	<li>Install Autopatcher IG. 
	NOTE: System requirement: Autopatcher IG was only tested on a PC running Windows 7. It has not been validated for other operating systems. The described procedure applies specifically to the hardware listed in the Table of Materials.
	<ol>
		<li>Download Autopatcher-IG from GitHub (https://github.com/chubykin/AutoPatcher_IG).</li>
		<li>Install Python (see the Table of Materials for the version and download address).</li>
		<li>Uninstall the PyQt4 library by typing &#34;pip uninstall PyQt4&#34; in a command line terminal. 
		NOTE: The system uses an older version of the PyQt4 library to achieve compatibility with the Qwt and Opencv libraries.</li>
		<li>Install Python libraries from historic wheel files (http://www.lfd.uci.edu/~gohlke/pythonlibs/). Find the following files: Numpy (pymc-2.3.6-cp27-cp27m-win32.whl), Opencv (opencv_python-2.4.13.2-cp27-cp27m-win32.whl), Pyqt (PyQt4-4.11.4-cp27-none-win32.whl), and Qwt (PyQwt-5.2.1-cp27-none-win32.whl).
		<ol>
			<li>To install the wheel files, go to the directory where the files are saved and type &#34;pip install ***wheelfilename***.whl.&#34; Substitute &#34;***wheelfilename***&#34; with the actual name of the file. 
			NOTE: &#34;cp27&#34; in the wheel file name indicates Python 2.7 and &#34;win32&#34; indicated Windows 32-bit. If &#34;win32&#34; does not work, try &#34;win64.&#34;</li>
		</ol>
		</li>
		<li>To control the CCD camera, download and install the installer for 64-bit (https://www.qimaging.com/support/software/). Then download MicroManager for 64-bit (https://micro-manager.org/wiki/Download_Micro-Manager_Latest_Release) to control the camera in Python.</li>
		<li>To control the manipulators and the microscope stage, install control software provided by the manufacturer. 
		NOTE: By doing this, the driver necessary to control the manipulators is also installed. The installation package is commonly provided in a CD-ROM.</li>
		<li>To control the secondary DAQ board, install the Universal Library from CD-ROM, provided with the purchase of the DAQ board.</li>
	</ol>
	</li>
	<li>Configure the hardware for Autopatcher IG.
	<ol>
		<li>Connect the microscope stage and manipulator controllers to the computer via USB ports.</li>
		<li>Assign COM port numbers to unit 0: microscope stage, unit 1: left manipulator, and unit 2: right manipulator, in this order, in the &#34;ports.csv&#34; configuration file in the &#34;configuration&#34; folder. Leave the other parameters in the ports.csv file (i.e.&#160;&#34;SCI&#34; and &#34;1&#34;) unchanged. 
		NOTE: The COM port number information can be found by running the manipulator configuration software provided by the manufacturer. Go to the &#34;settings&#34; tab, select &#34;settings&#34; and the &#34;Motion&#34; page, and read the labels for each tab at the top. Alternatively, this information can be found in the PC Device Manager.</li>
		<li>Assign analog input channel numbers on the DAQ board for a pressure sensor and patch channel 1 and 2 (corresponding to unit 1 and 2). Enter the channel number in the &#34;DAQchannels.csv&#34; file in the &#34;configuration&#34; folder. 
		NOTE: It is recommended to open the .csv files with the Notepad application instead of a spreadsheet, as it may alter the information when saving changes.</li>
	</ol>
	</li>
	<li>Run Autopatcher IG.
	<ol>
		<li>Turn on the amplifier, microscope controller, and manipulator controller. Ensure that the amplifier software is running.</li>
		<li>Run Autopatcher IG with Python from a command line terminal as follows: first, change the directory (command &#34;cd&#34; for most common terminals) where Autopatcher IG is installed, type &#34;python Autopatcher_IG.pyw&#34; in the command line terminal, and hit the &#34;Enter&#34; key. 
		NOTE: Do not run the manipulator control software before running Autopatcher IG because it will occupy the microscope stage and manipulator, causing Autopatcher IG to be unable to find the hardware. Manipulator control software can be run after Autopatcher IG is fully initiated if there are additional modules to be controlled (e.g., the inline heater).</li>
	</ol>
	</li>
	<li>Calibrate the primary pipette.
	<ol>
		<li>Pull patch pipettes as described previously9. Fill a pulled glass pipette with internal solution and load it onto the head stage. 
		NOTE: Empty glass pipettes have different contrasts under the microscope and may lead to inaccurate calibration.</li>
