INTRODUCTIONQuantitative proteomics has traditionally been performed using 2D gel electrophoresis, where quantitation is accomplished by recreating differences in the staining patterns of proteins derived from two states of cell populations or tissues from a similar biological system. More recently, mass spectrometry (MS) methods based on stable isotope quantitation have been developed that show significant potential for differential expression proteomic studies. One such in vitro method, described in this protocol, involves the use of isotope-coded affinity tags (ICATs) with three functional moieties: a cysteine reactive moiety, a linker with either eight hydrogens (the light form of the reagent) or eight deuteriums (the heavy form of the reagent, having an isotope code or mass tag of 8 Da), and a biotin moiety (the affinity tag). Using this technique, the cysteine side chains in complex mixtures of proteins from two different states of a cell population (e.g., normal vs. disease) are reduced and alkylated using the light form of the reagent (d0-labeled tag) in one cell state and the heavy form of the reagent (d8-labeled tag) for proteins in the second cell state. The two mixtures (d0 and d8 labeled) are then combined and subjected to proteolytic digestion (typically, with trypsin and/or Lys-C). Generated cysteine-containing peptides are affinity-purified using an avidin column, resulting in a "simplified" mixture of peptides that contains ~10-fold fewer peptides than the original mixture. These peptides are analyzed by MS, and quantitation information based on the relative abundance of the d0 and d8 isotopes is obtained. The identification of proteins is obtained from the peptide molecular mass and MS/MS-derived amino acid sequence.