Phosphorylation is the most common post-translational modification of proteins, in which a phosphate group is added to the polar side chain of amino acid residues by protein kinase. [1,2] The groups are typically linked to serine (Ser), threonine (Thr), and tyrosine, but some kinases target uncommon residues such as histidine, arginine, lysine, cysteine, aspartic acid (Asp), and glutamic acid (Glu). [3] And the phosphate group can be removed via the catalysis of phosphatases which aid in the reversibility of phosphorylation. Depending on the balance of two enzymes, phosphorylation dynamically regulates protein activity, stability, and subcellular localization in organisms, playing a critical role in cell signal transduction. [[4], [5], [6]] Consequently, investigating altered phosphoproteins promotes the comprehension of life mechanisms and disease impacts, and hastens the discovery of therapeutic targets, diagnostic biomarkers, and prognostic biomarkers. [2,[6], [7], [8], [9], [10]]

The proteomics approach is currently the primary high-throughput technique for analyzing phosphorylation changes. A typical procedure involves digesting proteins into peptides via specific proteases and then using mass spectrometry (MS) to identify phosphopeptides. However, accurately detecting phosphopeptides in positive ion mode is frequently challenging due to their low abundance and highly negative groups from multiple modification sites. [11] Additionally, the phosphate group often undergoes neutral loss during tandem mass spectrometry (MS/MS) analysis, potentially resulting in the misinterpretation of phosphorylation information. [12] Therefore, there is a significant need to improve the accuracy of identifying phosphoproteins and phosphopeptides and analyzing them quantitatively.

A common technique for the quantification of peptides involves combining stable isotope labeling with liquid chromatography-mass spectrometry (LC-MS) or matrix-assisted laser desorption/ionization-mass spectrometry (MALDI-MS). Commercial kits that offer light/heavy labeling or isobaric labeling are available for treating individual samples, with subsequent MS analysis of a pooled sample to achieve relative quantification. [13,14] However, these kits are not very suitable for phosphoproteomics because they cannot label modification sites. Site-specific labeling is a promising approach that enables accurate identification and relative quantification of phosphopeptides based on stable isotope labeling. Several chemical derivatization techniques have been developed to label phosphorylation sites, which convert a phosphoserine (pS) or phosphothreonine (pT) residue to a neutral one through the Michael addition of ethanedithiol (EDT), dithiothreitol (DTT), or cysteamine following dephosphorylation. [[15], [16], [17], [18]] However, these methods have limitations, such as incomplete conversion or the requirement for expensive isotopic reagents, which restricts their application on a large scale. Additionally, enhancing the stability of the peptide-phosphate ester bond has proven to be an effective strategy. [19]

A previous research has shown that dimethylamine (DMA) can react with the modification site of O-glycopeptide after de-O-glycosylation by β-elimination. [20] A separate study indicated that amidation of carboxyl groups in peptides can occur without losing phosphorylation. [21] These findings informed our development of a two-step chemical derivatization strategy in this study, utilizing methylamine and DMA as derivatization reagents, respectively. Methylamine shields carboxyl groups, thereby reducing the adsorption of non-phosphorylated peptides during enrichment. DMA provides a cost-effective reagent to achieve high conversion rates and stable isotope labeling of phosphorylation sites. The results indicate that the two-step derivatization method shares the advantages of the EDT/DTT protocol and facilitates site-specific identification of phosphopeptides by utilizing reporter ions and comprehensive product ion coverage.