Glycoproteins secreted by cells play essential roles in the regulation of extracellular activities. Secreted
glycoproteins are often reflective of cellular status, and thus
glycoproteins from easily accessible bodily fluids can serve as excellent
biomarkers for disease detection. Cultured cells have been extensively employed as models in the research fields of biology and biomedicine, and global analysis of
glycoproteins secreted from these cells provides insights into cellular activities and
glycoprotein functions. However, comprehensive identification and quantification of secreted
glycoproteins is a daunting task because of their low abundances compared with the high-abundance
serum proteins required for cell growth and proliferation. Several studies employed
serum-free media to analyze secreted
proteins, but it has been shown that serum
starvation, even for a short period of time, can alter
protein secretion. To overcome these issues, we developed a method to globally characterize secreted
glycoproteins and their N-glycosylation sites from cultured cells by combining selective enrichment of secreted
glycoproteins with a boosting approach. The results demonstrated the importance of the boosting sample selection and the boosting-to-sample ratio for improving the coverage of secreted
glycoproteins. The method was applied to globally quantify secreted
glycoproteins from THP-1 monocytes and macrophages in response to
lipopolysaccharides (LPS) and from Hep G2 cells treated with TGF-β without serum
starvation. We found differentially secreted
glycoproteins in these model systems that showed the cellular response to the immune activation or the epithelial-to-mesenchymal transition. Benefiting from the selective enrichment and the signal enhancement of low-abundance secreted
glycoproteins, this method can be extensively applied to study secreted
glycoproteins without serum
starvation, which will provide a better understanding of
protein secretion and cellular activity.