Protein lysine methyltransferases (PKMTs) catalyze the methylation of protein substrates, and their dysregulation has been linked to many diseases, including cancer. Accumulated evidence suggests that the reaction path of PKMT-catalyzed methylation consists of the formation of a cofactor(cosubstrate)-PKMT-substrate complex, lysine deprotonation through dynamic water channels, and a nucleophilic substitution (S N 2) transition state for transmethylation. However, the molecular characters of the proposed process remain to be elucidated experimentally. Here we developed a matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF-MS) method and corresponding mathematic matrix to determine precisely the ratios of isotopically methylated peptides. This approach may be generally applicable for examining the kinetic isotope effects (KIEs) of posttranslational modifying enzymes. Protein lysine methyltransferase SET8 is the sole PKMT to monomethylate histone 4 lysine 20 (H4K20) and its function has been implicated in normal cell cycle progression and cancer metastasis. We therefore implemented the MSbased method to measure KIEs and binding isotope effects (BIEs) of the cofactor S-adenosyl-L-methionine (SAM) for SET8-catalyzed H4K20 monomethylation. A primary intrinsic 13 C KIE of 1.04, an inverse intrinsic α-secondary CD 3 KIE of 0.90, and a small but statistically significant inverse CD 3 BIE of 0.96, in combination with computational modeling, revealed that SET8-catalyzed methylation proceeds through an early, asymmetrical S N 2 transition state with the C-N and C-S distances of 2.35-2.40 Å and 2.00-2.05 Å, respectively. This transition state is further supported by the KIEs, BIEs, and steadystate kinetics with the SAM analog Se-adenosyl-L-selenomethionine (SeAM) as a cofactor surrogate. The distinct transition states between protein methyltransferases present the opportunity to design selective transition-state analog inhibitors.S tepwise progression of an enzyme-catalyzed chemical reaction is accompanied by changes of bond orders and vibrational modes involved with specific atoms of the reactant(s) (1, 2). Such changes can be traced experimentally by measuring the ratios of turnover rates [kinetic isotope effects (KIEs)] or binding affinities [binding isotope effects (BIEs)] of the reactant(s) when the relevant atoms are replaced by heavy isotopes (3, 4). KIEs and BIEs are thus useful parameters for elucidating transition-state (TS) structures and catalytic mechanisms, which sometimes cannot be elucidated readily through sole measurement of steady-state kinetics (5-9). A sufficient set of KIEs and BIEs at the positions involved with bond motions can afford electrostatic and geometric constraints, when combined with computational modeling, to define an enzymatic TS (10-12). This information provides not only the atomic resolution of the transient structure at the highest energy summit along the reaction path, but also structural guidance for designing tight-binding TS analog inhibitors (13...
Kinase-catalyzed protein phosphorylation is involved in a wide variety of cellular events. Development of methods to monitor phosphorylation is critical to understand cell biology. Our lab recently discovered kinase-catalyzed biotinylation, where ATP-biotin is utilized by kinases to label phosphopeptides or phosphoproteins with a biotin tag. To exploit kinase-catalyzed biotinylation for phosphoprotein purification and identification in a cellular context, the susceptibility of the biotin tag to phosphatases was characterized. We found that the phosphorylbiotin group on peptide and protein substrates was relatively insensitive to protein phosphatases. To understand how phosphatase stability would impact phosphoproteomics research applications, kinase-catalyzed biotinylation of cell lysates was performed in the presence of kinase or phosphatase inhibitors. We found that biotinylation with ATP-biotin was sensitive to inhibitors, although with variable effects compared to ATP phosphorylation. The results suggest that kinase-catalyzed biotinylation is well suited for phosphoproteomics studies, with particular utility towards monitoring low abundance phosphoproteins or characterizing the influence of inhibitor drugs on protein phosphorylation.
Kinase-catalyzed protein phosphorylation is involved in a wide variety of cellular events. Development of methods to monitor phosphoproteins in normal and diseased states is critical to fully characterize cell signalling. Towards phosphoprotein analysis tools, our lab reported kinase-catalyzed labeling where γ-phosphate modified ATP analogs are utilized by kinases to label peptides or protein substrates with a functional tag. In particular, the ATP-biotin analog was developed for kinase-catalyzed biotinylation. However, kinase-catalyzed labeling has been tested rigorously with only a few kinases, preventing use of ATP-biotin as a general tool. Here, biotinylation experiments, gel or HPLC-based quantification, and kinetic measurements indicated that twenty-five kinases throughout the kinome tree accepted ATP-biotin as a cosubstrate. With this rigorous characterization of ATP-biotin compatibility, kinase-catalyzed labeling is now immediately useful for studying phosphoproteins and characterizing the role of phosphorylation in various biological events.
Kinase‐catalyzed protein phosphorylation plays an essential role in a variety of biological processes. Methods to detect phosphoproteins and phosphopeptides in cellular mixtures will aid in cell biological and signaling research. Our laboratory recently discovered the utility of γ‐modified ATP analogues as tools for studying phosphorylation. Specifically, ATP‐biotin can be used for labeling and visualizing phosphoproteins from cell lysates. Because the biotin tag is suitable for protein detection, the biotinylation reaction can be applied to multiple phosphoproteomics applications. Herein, we report a general protocol for labeling phosphopeptides and phosphoproteins in biological samples using kinase‐catalyzed biotinylation. Curr. Protoc. Chem. Biol. 4:83‐100 © 2012 by John Wiley & Sons, Inc.
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