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><tbody><tr><td>CCD Camera</td><td>QImaging</td><td>Rolera Bolt</td><td></td></tr><tr><td>Electrophysiology rig</td><td>Scientifica</td><td>SliceScope Pro 2000</td><td>Include microscope and manipulators. The manufacturer provided manipulator control software demonstrated in this manuscript is &amp;ldquo;Linlab2&amp;rdquo;.</td></tr><tr><td>Amplifier</td><td>Molecular Devices</td><td>MultiClamp 700B</td><td>computer-controlled microelectrode amplifier</td></tr><tr><td>Digitizer</td><td>Molecular Devices</td><td>Axon Digidata 1550</td><td></td></tr><tr><td>LED light source</td><td>Cool LED</td><td>pE-100</td><td>488 nm wavelength</td></tr><tr><td>Data acquisition board</td><td>Measurement Computing</td><td>USB1208-FS</td><td>Secondary DAQ.&lt;br/&gt; See manual at : http://www.mccdaq.com/pdfs/manuals/USB-1208FS.pdf</td></tr><tr><td>Solenoid valves</td><td>The Lee Co.</td><td>LHDA0531115H</td><td></td></tr><tr><td>Air pump</td><td>Virtual industry</td><td>VMP1625MX-12-90-CH</td><td></td></tr><tr><td>Air pressure sensor</td><td>Freescale semiconductor</td><td>MPXV7025G</td><td></td></tr><tr><td>Slice hold-down</td><td>Warner instruments</td><td>64-1415 (SHD-40/2)</td><td>Slice Anchor Kit, Flat for RC-40 Chamber, 2.0 mm, 19.7 mm</td></tr><tr><td>Python</td><td>Anaconda</td><td>version 2.7 (32-bit for windows)</td><td>https://www.continuum.io/downloads</td></tr><tr><td>Screw Terminals</td><td>Sparkfun</td><td>PRT - 08084</td><td>Screw Terminals 3.5 mm Pitch (2-Pin)</td></tr><tr><td>(2-Pin)</td><td></td><td></td><td></td></tr><tr><td>N-Channel MOSFET 60 V 30 A</td><td>Sparkfun</td><td>COM - 10213</td><td></td></tr><tr><td>DIP Sockets Solder Tail - 8-Pin</td><td>Sparkfun</td><td>PRT-07937</td><td></td></tr><tr><td>LED - Basic Red 5 mm</td><td>Sparkfun</td><td>COM-09590</td><td></td></tr><tr><td>LED - Basic Green 5mm</td><td>Sparkfun</td><td>COM-09592</td><td></td></tr><tr><td>DC Barrel Power Jack/Connector (SMD)</td><td>Sparkfun</td><td>PRT-12748</td><td></td></tr><tr><td>Wall Adapter Power Supply - 12 V DC 600 mA</td><td>Sparkfun</td><td>TOL-09442</td><td></td></tr><tr><td>Hook-Up Wire - Assortment (Solid Core, 22 AWG)</td><td>Sparkfun</td><td>PRT-11367</td><td></td></tr><tr><td>Locking Male x Female x Female Stopcock</td><td>ARK-PLAS</td><td>RCX10-GP0</td><td></td></tr><tr><td>Fisherbrand Tygon S3 E-3603 Flexible Tubings</td><td>Fisher scientific</td><td>14-171-129</td><td>Outer Diameter: 1/8 in.&lt;br/&gt; Inner Diameter: 1/16 in.</td></tr><tr><td>BNC male to BNC male coaxial cable</td><td>Belkin Components</td><td>F3K101-06-E</td><td></td></tr><tr><td>560 Ohm Resistor (5% tolerance)</td><td>Radioshack</td><td>2711116</td><td></td></tr><tr><td>Picospritzer</td><td>General Valve</td><td>Picospritzer II</td><td></td></tr></tbody></table>

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...

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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

11.5K Views.  Purdue University.  Dit protocol beschrijft hoe u automatische beeldgeleide patch-clamp experimenten kunt uitvoeren met behulp van een systeem dat onlangs is ontwikkeld voor standaard in vitro elektrofysiologie apparatuur. 

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

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 ...

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 ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

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

Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze motor-learning induced plasticity, we trained rats using an accelerated rotor rod task. Rats performed the task 10 times at 30-s intervals for 1 or 2 days. Performance was significantly improved on the training days compared to the first trial. We then prepared acute brain slices of the primary motor cortex (M1) in untrained and trained rats. Current-clamp analysis showed dynamic changes in resting membrane potential, spike threshold, afterhyperpolarization, and membrane resistance in layer II/III pyramidal neurons. Current injection induced many more spikes in 2-day trained rats than in untrained controls.
To analyze contextual-learning induced plasticity, we trained rats using an inhibitory avoidance (IA) task. After experiencing foot-shock in the dark side of a box, the rats learned to avoid it, staying in the lighted side. We prepared acute hippocampal slices from untrained, IA-trained, unpaired, and walk-through rats. Voltage-clamp analysis was used to sequentially record miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs) from the same CA1 neuron. We found different mean mEPSC and mIPSC amplitudes in each CA1 neuron, suggesting that each neuron had different postsynaptic strengths at its excitatory and inhibitory synapses. Moreover, compared with untrained controls, IA-trained rats had higher mEPSC and mIPSC amplitudes, with broad diversity. These results suggested that contextual learning creates postsynaptic diversity in both excitatory and inhibitory synapses at each CA1 neuron.
AMPA or GABAA receptors seemed to mediate the postsynaptic currents, since bath treatment with CNQX or bicuculline blocked the mEPSC or mIPSC events, respectively. This technique can be used to study different types of learning in other regions, such as the sensory cortex and amygdala.

