Solid Phase Isobaric Mass Tag Reagent for Simultaneous Protein

Jun 7, 2010 - Leonardo Di Donna,† Fabio Mazzotti,† and Giovanni Sindona†. Dipartimento di Chimica, Universita` della Calabria, Arcavacata di Ren...
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Anal. Chem. 2010, 82, 5552–5560

Solid Phase Isobaric Mass Tag Reagent for Simultaneous Protein Identification and Assay Anna Napoli,*,† Constantinos M. Athanassopoulos,*,‡ Petros Moschidis,‡ Donatella Aiello,† Leonardo Di Donna,† Fabio Mazzotti,† and Giovanni Sindona† Dipartimento di Chimica, Universita` della Calabria, Arcavacata di Rende, Italy, and Department of Chemistry, University of Patras, Patras, Greece The solid phase isobaric mass tagging (SPIMT) approach is presented for simultaneous protein quantitation and identification. The novelty of the SPIMT strategy relies on a CID-based differentiation of regioisomeric species for quantitation of tagged proteolytic peptides. SPIMTs are unlabeled mass-tagging reagents, which consist of a reporter group, a mass balance group, and a spacer with a amine-specific reactive group, able to be linked to any N-terminal peptide. Therefore SPIMT-linked peptides from a two-plex set appear as a single unresolved precursor ion in MS, whereas the reporter groups lead to quantitation signals of m/z 168.2 and 182.2 Da upon tandem mass spectrometry (MS/MS) analysis with matrixassisted laser desorption time-of-flight/time-of-flight (MALDI TOF/TOF). This strategy allows ease protein identification by direct submission of MS and MS/MS data to the MASCOT database. SPIMT approach showed an excellent quantitation linearity, detecting any relative concentration differences of peptides in two solutions over a 5-fold concentration range without losing sequencing information. Therefore, SPIMTs are an attractive, simple, and low cost alternative for two-plex quantitation of proteins and offer possibilities of tuning the two-plex signal mass window by replacing the spacer. The development of reliable methodologies for simultaneous identification and quantitation of a large number of proteins is an important challenge in the proteomics field. Several methods utilizing the automated liquid chromatography coupled to a tandem mass spectrometer (LC-MS/MS) have been developed to achieve quantitative measurements. Stable-isotope labeling methods are the most broadly applied for sample preparation due to their high accuracy for relative quantitation.1 Stable isotopes are generally introduced into proteins or peptides by metabolic * To whom correspondence should be addressed. Prof. Anna Napoli, Ph.D., Via P. Bucci, Cubo 12/C, Dipartimento di Chimica, Universita` della Calabria, I-87030 Arcavacata di Rende, Italy. Phone: + 39 0984 492852. Fax +39 0984 493307. E-mail: [email protected]. Assistant Prof. Constantinos M. Athanassopoulos, Ph.D., University Campus, Department of Chemistry, University of Patras, GR-26500, Rio, Patras, Greece. Phone: +302610-997909. E-mail: [email protected]. † Universita` della Calabria. ‡ University of Patras. (1) Ong, S. E.; Mann, M. Nat. Chem. Biol. 2005, 1 (5), 252–262.

