An approach to quantifying N-linked glycoproteins by enzyme-catalyzed 18O3-labeling of solid-phase enriched glycopeptides.

Global analysis of glycoproteins shows great promise for the discovery of therapeutic targets and clinical biomarkers. Selective capture of glycopeptides by hydrazide resin followed by mass spectrometric identification of the peptides released by PNGaseF treatment has been most widely used. However, the majority of the reports using this approach focus on global profiling, rather than relative quantitation of glycoprotein alternations in pathological states. We describe an integrated strategy allowing for relative quantitation of glycoproteins in complex biological mixtures using this approach. The strategy includes periodate oxidation of tryptic digests, solid-phase enrichment of glycopeptides via hydrazide-coupled magnetic beads, in conjunction with (18)O stable isotope labeling catalyzed by both trypsin and PNGaseF, and subsequent identification and quantitation by LC-MS/MS analysis. Three (18)O atoms ((18)O(3)) are incorporated into N-linked glycopeptides for samples treated in (18)O-water, two at the carboxyl terminus by trypsin during hydrazide coupling and the third at the N-glycosylation site through PNGaseF-mediated deglycosylation. Thus, mass shifts of 6 and 8 Da are indicative of singly and doubly glycosylated peptides, respectively. Experimental conditions were optimized to promote the trypsin-mediated (18)O(2) incorporation and prevent backbone exchange. The accuracy, reproducibility, and linearity of relative quantitation were evaluated by using 15 glycoproteins spiked into mouse serum at different concentration ratios. Using this approach, we were able to identify and quantitate 224 N-glycopeptides representing 130 unique glycoproteins from 20 μL of the undepleted mouse serum samples. The strategy can be easily adapted to the analysis of glycoproteins in tissues, cell lines, and other sample origins.