Divalent cation block of inward currents and low-affinity K+ uptake in Saccharomyces cerevisiae.

We have used the patch clamp technique to characterize whole-cell currents in spheroplasts isolated from a trk1Delta trk2Delta strain of Saccharomyces cerevisiae which lacks high- and moderate-affinity K+ uptake capacity. In solutions in which extracellular divalent cation concentrations were 0.1 mM, cells exhibited a large inward current. This current was not the result of increasing leak between the glass pipette and membrane, as there was no effect on the outward current. The inward current comprised both instantaneous and time-dependent components. The magnitude of the inward current increased with increasing extracellular K+ and negative membrane potential but was insensitive to extracellular anions. Replacing extracellular K+ with Rb+, Cs+, or Na+ only slightly modulated the magnitude of the inward current, whereas replacement with Li+ reduced the inward current by approximately 50%, and tetraethylammonium (TEA+) and choline were relatively impermeant. The inward current was blocked by extracellular Ca2+ and Mg2+ with apparent Kis (at -140 mV) of 363 +/- 78 and 96 +/- 14 microM, respectively. Furthermore, decreasing cytosolic K+ increased the magnitude of the inward current independently of the electrochemical driving force for K+ influx, consistent with regulation of the inward current by cytosolic K+. Uptake of 86Rb+ by intact trk1Delta trk2Delta cells was inhibited by extracellular Ca2+ with a Ki within the range observed for the inward current. Furthermore, increasing extracellular Ca2+ from 0.1 to 20 mM significantly inhibited the growth of these cells. These results are consistent with those of the patch clamp experiments in suggesting that low-affinity uptake of alkali cations in yeast is mediated by a transport system sensitive to divalent cations.