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labeling strategy (SILAC),2-4 enzymatic 18O-labeling of peptide C-termini,5-7 and chemical modification of specific functional groups using reagents such as isotope-coded affinity tags (ICATs).8 The restricted specificity of ICAT reagents only for cysteinyl peptides, in combination with the differential peptide elution profile, seems to be troublesome for a global proteomic analysis. Amine- reactive isobaric tags (TMSs)9,10 and isobaric mass tagging reagents (iTRAQ)11-14 have been designed to overcome these limitations. The introduction of iTRAQ has called the attention of the proteomics community because of the ability to perform relative (or absolute) quantification in up to four (and soon eight) phenotypes.11 Furthermore, peptides differentially labeled with isobaric tags coelute and tandem mass spectra display all of the sequence ions and quantitation signals, thus allowing simultaneous protein quantitation and identification. iTRAQ-based proteomics has been widely applied in systems ranging from microorganism stress responses15-18 to evaluating mammalian (2) Pasa-Tolic, L.; Jensen, P. K.; Lerson, G. A.; Lipton, M. S.; Peden, K. K.; Martinovic, S.; Tolic, N.; Bruce, J. E.; Smith, R. D. J. Am. Chem. Soc. 1999, 121, 7949–7950. (3) Oda, Y.; Huang, K.; Cross, F. R.; Cowburn, D.; Chait, B. T. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 6591–6596. (4) Ong, S. E.; Blagoev, B.; Kratchmarova, I.; Kristensen, D. B.; Steen, H.; Pandey, A.; Mann, M. Mol. Cell. Proteomics 2002, 1 (5), 376–386. (5) Yao, X.; Freas, A.; Ramirez, J.; Demirev, P. A.; Fenselau, C. Anal. Chem. 2001, 73 (13), 2836–2842. (6) Qian, W. J.; Monroe, M. E.; Liu, T.; Jacobs, J. M.; Anderson, G. A.; Shen, Y.; Moore, R. J.; Anderson, D. J.; Zhang, R.; Calvano, S. E.; Lowry, S. F.; Xiao, W.; Moldawer, L. L.; Davis, R. W.; Tompkins, R. G.; Camp, D. G.; Smith, R. D. Mol. Cell. Proteomics 2005, 4 (5), 700–709. (7) Mason, C. J.; Therneau, T. M.; Eckel-Passow, J. E.; Johnson, K. L.; Oberg, A. L.; Olson, J. E.; Nair, K. S.; Muddiman, D. C.; Bergen, H. R., III. Mol. Cell. Proteomics 2007, 6 (2), 305–18. (8) Gygi, S. P.; Rist, B.; Gerber, S. A.; Turecek, F.; Gelb, M. H.; Aebersold, R. Nat. Biotechnol. 1999, 17 (10), 994–999. (9) Thompson, A.; Schafer, J.; Kuhn, K.; Kienle, S.; Schwarz, J.; Schmidt, G.; Neumann, T.; Hamon, C. Anal. Chem. 2003, 75, 1895–1904. (10) Seo, J.; Suh, M. S.; Thangadurai, T. D.; Kim, J.; Rhee, Y. H.; Yoon, H. J.; Shin, S. K. Anal. Chem. 2008, 80, 6145–6153. (11) Ross, P. L.; Huang, Y. N.; Marchese, J. N.; Williamson, B.; Parker, K.; Hattan, S.; Khainovski, N.; Pillai, S.; Dey, S.; Daniels, S.; Purkayastha, S.; Juhasz, P.; Martin, S.; Barlet-Jones, M.; He, F.; Jacobson, A.; Pappin, D. J. Mol. Cell. Proteomics 2004, 3, 1154–1169. (12) Zhang, Y.; Wolf-Yadlin, A.; Ross, P. L.; Pappin, D. J.; Rush, J.; Lauffenburger, D. A.; White, F. M. Mol. Cell. Proteomics 2005, 4 (9), 1240–1250. (13) DeSouza, L.; Diehl, G.; Rodrigues, M. J.; Guo, J.; Romaschin, A. D.; Colgan, T. J.; Siu, K. W. J. Proteome Res. 2005, 4 (2), 377–386. (14) Choe, L.; D’Ascenzo, M.; Relkin, N. R.; Pappin, D. J.; Ross, P.; Williamson, B.; Guertin, S.; Pribil, P.; Lee, K. H. Proteomics 2007, 7, 3651–3660. (15) Lee, J.; Cao, L.; Ow, S. Y.; Barrios-Llerena, M. E.; Chen, W.; Wood, T. K.; Wright, P. C. J. Proteome Res. 2006, 5 (6), 1388–1397. (16) Rudella, A.; Friso, G.; Alonso, J. M.; Ecker, J. R.; van Wijk, K. J. Plant Cell 2006, 18 (7), 1704–1721. 10.1021/ac1004212  2010 American Chemical Society Published on Web 06/07/2010