The slice patch clamp technique is an effective method for analyzing learning-induced changes in the intrinsic properties and plasticity of excitatory or inhibitory synapses.

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

Evaluation of Synaptic Multiplicity Using Whole-cell Patch-clamp Electrophysiology

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic multiplicity). Synaptic multiplicity is plastic and changes throughout development and in different physiological conditions, being an important determinant for the efficacy of synaptic transmission. Here, we outline experiments for estimating the degree of multiplicity of synapses terminating onto a given postsynaptic neuron using whole-cell patch clamp electrophysiology in acute brain slices. Specifically, voltage-clamp recording is used to compare the difference between the amplitude of spontaneous excitatory postsynaptic currents (sEPSCs) and miniature excitatory postsynaptic currents (mEPSCs). The theory behind this method is that afferent inputs that exhibit multiplicity will show large, action potential-dependent sEPSCs due to the synchronous release that occurs at each synaptic contact. In contrast, action potential-independent release (which is asynchronous) will generate smaller amplitude mEPSCs. This article outlines a set of experiments and analyses to characterize the existence of synaptic multiplicity and discusses the requirements and limitations of the technique. This technique can be applied to investigate how different behavioral, pharmacological or environmental interventions in vivo affect the organization of synaptic contacts in different brain areas.

Here, we present a protocol for evaluating the functional synaptic multiplicity using whole-cell patch clamp electrophysiology in acute brain slices.

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic ...

Establishing a Whole-Cell Voltage-Clamp Configuration for Electrophysiological Recordings in Brain Slices

Source: Sunstrum, J. K., et al. <a href="https://app.jove.com/v/59461">Evaluation of synaptic multiplicity using whole-cell patch-clamp electrophysiology.</a> J. Vis. Exp. (2019)
This video demonstrates positioning a fine-pointed pipette near brain slices, forming a seal with a neuronal cell. Suction breaks the cell membrane, thereby creating a whole-cell patch clamp that enables current measurements for electrophysiology experiments.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review ...

JoVE Encyclopedia of Experiments

Neurophysiology

Electrophysiology Techniques

Automated Image-Guided Patch Clamp Technique for Studying Neurons in Brain Slices

Source: Wu, Q., et al. <a href="https://app.jove.com/v/56010">Application of automated image-guided patch clamp for the study of neurons in brain slices.</a> J. Vis. Exp. (2017).
In this video, brain slices are immobilized in a recording chamber, and cells of interest are located and focused on using a computer-controlled microscope. The automated image processing system then precisely positions a patch pipette to seal the cell membrane and monitor electrical signals.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. System Setup

	Construct ...

Paired Patch Clamp Recording of the Neuronal Connections in Acute Brain Slices

Source: Qi, G., et al. <a href="https://app.jove.com/v/52358">Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings.</a> J. Vis. Exp. (2015)
This video demonstrates the procedure to map synaptic connections by securing a brain slice, positioning pipettes to achieve whole-cell configurations, and stimulating presynaptic neurons to record postsynaptic responses, thereby elucidating neuronal connectivity.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Paired Patch-Clamp ...

Patch-Clamp Recordings from the Dendrite of a Dopaminergic Neuron in a Brain Slice

In this procedure, transfer a brain slice to the recording chamber. Anchor the slice at the bottom of the recording chamber with a platinum ring, and ensure that the slice is of good quality with a smooth, even surface. Then, select a neuron with a dendrite extending over a long distance in the same plane, and ensure that the dendrite of interest can be followed to a well-defined soma. Next, fill the patch pipette with electrode solution.
To patch the dendrite in cell-attached mode, identify and focus on a portion of the dendrite. Apply positive pressure to the pipette, and lower it using the micromanipulator. Then, position the pipette close to the membrane, and adjust the pressure in order to create a small dimple. Subsequently, release the pressure on the pipette tip and patch the dendrite while controlling the pipette resistance. Try to obtain a seal resistance larger than 1 giga ohm, at best between 3 and 10 giga ohms for cell-attached recordings.
Afterward, retract the pipette away from the membrane by a couple of micrometers to avoid deformation of the dendrite. The action currents seen here represent the spontaneous action potentials of nigral neurons. To suppress the action currents, apply calcium and sodium channel blockers to ACSF. Then, apply a voltage step of negative 90 millivolts from a holding potential of 0 millivolts to evoke the hyperpolarization-activated cation current.

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

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