		<li>Move the pipette tip to the microscope visual field and bring it into focus. If the dial pad is used to move the manipulators and/or microscope stage, update the coordinates by pressing &#34;z&#34; on the keyboard. 
		NOTE: This action is not necessary if the keyboard (microscope stage: A/D - x-axis, W/S - y-axis, R/F - z-axis; manipulators: H/K - x-axis, U/J - y-axis, O/L - z-axis, 1/2 - unit number) is used to control movement because the coordinates will be updated in real time.</li>
		<li>Click the &#34;Start calibration&#34; button on the main Graphic User Interface (GUI) for the corresponding unit on which the pipette is loaded (Figure 2). 
		NOTE: A pop-up window will appear when the calibration is finished. 
		NOTE: Calibration will be carried out automatically, which will take about 3.5 min. Clicking on the same button (switched now to &#34;cancel calibration&#34; after initiating calibration) will abort the calibration attempt.</li>
		<li>Save the calibration by clicking &#34;save calibration&#34; at the bottom of the main GUI (it saves the current calibration for both manipulators and can be loaded in the future). 
		NOTE: The field of view under low (4 or&#160;10X) and high (40X) magnification must be aligned for secondary calibration to function properly. Please refer to the user manual of the optical system in use for the alignment procedures.</li>
	</ol>
	</li>
</ol>
2. Automatic Patch Clamp Procedure
<ol>
	<li>Prepare acute brain slices, as described previously10.</li>
	<li>Prepare glass pipettes for the patch clamp.</li>
	<li>Place one brain slice in the center of the recording chamber. Stabilize the slice with a slice hold-down, or &#34;harp.&#34;</li>
	<li>Detect the fluorescent cell.
	<ol>
		<li>Find the area of interest under the 4X lens. Move the microscope stage by turning on click-to-move (&#34;CTM&#34;) mode and click the center of the area of interest. Alternatively, use the keypad to move the microscope stage (A/D - x-axis, W/S - y-axis, R/F - z-axis).</li>
		<li>Switch to the high-magnification lens and adjust the focus by moving the microscope in the z-axis, using R/F on the keypad. 
		NOTE: It is recommended to adjust the water bath level so that the focal plane under the low- and high-magnification lenses are the same or similar in the z-axis.</li>
		<li>Click the &#34;Detect Cell&#34; button on the main GUI column, &#34;Unit 0.&#34; If the LED or laser light source of the setup cannot be controlled by the TTL signal, manually turn on the LED/laser; a pop-up window will appear when the cell detection is finished.
		<ol>
			<li>Turn off the LED/laser if necessary; a list of cell coordinates will appear in the &#34;Memory positions&#34; GUI. Remove undesired cells from the list by clicking the &#34;X&#34; button next to the coordinates.</li>
			<li>Alternatively, if target cells are not fluorescently labeled, click &#34;Mouse mode&#34; on the main GUI. Click on the cell of interest; a yellow dot with a number will appear on the cell, and the coordinates of the cell will appear in the &#34;Memory positions&#34; GUI.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Calibrate the secondary pipette.
	<ol>
		<li>Fill 1/3 of a glass pipette with internal solution. Load the pipette onto the pipette holder attached to the head stage.</li>
		<li>Use the low-magnification lens. Bring the pipette into the visual field and adjust the focus using the keypad (H/K - x-axis, U/J - y-axis, O/L - z-axis). Use &#34;1&#34; and &#34;2&#34; to switch between unit 1 and unit 2.</li>
		<li>Load the primary calibration by clicking on &#34;Load calibration.&#34; Click the &#34;Secondary calibration&#34; button on the main GUI under the unit that is in use. For example, if unit 2 is in use, click the &#34;Secondary calibration&#34; button in the &#34;Unit 2&#34; column. Follow the pop-up window instructions to switch to the high-magnification lens.</li>
	</ol>
	</li>
	<li>Patch a target cell.
	<ol>
		<li>Make sure that the &#34;MultiClamp&#34; (i.e.&#160;amplifier) software is running. Click on the &#34;Patch control&#34; button to open the &#34;Patch control&#34; GUI; it may take up to a few min to open this GUI.</li>
		<li>Use the 40X magnification lens by checking &#34;40X&#34; on the main GUI &#34;Unit 0&#34; column. Click the &#34;go to&#34; button next to the cell of interest on the coordinate list in the &#34;Memory position&#34; GUI; the microscope will move to the cell.</li>
