Membrane currents were recorded from isolated pig ventricular myocytes using the whole-cell configuration of the patch clamp technique.

Isolated cells were placed in a perfusion chamber (Warner Instruments) on the stage of an inverted microscope (Olympus IX71) and bathed in a K⁺-free extracellular solution containing (in mM) NaCl 140, CsCl 5.4, CaCl₂2.5, MgCl₂ 0.5, glucose 5.5, and HEPES 5 (pH 7.4), maintained at room temperature. Nisoldipine (1 µM) was added to the extracellular solution to block L-type Ca²⁺ channels.

Cells were patched using microelectrodes with resistances between 1 and 2 MΩ. Access resistance was compensated to obtain series resistance errors of less than 5 mV.

Cells were dialyzed with a K⁺-free microelectrode solution containing (in mM): Cs-glutamate 130, TEA-Cl 20, EGTA 5, MgATP 5, TrisGTP 0.06, and HEPES 5 (pH 7.2).

Whole cell currents were recorded under voltage-clamp conditions using a Multiclamp 700B voltage clamp amplifier, Digidata 1440A computer interface, and pClamp 11 data acquisition and analysis software (Molecular Devices). Data were lowpass filtered at 4 kHz, and sampled at 10 kHz.

The membrane potential was held at -30 mV. A 100 ms test pulse to +50 mV was used to measure the cAMP-regulated Cl^‑ current.

The time course of changes in the amplitude of the Cl^‑ current was monitored by measuring the amplitude of the current recorded during the test pulse to +50 mV applied once every 5 s.

A stable baseline (about 5 minutes) was obtained before application of test drugs.

The Cl^‑ current was defined as the cAMP-induced current obtained by subtracting the current recorded before application of test-drug(s) from those recorded after.