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Intracellular signal transduction is often regulated by transient protein phosphorylation in response to external stimuli. Insulin signaling is dependent on specific protein phosphorylation events, and analysis of insulin receptor substrate-1 (IRS-1) phosphorylation reveals a complex interplay between tyrosine, serine, and threonine phosphorylation. The phospho-specific antibody-based quantification approach for analyzing changes in site-specific phosphorylation of IRS-1 is difficult due to the dearth of phospho-antibodies compared with the large number of known IRS-1 phosphorylation sites. We previously published a method detailing a peak area-based mass spectrometry approach, using precursor ions for peptides, to quantify the relative abundance of site-specific phosphorylation in the absence or presence of insulin. We now present an improvement wherein site-specific phosphorylation is quantified by determining the peak area of fragment ions respective to the phospho-site of interest. This provides the advantage of being able to quantify co-eluting isobaric phosphopeptides (differentially phosphorylated versions of the same peptide), allowing for a more comprehensive analysis of protein phosphorylation. Quantifying human IRS-1 phosphorylation sites at Ser303, Ser323, Ser330, Ser348, Ser527, and Ser531 shows that this method is linear (n = 3; r(2) = 0.85 +/- 0.05, 0.96 +/- 0.01, 0.96 +/- 0.02, 0.86 +/- 0.07, 0.90 +/- 0.03, 0.91 +/- 0.04, respectively) over an approximate 10-fold range of concentrations and reproducible (n = 4; coefficient of variation = 0.12, 0.14, 0.29, 0.30, 0.12, 0.06, respectively). This application of label-free, fragment ion-based quantification to assess relative phosphorylation changes of specific proteins will prove useful for understanding how various cell stimuli regulate protein function by phosphorylation.

Label-free relative quantification of co-eluting isobaric phosphopeptides of insulin receptor substrate-1 by HPLC-ESI-MS/MS