Abstract

A fluorometric method that allows repeatable measurement of sugar transport rates and parameters in single living cells is presented. Intracellular sugar concentrations were estimated in real time from changes in cell volume that occur secondary to permeation of sugars across the plasma membrane. In turn, the cell volume changes were estimated from variations of intracellular calcein fluorescence measured by confocal microscopy. Using HeLa cells, the assay allowed reproducible measurement of the uptake and exit of D-galactose and 3-O-methyl-D-glucose. The rate of zero-trans uptake (i.e. at an intracellular concentration of zero) of galactose at an extracellular concentration of 200 mM was 0.34±0.05 mM/s (n=8). Apparent V(max) and K(m) for galactose exit were 0.32±0.05 mM/s (n=9) and 30±7.2 mM (n=9), respectively. The apparent affinity of infinite-trans (i.e. at a very high intracellular concentration) uptake of 3-O-methyl-D-glucose was 3.8±0.47 mM (n=8). Galactose uptake was 93±8% (n=8) inhibited in the presence of 50 ?M phloretin, whereas galactose exit was 96±6% (n=5) trans-inhibited by 100 mM 4,6-ethylidine-D-glucose. This technique may help to characterize sugar transport in freshly isolated cells, co-cultures and heterogeneous cell explants. It may also allow available cell microinjection technology to be used to study the regulation of sugar transporters' intrinsic activity. In principle, similar approaches might also be applied in functional studies of other transporters for which non-metabolized substrates are available.