		<li>Click on the CTM button of the unit in use in the main GUI to enable movement following a mouse click. Click on the cell of interest; the pipette tip will move to the cell.</li>
		<li>Click on the &#34;Patch&#34; button on the &#34;Patch control&#34; GUI. 
		NOTE: Automatic patching will begin, and the pressure and resistance can be monitored on the &#34;Patch control&#34; GUI.
		<ol>
			<li>Use the &#34;Unit 1 selected&#34; button to switch the input signal between the two units. 
			NOTE: The system will approach the target cell, apply negative pressure, match the cell membrane potential, and detect gigaseal formation based on a series of pressure and resistance thresholds and logic.</li>
			<li>Manipulate the automatic process at any point by clicking on the respective buttons on the &#34;Patch control&#34; GUI. For example, click on the &#34;Patch&#34; button again to cancel the patch trial and click on &#34;Next stage&#34; to advance the patching process to the next step, regardless of the threshold. 
			NOTE: A pop-up window will notify the user when a gigaseal has formed, and the option to apply zap along with large negative pressure will be presented.</li>
		</ol>
		</li>
		<li>Select &#34;Yes&#34; to break in with combined zap and suction. Alternatively, select &#34;No&#34; to break in with suction only. 
		NOTE: When a successful whole-cell patch is completed, a pop-up window will remind the user to save the experiment patch log.</li>
		<li>Save the experiment patch log. 
		NOTE: If a patching trial is unsuccessful, a pop-up window will notify the user, and the patch process will reset.</li>
		<li>Go back to step 2.4 and repeat the steps with a different cell.</li>
	</ol>
	</li>
	<li>Refine the patching thresholds (optional). 
	NOTE: Thresholds for the initial pipette resistance range, positive and negative pressure, gigaseal resistance, etc. can be modified from a configuration file.
	<ol>
		<li>Open the &#34;PatchControlConfiguration.csv&#34; file in the &#34;Configuration&#34; folder at the destination where the system is installed. Change the numbers corresponding to each threshold value. Do not change the names of the values; this will result in unrecognizable entries in the system.</li>
		<li>Implement the new threshold values immediately by pressing &#34;Ctrl+L&#34; without restarting the program; restarting the program will implement the newest threshold value from the file.</li>
	</ol>
	</li>
</ol>
3. Performing Recordings
NOTE: The mode in the computer-controlled microelectrode amplifier will be set automatically to Current Clamp (&#34;IC&#34;) by the autopatcher software once a successful patch has been achieved. Whole-cell patch clamp recordings can be done using the recording software of choice (this system does not include a recording function). If multiple target cells were identified, after finishing a recording, go back to step 2.4 and try another cell.
<ol>
	<li>Perform automatic local drug application experiments using a picospritzer (optional). 
	NOTE: Here, a local drug application experiment is used as an example to describe how to use the additional &#34;Command Sequence&#34; function to control external hardware through TTL signals.
	<ol>
		<li>Connect port A channel 0 and the ground on the secondary DAQ board to the start trigger input on the digitizer using a stripped BNC cable (as described in step 1.2.3). Connect one digital output channel on the digitizer to the external trigger on the picospritzer.</li>
		<li>Prepare the picospritzer according to the user manual. Connect the picospritzer air output to the drug-puff pipette holder attached to a micromanipulator.</li>
		<li>Load a pipette filled with a drug of choice. Attach it to the pipette holder. Calibrate the pipette, as described in step 1.7.</li>
		<li>After patching a cell as described in step 2.6, select a desired location for drug delivery by mouse-clicking in the camera view GUI under &#34;mouse mode&#34; (switched from the main GUI). Alternatively, use the &#34;Grid&#34; GUI to design a grid, with each pixel in the grid as one of the target locations. 
		NOTE: The grid can be manipulated in the camera view GUI by dragging with the mouse.</li>
		<li>In the &#34;Command Sequence&#34; GUI, select the manipulator unit that the drug pipette is mounted on. Click &#34;load mouse points&#34; or &#34;load grid points&#34; to import all coordinates for drug delivery.
