Initial rates of sugar uptake (zero-trans rates) are often measured by incubating yeast cells with radiolabeled sugars for 5 to 30 s and determining the radioactivity entering the cells. The yeast cells used are usually harvested from growth medium, washed, suspended in nutrient-free buffer, and stored on ice before they are assayed. With this method, the specific rates of zero-trans lactose uptake by Kluyveromyces lactis or recombinant Saccharomyces cerevisiae strains harvested from lactose fermentations were three- to eightfold lower than the specific rates of lactose consumption during fermentation. No significant extracellular beta-galactosidase activity was detected. The ATP content and adenylate energy charge (EC) of the yeasts were relatively low before the [(14)C]lactose uptake reactions were started. A short (1- to 7-min) preincubation of the yeasts with 10 to 30 mM glucose caused 1.5- to 5-fold increases in the specific rates of lactose uptake. These increases correlated with increases in EC (from 0.6 to 0.9) and ATP (from 4 to 8 micromol x g dry yeast(-1)). Stimulation by glucose affected the transport V(max) values, with smaller increases in K(m) values. Similar observations were made for maltose transport, using a brewer's yeast. These findings suggest that the electrochemical proton potential that drives transport through sugar/H(+) symports is significantly lower in the starved yeast suspensions used for zero-trans assays than in actively metabolizing cells. Zero-trans assays with such starved yeast preparations can produce results that seriously underestimate the capacity of sugar/H(+) symports. A short exposure to glucose allows a closer approach to the sugar/H(+) symport capacity of actively metabolizing cells.