We present a novel method for characterization of uncharged colloids in aqueous suspension that relies on the shifting the electrophoretic mobility of colloidal particles by attaching DNA strands of a known length in the presence of electric fields. DNA attachment is achieved by covalently attaching lipophilic groups to the DNA, including n-alkanes, cholesterol, and bile salts. Buffers containing the colloid of interest are loaded into the capillary and a small sample of the alkylated DNA is loaded into the front end of the capillary. On application of electric field, the DNA transiently binds to colloidal particle, momentarily imparting a significant negative mobility on the colloid-DNA complex. The mobility shift is largest for smaller particles (which are more greatly impacted by DNA attachment) and for those that bind the alkyl group with highest affinity. These two effects are decoupled by performing separate runs with varying concentrations of the colloid of interest. Mobility is measured by analyzing the migration time of labeled DNA in the presence of the colloidal suspension using a equilibrium-based model for the DNA attachment.

Peaks in the CE traces remain sharp even when using polydisperse colloidal dispersions, as each DNA strand binds with thousands of distinct particles during the run and the measured mobility represents their time average. Size and binding affinity measurements are presented for nonionic micelles (Triton X-100 and C16E6), oil-in-water nanoemulsions, and carbon nanotubes, and are compared to light scattering and other independent techniques. The CE-based DNA tagging method emerges as a reliable method to size colloidal dispersions without labeling, and to quantify adsorption equilibria between nonpolar solutes and these colloids. Measurements can be performed in less than 5 minutes, require only nanoliter volumes, and do not require any labeling of the colloids.