Neuroscience
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Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices
Chapters
Summary July 31st, 2017
This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.
Transcript
The overall goal of this technology is to use computer vision to automatically detect fluorescent cells and perform automated patch clamp of these cells in acute brain slices. This method can help overcome key difficulties in the patch clamp electrophysiology field, such as identification, targeting, and patch clamp of fluorescently-labeled cells. The main advantage of this technique is that it can automatically detect patch pipets, fluorescent cells, and integrate the steps with automatic patch clamp.
This innovation significantly increases the experimental throughput. The implications of this technique extend towards therapy or diagnosis of neurological disorders, because high throughput image-guided automatic patch clamp can be used for pharmacological screenings in more physiological neuronal preparations. This method can also be applied to other in vitro preparations, such as neuronal organelle typic, or slice cultures, dissociate neurons as well as any non-neuronal cells.
To begin this procedure, turn on the amplifier, microscope controller, and the manipulator controller. Then, run Autopatcher IG with python from a command line terminal by first changing the directory where Autopatcher IG is installed. Next, type Python Autopatcher_IG.
PYW in the command line terminal and hit the enter key. After that, fill up all the glass pipet with internal solution and load it on to the head stage. Move the pipet tip to the microscope visual field and bring it into focus.
If the dial pad is used to move the manipulators in the microscope stage, update the coordinates by pressing Z on the keyboard. The primary calibration converts the angle and distance of movement by the manipulator to the microscope coordinate system. The manipulator travels in three directions with preset distance and the software detects the coordinates of the pipet tip within the microscope visual field.
Click the start calibration button on the main GUI for the corresponding unit on which the pipet is loaded. Subsequently, save the calibration by clicking save calibration at the bottom of the main GUI. This primary calibration is only necessary to be performed once unless the physical organization of the rig is altered.
In this step, place one brain slice in the center of the recording chamber. Stabilize the brain slice with a slice hold down or a harp. To detect the fluorescent cell, find the area of interest under the four times magnification.
Then, move the microscope stage by turning on click to move mode and click the center of the area of interest. Alternatively, use the key pad to move the microscope stage. Next, switch to the high magnification lens and adjust the focus by moving the microscope in the Z-axis using RF on the keypad.
The software directs the microscope and the camera to take a series of images at different depths. Then, these images are subjected to computer vision processing to find fluorescently-labeled cells. The final output is a list of detected cell coordinates.
Click the detect cell button on the main GUI column unit zero. If the LED or laser light source of the setup cannot be controlled by the TTL signal, manually turn on the LED or laser. Turn off the LED or laser if necessary.
A list of cell coordinates will appear in the memory positions GUI. Remove undesired cells from the list by clicking the X button next to the coordinates. Alternatively, if target cells are not fluorescently-labeled, click mouse mode on the main GUI.
Then, 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 memory positions GUI. To perform secondary calibration in order to detect pipet offset coordinates, fill 1/3 of a glass pipet with internal solution. Then, load the pipet onto the pipet holder attached to the head stage.
At low magnification, use one and two on the keypad to switch between unit one and unit two. Then, bring the pipet into the visual field and adjust the focus using the keypad. Next, load the primary calibration by clicking on load calibration.
Switch the microscope lens to high magnification and click 40 times on the main GUI. Bring the pipet tip to the center. Then, click the secondary calibration button on the main GUI, under the unit that is in use.
To patch a target cell, click on the patch control button to open the patch control GUI. Click the go to button next to the cell of interest on the coordinate list in the memory position GUI. Subsequently, click on the CTM button of the unit in the main GUI to enable movement.
Click on the cell of interest to move the pipet tip to the cell. Due to mechanical limitation of the manipulator, when traveling longer than 500 micrometer distance, there may be a slight positioning error. To prevent a large mechanical positioning error, secondary calibration should be performed near the target cell.
Next, use the unit one selected button to switch the input signal between the two units. Click on the patch button on the patch control GUI. Automatic patching will begin and the pressure and resistance can be monitored on the patch control GUI.
The automatic patching algorithm monitors resistance and controls pressure through a series of logic loops, to form a gigaseal. A popup window notifies the formation of a gigaseal. The system utilizes resistance change to recognize cell-surface contact.
In the situation that the cell-surface contact is not detected in time, use the next button to advance patching stage while remaining in the same automatic patching trial. Manipulate the automatic process at any point by clicking on the respective buttons on the patch control GUI. Then, select yes to break in with combined Zap and suction.
Alternatively, select no to break in with suction only. Then, save the experiment patch log. This figure shows the selected locations loaded to the command sequence GUI.
The left column shows the list of coordinates and the right column shows the list of commands in the form of TTL signals for each location. Here are the screenshots during the drug application experiment, corresponding to the three selected locations. KCL solution was mixed with red fluorescent dye for the purpose of visualization.
Unit one was the KCL-containing pipet and unit two was the patching pipet. In this figure, the step current injections show a regular spiking neuron. Shown here are the voltage clamp recording traces from the local application of 500 millimolar potassium chloride solution at three locations.
The red trace with inward current was recorded from the trial when potassium chloride application was close to the patch cell. The red arrow indicates the timing of potassium chloride application. Using this method, one can perform a patch clamp trial within four minutes without extensive training compared to manual patching.
Following this procedure, other methods like optogenetics, chemogenetics, and pharmacological manipulations, can be performed in order to answer questions related to your research project. After its development, this technique may pave the way for researchers in the field of neuroscience to explore high-throughput studies of synaptic receptors and ion channels in different in vitro preparations. After watching this video, you should have a good understanding of how to perform image-guided automated patch clamp experiments.
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