		<ol>
			<li>Click on each coordinate entry to edit the specific command TTL signal in the right column. In the first command entry, click on the right-most digit to turn it from &#34;0&#34; to &#34;1,&#34; sending a +5-V TTL signal. Set the time (T) to the desired TTL signal duration, in ms.</li>
			<li>In the second command entry, set all digits to &#34;0&#34; and set T to the duration of the recording length of the trial. Edit commands for all coordinate entries. Add command entries by clicking &#34;+&#34; if multiple TTL signals are necessary. 
			NOTE: The 8-digit bits represent port A channels 0-7 on the secondary DAQ (ports 1 - 3 are occupied by the pump and two valves) can be switched, if necessary.</li>
		</ol>
		</li>
		<li>Create a recording protocol in the data acquisition module so that the start of a sweep is triggered by the external start trigger. Edit the protocol to deliver the drug at the desired time.</li>
		<li>In the &#34;Command Sequence&#34; GUI, click &#34;Run&#34; on the left to run all coordinates. Alternatively, click &#34;Run&#34; on the right to run only the selected sequence. 
		NOTE: The pipette will move to each coordinate and execute the TTL signal as defined to start the recording sweep.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Sakmann, B., Neher, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patch+clamp+techniques+for+studying+ionic+channels+in+excitable+membranes.">Patch clamp techniques for studying ionic channels in excitable membranes.</a> Annu Rev Physiol. 46, 455-472 (1984).</li><li>Collins, M. D., Gordon, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+liposome+preparation+for+imaging+and+patch-clamp+electrophysiology.">Giant liposome preparation for imaging and patch-clamp electrophysiology.</a> J Vis Exp. (76), (2013).</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 (6), 585-587 (2012).</li><li>Desai, N. S., Siegel, J. J., Taylor, W., Chitwood, R. A., Johnston, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MATLAB-based+automated+patch-clamp+system+for+awake+behaving+mice.">MATLAB-based automated patch-clamp system for awake behaving mice.</a> J Neurophysiol. 114 (2), 1331-1345 (2015).</li><li>Kodandaramaiah, S. B., 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=Assembly+and+operation+of+the+autopatcher+for+automated+intracellular+neural+recording+in+vivo.">Assembly and operation of the autopatcher for automated intracellular neural recording in vivo.</a> Nat Protocols. 11 (4), 634-654 (2016).</li><li>Wu, Q., 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=Integration+of+autopatching+with+automated+pipette+and+cell+detection+in+vitro.">Integration of autopatching with automated pipette and cell detection in vitro.</a> J Neurophysiol. 116 (4), 1564-1578 (2016).</li><li>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=A+computer-assisted+multi-electrode+patch-clamp+system.">A computer-assisted multi-electrode patch-clamp system.</a> J Vis Exp. (80), e50630 (2013).</li><li>Fertig, N., Blick, R. H., Behrends, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole+cell+patch+clamp+recording+performed+on+a+planar+glass+chip.">Whole cell patch clamp recording performed on a planar glass chip.</a> Biophys J. 82 (6), 3056-3062 (2002).</li><li>Brown, A. L., Johnson, B. E., Goodman, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Making+patch-pipettes+and+sharp+electrodes+with+a+programmable+puller.">Making patch-pipettes and sharp electrodes with a programmable puller.</a> J Vis Exp. (20), (2008).</li><li>Segev, A., Garcia-Oscos, F., Kourrich, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-cell+Patch-clamp+Recordings+in+Brain+Slices.">Whole-cell Patch-clamp Recordings in Brain Slices.</a> J Vis Exp. (112), (2016).</li><li>Campagnola, L., Kratz, M. B., Manis, P. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ACQ4:+an+open-source+software+platform+for+data+acquisition+and+analysis+in+neurophysiology+research.">ACQ4: an open-source software platform for data acquisition and analysis in neurophysiology research.</a> Front Neuroinform. 8 (3), (2014).</li><li>Kolb, I., 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=Cleaning+patch-clamp+pipettes+for+immediate+reuse.">Cleaning patch-clamp pipettes for immediate reuse.</a> Sci Rep. 6, (2016).</li></ol>

