How to Set Up a Detached Patch-Clamp Experiment

Obtain a plate for cells. Wash the plate two times with 10 milliliters of PBS (phosphate buffered saline) without calcium and magnesium. Add 2 milliliters of detacher (trypsin). Incubate the plate for three minutes at 37 degrees Celsius. Observe the plate under the microscope to ensure that the cells have detached. Gently move the plate to detach all cells from the bottom of the plate. Add 10 milliliters of HEK (human epidermal keratinocyctes) and FCS (fetal cow serum) medium. Use a 10-milliliter pipette to move the cells up and down.

Observe the cells under the microscope to ensure that they are single cells. Pipette using the 10-milliliter pipette until the cells are single, if needed. Centrifuge the cells for two minutes at 1,000 rotations per minute. Discard the supernatant. Place the cells in 10 milliliters of fetal calf serum. Centrifuge the cells again for two minutes at 1,000 rotations per minute. Discard the supernatant. Place the cells in 200 microliters of external recording solution. Observe the cells under the microscope for single cells with a smooth membrane.

Place the saline solution inside a patch clamp chip. Add a drop of intracellular electrolyte solution to the bottom part of the chip using a pipette. Mount the chip onto a twist cap. The twist cap should contain the reference electrode and suction line. Add a drop of extracellular electrolyte solution to the top of the chip using a pipette. The electrolyte solution allows for the chip to make contact with the electrode. Add 5 microliters of cell suspension to the saline solution in the chip. Position a single cell using suction through a pipette. Suction can be performed manually by sucking air through the pipette. Positioning the cell increases resistance to form a seal. The seal allows for investigation of the whole cell.