Several mass spectrometry based assays have emerged for the quantitative profiling of cellular tyrosine phosphorylation. Ideally, these methods should reveal the exact sites of tyrosine phosphorylation, be quantitative and not cost-prohibitive. The latter is often an issue as typically several milligrams of (stable isotope labeled) starting protein material is required to enable the detection of low abundant phosphotyrosine peptides. Here, we adopted and refined a peptide centric immuno-affinity purification approach for the quantitative analysis of tyrosine phosphorylation by combining it with a cost-effective stable isotope dimethyl labeling method. We were able to identify by mass spectrometry, using just two LC-MS/MS runs, more than 1100 unique non-redundant phosphopeptides in HeLa cells from about 4 mg of starting material without requiring any further affinity enrichment as close to 80% of the identified peptides were tyrosine phosphorylated peptides. Stable isotope dimethyl labeling could be incorporated prior to the immuno-affinity purification, even for the used large quantities (mg) of peptide material, enabling the quantification of differences in tyrosine phosphorylation upon pervanadate treatment or EGF stimulation. Analysis of the EGF-stimulated HeLa cells, a frequently used model system for tyrosine phosphorylation, resulted in the quantification of 73 regulated unique phosphotyrosine peptides. The quantitative data was found to be exceptionally consistent with the literature, evidencing that such a targeted quantitative phosphoproteomics approach can provide reproducible results. In general, the combination of immuno-affinity purification of tyrosine phosphorylated peptides with large scale stable isotope dimethyl labeling provides a cost-effective approach that can alleviate variation in sample preparation and analysis as samples can be combined early on. Using this approach a rather complete qualitative and quantitative picture of tyrosine phosphorylation signaling events can be generated.