A non-provisional patent application &#34;SYSTEMS AND METHODS FOR AUTOMATED IMAGE-GUIDED PATCH-CLAMP ELECTROPHYSIOLOGY IN VITRO,&#34; U.S. Serial No.: 15/353,719, was filed on November 16, 2016, Ref. No.: PRF 67270-02.

Here, we describe a method for automatic image-guided patch clamp recordings in vitro. The key steps in this process are summarized as follows. First, computer vision is used to automatically recognize the pipette tip using a series of images acquired via a microscope. This information is then used to calculate the coordinate transformation function between the microscope and the manipulator coordinate systems. Computer vision is used to automatically detect fluorescently labeled cells and to identify their coor...

The patch clamp technique was first developed by Neher and Sakmann in the 1970s to study the ionic channels of excitable membranes1. Since then, patch clamping has been applied to the study of many different subjects at the cellular, synaptic, and circuit level&#8212;both in vitro and in vivo&#8212;in many different cell types, including neurons, cardiomyocytes, Xenopus oocytes, and artificial liposomes2. This process involves the correct identification and targeting of a cell of interest, intricate micromanipulator control to move the patch pipette in close proximity to the cell, the application of positive and negative pressure to the pipette at the proper time to establish a tight gigaseal patch, and a break-in to establish a whole-cell patch configuration. Patch clamping is typically conducted manually and requires extensive training to master. Even for a researcher experienced with the patch clamp, the success rate is relatively low. More recently, several attempts have been made to automate patch-clamp experiments. Two main strategies have evolved to accomplish automation: augmenting standard patch clamp equipment to provide automatic control of the patching process and the design of new equipment and techniques from the ground up. The former strategy is adaptable to existing hardware and can be used in a variety of patch clamp applications, including in vivo blind patch clamp3,4,5, in vitro patch clamp of acute brain slices, organotypic slice cultures, and cultured dissociated neurons6. It enables the interrogation of complex local circuits by using multiple micromanipulators simultaneously7. The planar patch method is an example of the new development strategy, which can achieve the high-throughput simultaneous patch clamp of cells in suspension for drug screening purposes8. However, the planar patch method is not applicable to all cell types, particularly neurons with long processes or intact circuits containing extensive connections. This limits its application to mapping the intricate circuitry of the nervous system, which is a key advantage of traditional patch clamp technology.
We have developed a system that automates the manual patch clamp process in vitro by augmenting standard patch clamp hardware. Our system, Autopatcher IG, provides automatic pipette calibration, fluorescent cell target identification, automatic control of pipette movement, automatic whole-cell patching, and data logging. The system can automatically acquire multiple images of brain slices at different depths; analyze them using computer vision; and extract information, including the coordinates of fluorescently labeled cells. This information can then be used to target and automatically patch cells of interest. The software is written in Python&#8212;a free, open-source programming language&#8212;using several open-source libraries. This ensures its accessibility to other researchers and improves the reproducibility and rigor of electrophysiology experiments. The system has a modular design, such that additional hardware can easily be interfaced with the current system demonstrated here.

