Patch-clamp recording

We adopted the standard whole-cell patch-clamp technique to record sodium currents. For I_Nav1.5 recording, cells on cover slips were transferred to a continuously perfused recording chamber mounted on the stage of an inverted microscope and perfused with 10 mL of bath solution at a rate of about 1 mL/min for electrophysiological recording. Glass pipettes (1.5 mm diameter) were pulled with a two-stage microelectrode puller (PC-10, Narishige, Japan), and the resistance of the pipettes ranged from 2–4 MΩ when they were filled with the internal solution and immersed in the bath solution. For I_Na recording in human atrial myocytes, the solution containing the isolated cells was dropped in an open perfusion chamber (1 mL) mounted on the stage of an inverted microscope. The myocytes were allowed to adhere to the bottom of the chamber for 10–15 min and were then superfused with bath solution at a rate of 2–3 mL/min. The resistance of the pipettes ranged from 2–3 MΩ when they were filled with the internal solution. Only quiescent rod-shaped cells showing clear cross-striations were used. Membrane currents were recorded with an EPC-10 amplifier and Pulse software (HEKA Electronik, Lambrecht, Germany) at room temperature (22–24 °C). The liquid junction potentials between the pipette and bath solutions were compensated for before the pipette touched the cell. After obtaining a gigaseal, we ruptured the membrane via gentle suctioning to form the whole-cell configuration. The current signal was filtered at 3 kHz and sampled at 10 kHz. The whole-cell capacitance and resistance were compensated for, and the leak currents were subtracted. Currents were recorded 5 min after the whole-cell patch-clamp configuration was achieved.