Abstract

Antibiotic resistance is a major challenge for modern medicine. One of the first biochemical mechanisms discovered in multi-drug resistance involved a decrease in drug permeation across the bacterial membrane. Therefore, the quantification of drug transport across the bacterial membrane is critical. We present an optofluidic permeability assay which quantifies the permeability coefficient of drugs crossing lipid membranes. We deconstruct the bacterial membrane and study drug transport across its various components, using lipid vesicles as model systems. The drug molecules are tracked directly by stimulating their autofluorescence in the ultraviolet. Lipid vesicles are mixed with the drug in a controlled manner in microchannels and viewed at various time points; the increase in drug autofluorescence intensity within the vesicles over time is used to calculate the drug’s permeability. We show that changing the lipid composition of the membrane can change drug permeability by an order of magnitude. We further show that incorporating the Escherichia coli outer membrane porin OmpF in the vesicles significantly enhances the flux of the fluoroquinolone norfloxacin across the membrane, demonstrating the importance of such membrane porins in antibiotic transport. Finally, we end with a glimpse of how similar techniques can also be used to study drug uptake in bacteria on the single cell level, thus providing a new tool to identify bacteria tolerant to drugs on the basis of reduced antibiotic uptake.