<table width="747">
	<tbody>
		<tr height="21">
			<td height="21" style="width:231px;height:21px;">CCD Camera</td>
			<td style="width:115px;">QImaging</td>
			<td style="width:147px;">Rolera Bolt</td>
			<td style="width:256px;"></td>
		</tr>
		<tr height="101">
			<td height="101" style="width:231px;height:101px;">Electrophysiology rig</td>
			<td style="width:115px;">Scientifica</td>
			<td style="width:147px;">SliceScope Pro 2000</td>
			<td style="width:256px;">Include microscope and manipulators. The manufacturer provided manipulator control software demonstrated in this manuscript is &#8220;Linlab2&#8221;.</td>
		</tr>
		<tr height="41">
			<td height="41" style="width:231px;height:41px;">Amplifier</td>
			<td style="width:115px;">Molecular Devices</td>
			<td style="width:147px;">MultiClamp 700B</td>
			<td style="width:256px;">computer-controlled microelectrode amplifier</td>
		</tr>
		<tr height="41">
			<td height="41" style="width:231px;height:41px;">Digitizer</td>
			<td style="width:115px;">Molecular Devices</td>
			<td style="width:147px;">Axon Digidata 1550</td>
			<td style="width:256px;"></td>
		</tr>
		<tr height="55">
			<td height="55" style="width:231px;height:55px;">LED light source</td>
			<td style="width:115px;">Cool LED</td>
			<td style="width:147px;">pE-100</td>
			<td style="width:256px;">488nm wavelength</td>
		</tr>
		<tr height="98">
			<td height="98" style="width:231px;height:98px;">Data acquisition board</td>
			<td style="width:115px;">Measurement Computing</td>
			<td style="width:147px;">USB1208-FS</td>
			<td style="width:256px;">Secondary DAQ. 
			See manual at : http://www.mccdaq.com/pdfs/manuals/USB-1208FS.pdf</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:231px;height:21px;">Solenoid valves</td>
			<td style="width:115px;">The Lee Co.</td>
			<td style="width:147px;">LHDA0531115H</td>
			<td style="width:256px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="width:231px;height:41px;">Air pump</td>
			<td style="width:115px;">Virtual industry</td>
			<td style="width:147px;">VMP1625MX-12-90-CH</td>
			<td style="width:256px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="width:231px;height:41px;">Air pressure sensor</td>
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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.

