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Phosphoprotein Isotope-Coded Solid-Phase Tag Approach for Enrichment and Quantitative Analysis of Phosphopeptides from Complex Mixtures Wei-Jun Qian,† Michael B. Goshe,*,‡ David G. Camp II,† Li-Rong Yu,†,§ Keqi Tang,† and Richard D. Smith*,†
Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, P.O. Box 999, MSIN: K8-98, Richland, Washington 99352, and Department of Molecular and Structural Biochemistry, North Carolina State University, 128 Polk Hall, Campus Box 7622, Raleigh, North Carolina 27695-7622
Many cellular processes are regulated by reversible protein phosphorylation, and the ability to broadly identify and quantify phosphoproteins from proteomes would provide a basis for gaining a better understanding of these dynamic cellular processes. However, such a sensitive, efficient, and global method capable of addressing the phosphoproteome has yet to be developed. Here we describe an improved stable-isotope labeling method using a phosphoprotein isotope-coded solid-phase tag (PhIST) for isolating and measuring the relative abundances of phosphorylated peptides from complex peptide mixtures resulting from the enzymatic digestion of extracted proteins. The PhIST approach is an extension of the previously reported phosphoprotein isotope-coded affinity tag (PhIAT) approach developed by our laboratory,1,2 where phosphoseryl and phosphothreonyl residues were derivatized by hydroxide ion-mediated β-elimination followed by the Michael addition of 1,2-ethanedithiol (EDT). Instead of using the biotin affinity tag, peptides containing the EDT moiety were captured and labeled in one step using isotope-coded solid-phase reagents containing either light (12C6, 14N) or heavy (13C6, 15N) stable isotopes. The captured peptides labeled with the isotope-coded tags were released from the solid-phase support by UV photocleavage and analyzed by capillary liquid chromatography-tandem mass spectrometry. The efficiency and sensitivity of the PhIST labeling approach for identification of phosphopeptides from mixtures were determined using casein proteins. Its utility for proteomic applications was demonstrated by the labeling of soluble phosphoproteins from a human breast cancer cell line. Reversible phosphorylation of proteins on serine, threonine, and tyrosine residues is one of the most prevalent posttranslational modifications, and nearly 30% of all proteins expressed in eukaryotic cells are phosphorylated to some extent.3,4 The general significance of phosphorylation has been well appreciated as a central mechanism that regulates nearly every aspect of cellular life and touches almost every known signaling pathway.5 There10.1021/ac0342774 CCC: $25.00 Published on Web 09/05/2003
© 2003 American Chemical Society
fore, our understanding of signaling networks and cellular function will be greatly enhanced by the development of high-throughput techniques that allow quantitative mapping of phosphoproteins and their respective sites of phosphorylation. Despite its importance, the global analysis of protein phosphorylation remains a major analytical challenge for several reasons including the low abundance of many signaling phosphoproteins within cells, the low stoichiometry of phosphorylation at any specific site for a given protein, and the limited dynamic range for current analytical techniques employed for the study of protein phosphorylation. Traditionally, protein phosphorylation has been studied by metabolic 32P labeling of cellular proteins coupled with 2-D polyacrylamide gel electrophoresis and followed by Edman sequencing.6,7 These techniques are insufficient when dealing with complex mixtures of proteins, are labor intensive, and suffer from the difficulties associated with the use of radioactive isotopes. In recent years, mass spectrometry (MS) has become the method of choice for protein phosphorylation analysis.8,9 Although various MS-based techniques (e.g., peptide mapping, precursor ion scanning, and neutral loss scanning) have been exploited for the detection of phosphorylation, these techniques alone are generally best suited for the analysis of single proteins or simple mixtures of phosphoproteins. Recently, several novel methodologies have been developed for selective enrichment of phosphopeptides from complex †
