The K(+)-stimulated ATPase was partially purified from a plasma membrane fraction of suspension cultured cells of rose (Rosa damascena) by two different solubilization procedures. Solubilization with 30 mm octyl-beta-d-glucopyranoside followed by precipitation with ammonium sulfate increased the specific activity of the enzyme about 6-fold. Solubilization with 1% cholate removed all but 1% of the phospholipids and resulted in an almost total loss of ATPase activity. The subsequent addition of polar lipids restored >90% of the ATPase activity with a doubling in specific activity. Fractionation of the cholate-solubilized ATPase activity on a Sephadex G-150 column resulted in 88% of the ATPase activity being recovered in two discrete, approximately equal peaks. Both ATPase activities were similar to plasma membrane ATPase activities in pH optimum, substrate specificity, ion stimulation, and inhibitor sensitivity. Assays of marker enzymes for Golgi apparatus, endoplasmic reticulum, and mitochondria revealed only a low contamination (<7%) from other membranes in the plasma membrane-enriched preparations. Lacking an unequivocal marker for the tonoplast, intact vacuoles were isolated, and their membrane density and ATPase activity were characterized and shown not to correspond to those of the putative plasma membrane preparation. These results suggest that there are two forms of ATPase separable by size in the plasma membrane of rose.