Our system has been tested on its ability to patch cells in acute brain slices, mouse induced Pluripotent Stem Cells (iPSCs) differentiated into neurons, and HEK 293 cells artificially expressing channels of interest. Figure 3 shows an experiment using Thy1-ChR2-YFP transgenic mice (B6.Cg-Tg(Thy1-COP4/EYFP)18Gfng/J) targeting fluorescently labeled layer 5 pyramidal neurons in the visual cortex. The target cell was one of the automatically identified green flu...

Watch this Scientific Journal Video about Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices at JoVE.com

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

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 ...

application-automated-image-guided-patch-clamp-for-study-neurons

Research

JoVE Journal

Neuroscience

11.4K Views.  Purdue University. This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.

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

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 ...

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.

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.

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

Local Application of Drugs to Study Nicotinic Acetylcholine Receptor Function in Mouse Brain Slices

Tobacco use leads to numerous health problems, including cancer, heart disease, emphysema, and stroke. Addiction to cigarette smoking is a prevalent neuropsychiatric disorder that stems from the biophysical and cellular actions of nicotine on nicotinic acetylcholine receptors (nAChRs) throughout the central nervous system. Understanding the various nAChR subtypes that exist in brain areas relevant to nicotine addiction is a major priority.
Experiments that employ electrophysiology techniques such as whole-cell patch clamp or two-electrode voltage clamp recordings are useful for pharmacological characterization of nAChRs of interest. Cells expressing nAChRs, such as mammalian tissue culture cells or Xenopus laevis oocytes, are physically isolated and are therefore easily studied using the tools of modern pharmacology. Much progress has been made using these techniques, particularly when the target receptor was already known and ectopic expression was easily achieved. Often, however, it is necessary to study nAChRs in their native environment: in neurons within brain slices acutely harvested from laboratory mice or rats. For example, mice expressing "hypersensitive" nAChR subunits such as &alpha;4 L9&prime;A mice 1 and &alpha;6 L9&prime;S mice 2, allow for unambiguous identification of neurons based on their functional expression of a specific nAChR subunit. Although whole-cell patch clamp recordings from neurons in brain slices is routinely done by the skilled electrophysiologist, it is challenging to locally apply drugs such as acetylcholine or nicotine to the recorded cell within a brain slice. Dilution of drugs into the superfusate (bath application) is not rapidly reversible, and U-tube systems are not easily adapted to work with brain slices.
In this paper, we describe a method for rapidly applying nAChR-activating drugs to neurons recorded in adult mouse brain slices. Standard whole-cell recordings are made from neurons in slices, and a second micropipette filled with a drug of interest is maneuvered into position near the recorded cell. An injection of pressurized air or inert nitrogen into the drug-filled pipette causes a small amount of drug solution to be ejected from the pipette onto the recorded cell. Using this method, nAChR-mediated currents are able to be resolved with millisecond accuracy. Drug application times can easily be varied, and the drug-filled pipette can be retracted and replaced with a new pipette, allowing for concentration-response curves to be created for a single neuron. Although described in the context of nAChR neurobiology, this technique should be useful for studying many types of ligand-gated ion channels or receptors in neurons from brain slices.

In this paper, we describe a useful method to study ligand-gated ion channel function in neurons of acutely isolated brain slices. This method involves the use of a drug-filled micropipette for local application of drugs to neurons recorded using standard patch clamp techniques.

Tobacco use leads to numerous health problems, including cancer, heart disease, emphysema, and stroke. Addiction to cigarette smoking is a prevalent ...

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 ...

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 ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

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 ...

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 ...

Whole-cell Currents Induced by Puff Application of GABA in Brain Slices

Pharmacological administration is commonly used when conducting whole-cell patch-clamp recording in brain slices. One of the best methods of drug application during electrophysiological recording is the puff technique, which can be used to study the effect of pharmacological reagents on neuronal activities in brain slices. The greatest advantage of puff application is that the drug concentration around the recording site increases rapidly, thus preventing desensitization of membrane receptors. Successful use of puff application involves careful attention to the following elements: the concentration of the drug, the parameters of the puff micropipette, the distance between the tip of a puff micropipette and the neuron recorded, and the duration and pressure driving the puff (pounds per square inch, psi). This article describes a step-by-step procedure for recording whole-cell currents induced by puffing gamma-aminobutyric acid (GABA) onto a neuron of a prefrontal cortical slice. Notably, the same procedure can be applied with minor modifications to other brain areas such as the hippocampus and the striatum, and to different preparations, such as cell cultures.

We describe the puff technique, by which pharmacological reagents can be administered during whole-cell patch-clamp recording, and highlight various aspects of the features that are crucial for its success.