Abstract
N-acyl-phosphatidylethanolamine (NAPE)-hydrolyzing phospholipase D (NAPE-PLD) is a zinc metallohydrolase enzyme that converts NAPEs to bioactive N-acyl-ethanolamides. Altered NAPE-PLD activity may contribute to pathogenesis of obesity, diabetes, atherosclerosis, and neurological diseases. Selective measurement of NAPE-PLD activity is challenging, however, because of alternative phospholipase pathways for NAPE hydrolysis. Previous methods to measure NAPE-PLD activity involved addition of exogenous NAPE followed by TLC or LC/MS/MS, which are time and resource intensive. Recently, NAPE-PLD activity in cells has been assayed using the fluorogenic NAPE analogs PED-A1 and PED6, but these substrates also detect the activity of serine hydrolase-type lipases PLA1 and PLA2. To create a fluorescence assay that selectively measured cellular NAPE-PLD activity, we synthesized an analog of PED-A1 (flame-NAPE) where the sn-1 ester bond was replaced with an N-methyl amide to create resistance to PLA1 hydrolysis. Recombinant NAPE-PLD produced fluorescence when incubated with either PED-A1 or flame-NAPE, whereas PLA1 only produced fluorescence when incubated with PED-A1. Furthermore, fluorescence in HepG2 cells using PED-A1 could be partially blocked by either biothionol (a selective NAPE-PLD inhibitor) or tetrahydrolipstatin (an inhibitor of a broad spectrum of serine hydrolase-type lipases). In contrast, fluorescence assayed in HepG2 cells using flame-NAPE could only be blocked by biothionol. In multiple cell types, the phospholipase activity detected using flame-NAPE was significantly more sensitive to biothionol inhibition than that detected using PED-A1. Thus, using flame-NAPE to measure phospholipase activity provides a rapid and selective method to measure NAPE-PLD activity in cells and tissues.