Whole-cell patch-clamp recordings

The whole-cell patch-clamp technique was described in our previously published papers (23). Trypsin was used to detach the cultured cells, which were planted onto glass coverslips at least 30 min prior to patch clamp recordings and continuously perfused with the standard bath solution (SBS). Whole-cell K⁺ currents were recorded at room temperature with a standard ramp-step protocol: a 200 ms step pulse to 20 mV, followed by a 500 ms ramp from –90 to +20 mV (HEK-293 cell) or +80 mV (U251MG cell and cardiac myocytes) every 10 s, from a holding potential of −90 mV. Electrical signal was amplified by a patch-clamp amplifier (EPC-10, HEKA, Lambrecht, Germany) and manipulated by Pulse software (HEKA, Lambrecht, Germany). The currents at +20 mV or +80 mV were recorded to analyze the differences between different treatments, and the currents of ramp pulse were used for I–V relationship analysis.

For dose-dependent analysis, the cells were perfused with SBS and a series of different concentrations of BL1249 or bepridil (0.1, 0.3, 1, 3, and 10 µM). The concentrations were determined according to previous studies; the highest concentration of bepridil was usually 10 µM and IC₅₀ toI_Ks =6.2 µM (15), IC₅₀ to BKca channels =1.866 µM (16), and IC₅₀ to K(ATP) channels =6.6 µM (17). The mean current of more than three cells from three different experiments was recognized as the final channel current of each experiment. The Origin and IGOR software were used to analyze all recordings. To normalize the membrane currents to cell size, the currents were divided by the capacitances of cell membrane to obtain the current density (pA/pF).