Pacific Northwest National Laboratory. North Carolina State University. § Current address: SAIC-Frederick Inc., National Cancer Institute at Frederick, Analytical Chemistry Laboratory, Mass Spectrometry Center, P.O. Box B, Frederick, MD 21702. (1) Goshe, M. B.; Conrads, T. P.; Panisko, E. A.; Angell, N. H.; Veenstra, T. D.; Smith, R. D. Anal. Chem. 2001, 73, 2578-2586. (2) Goshe, M. B.; Veenstra, T. D.; Panisko, E. A.; Conrads, T. P.; Angell, N. H.; Smith, R. D. Anal. Chem. 2002, 74, 607-616. (3) Hubbard, M. J.; Cohen, P. Trends Biochem. Sci. 1993, 18, 172-177. (4) Cohen, P. Trends Biochem. Sci. 2000, 25, 596-601. (5) Hunter, T. Cell 2000, 100, 113-127. (6) Wettenhall, R. E. H.; Aebersold, R. H.; Hood, L. E. Methods Enzymol. 1991, 201, 186-199. (7) van der Geer, P.; Hunter, T. Electrophoresis 1994, 15, 544-554. (8) Resing, K. A.; Ahn, N. G. Methods Enzymol. 1997, 283, 29-44. (9) Mann, M.; Ong, S. E.; Gronborg, M.; Steen, H.; Jensen, O. N.; Pandey, A. Trends Biotechnol. 2002, 20, 261-268. ‡
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mixtures using chemical modification strategies or immobilized metal affinity chromatography (IMAC).1,10-12 Our group1 and Oda et al.,10 concurrently and independently, developed two similar methods for modifying and enriching phosphoseryl (pSer) and phosphothreonyl (pThr) peptides. Both methods involve hydroxide ion-mediated β-elimination of the O-phosphate moiety and the addition of 1,2-ethanedithiol (EDT) followed by biotinylation and isolation using avidin affinity chromatography. Our method, termed PhIAT (phosphoprotein isotope-coded affinity tag), incorporated stable isotope labeling by using either light (HSCH2CH2SH, EDT-d0) or heavy (HSCD2CD2SH, EDT-d4) isotopic versions of EDT, allowing relative quantitation of protein phosphorylation states. One disadvantage of the β-elimination modification is that it is not applicable to phosphorylated tyrosine (pTyr) residues. Zhou et al.11 developed an alternative approach capable of enriching pSer, pThr, and pTyr peptides; however, the requirement of multiple chemical modifications and purification steps leads to substantial sample loss. More recently, Ficarro et al.12 reported an improved IMAC approach for enrichment of phosphopeptides by converting all peptides to their corresponding methyl esters, which reduces the nonspecific adsorption of nonphosphorylated peptides. Sample recovery remains a potential limitation of this method due to inefficient binding of the esterified phosphopeptides on the IMAC column. Another potential limitation for the approaches of Zhou et al. and Ficarro et al. is the difficulty in identifying the exact site of phosphorylation by tandem MS due to the lability of the phosphate group during collisioninduced dissociation (CID).13 In contrast, the PhIAT approach introduces a stable isotope-coded label at each site of phosphorylation, thus facilitating the exact identification and relative quantitation of each phosphorylated site.2 Although some promising results have been demonstrated with the PhIAT approach,1,2 several limitations associated with avidin affinity chromatography have been observed, including difficulty in removing all nonspecifically bound peptides as well as reduced sample recovery due to irreversible binding of a subpopulation of the biotinylated peptides. To address these issues, a novel isotope labeling strategy, termed the phosphoprotein isotopecoded solid-phase tag (PhIST) approach, has been developed using an isotope-coded solid-phase tag in place of the biotin affinity tag implemented in our original PhIAT approach. An isotope-coded solid-phase tag similar to that used by Zhou and co-workers14 was coupled with the PhIAT approach to more effectively isolate phosphopeptides from complex peptide mixtures. Improvements in the overall sensitivity and efficiency for isolating and analyzing phosphopeptides using the PhIST approach are demonstrated. EXPERIMENTAL SECTION Materials. The isotope-coded light and heavy solid-phase reagents (Figure 1) were synthesized according to the protocol described by Zhou et al.14 Aminopropyl controlled pore glass beads were purchased from CPG, Inc. (Lincoln Park, NJ). (10) Oda, Y.; Nagasu, T.; Chait, B. T. Nat. Biotechnol. 2001, 19, 379-382. (11) Zhou, H.; Watts, J. D.; Aebersold, R. Nat. Biotechnol. 2001, 19, 375-378. (12) Ficarro, S. B.; McCleland, M. L.; Stukenberg, P. T.; Burke, D. J.; Ross, M. M.; Shabanowitz, J.; Hunt, D. F.; White, F. M. Nat. Biotechnol. 2002, 20, 301-305. (13) DeGnore, J. P.; Qin, J. J. Am. Soc. Mass Spectrom. 1998, 9, 1175-1188. (14) Zhou, H. L.; Ranish, J. A.; Watts, J. D.; Aebersold, R. Nat. Biotechnol. 2002, 19, 512-515.
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N-Hydroxybenzotriazole, 4-[4-(1-(Fmoc-amino)ethyl)-2-methoxy5-nitrophenoxy)butanoic acid (Fmoc-aminoethyl photolinker), and N-Fmoc-D-leucine were purchased from Novabiochem (San Diego, CA). L-Leucine-N-Fmoc (U-13C6, 98%, 15N, 98%) was purchased from Cambridge Isotope Laboratories (Andover, MA). The 1,2ethanedithiol (>98%) was purchased from Fluka. Tris(2-carboxyethyl)phosphine hydrochloride (TCEP-HCl) and 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) were obtained from Pierce (Rockford, IL). β-Casein (from bovine milk with Rs1-, Rs2-, and κ-casein comprising