Chart 1

organelles.11,19-22 These are relatively costly reagents, and largescale validation of this technique dramatically increases experimental costs. Quantitative measurements by site-specific, stable isotopic labeling of the cysteinyl peptide strategy23-25 using solid-phase isotope tagging reagents have been introduced to overcome the initial, time-consuming chromatographic step. The adaptation of stable isotopic labeling strategies to a solid phase format has shown comparable reproducibility, efficiency, and sensitivity. However, the reagents and sample preparation steps of some solid phase handles require synthetic and/or cleavage techniques not routinely available. Furthermore a large excess of solid-phase reactive sites is added compared to the sample dimension, which increases the risks of unwanted side reactions with noncysteinyl peptides. In this report, two-plex solid-phase isobaric label-free reagents based on “pseudo” tripeptides are proposed for quantitative protein analysis (see Chart 1). These reagents attach isobaric mass tags at the N-termini of digest peptide mixtures to perform relative quantification. The solid phase isobaric mass tagging (SPIMT) reagents were designed according to the principle that they must be isobaric and label-free rather than labeled with expensive heavy atoms. Consequently, all the derivatized peptides are isobaric and yield signature or reporter ions after CID that can be used to identify and quantify individual members of the two-plex set. SPIMT, like other solid-phase mass tagging reagents, consists of three parts: a linker, an isobaric tag, and a specific reactive group (see Chart 1). These SPIMTs can conveniently be prepared by in house solid-phase peptide methodologies. The first com(17) Redding, A. M.; Mukhopadhyay, A.; Joyner, D. C.; Hazen, T. C.; Keasling, J. D. Brief. Funct. Genomics Proteomics 2006, 5 (2), 133–143. (18) Aggarwal, K.; Choe, L. H.; Lee, K. H. Proteomics 2005, 5 (9), 2297–2308. (19) Chen, X.; Walker, A. K.; Strahler, J. R.; Simon, E. S.; Tomanicek-Volk, S. L.; Nelson, B. B.; Hurley, M. C.; Ernst, S. A.; Williams, J. A.; Andrews, P. C. Mol. Cell. Proteomics 2006, 5 (2), 306–312. (20) Cong, Y. S.; Fan, E.; Wang, E. Mech. Ageing Dev. 2006, 127 (4), 332–343. (21) Islinger, M.; Li, K. W.; Loos, M.; Luers, G.; Volkl, A. Eur. J. Cell Biol. 2005, 84, 121–123. (22) Hardt, M.; Witkowska, H. E.; Webb, S.; Thomas, L. R.; Dixon, S. E.; Hall, S. C.; Fisher, S. J. Anal. Chem. 2005, 77 (15), 4947–4954. (23) Zhou, H. L.; Ranish, J. A.; Watts, J. D.; Aebersold, R. Nat. Biotechnol. 2002, 20, 512–515. (24) Qiu, Y. C.; Sousa, E. A.; Hewick, R. M.; Wang, J. H. Anal. Chem. 2002, 74, 4969–4979. (25) Shi, Y.; Xiang, R.; Crawford, J. K.; Colangelo, C. M.; Horva`th, C; Wilkins, J. A. J. Proteome Res. 2004, 3, 104–111.

mercially available amino acid is attached to Tentagel S RAM beads, supplied with an acid-cleavable “Rink amide” linker, by Fmoc-chemistry and the same approach is followed to anchor the second amino acid, and then the attachment of an adipic acid follows. Finally, the N-terminal carboxylic function is activated as N-hydroxysuccinimidyl ester. The unique features of these reagents consist, therefore, in the low cost of a convenient laboratory-scale synthetic procedure. Free amino acids, glutathione, and peptides mixture obtained using different proteases on bovine serum albumin (BSA), serotransferrin, and lactotransferrin were tested in order to study the chemical reactivity of the SPIMT reagents, the linearity of quantification, and the effect of the amide tripeptide tags on the fragmentation pattern. Finally, the methodology was extended to a mixture of standard proteins prepared at different concentration levels. In all cases, the amino acids as well as peptide and peptide mixtures were differentially tagged with SPIMT and combined before matrix-assisted laser desorption ionization (MALDI) MS and MS/MS analysis. EXPERIMENTAL SECTION Materials. Solvents (CH3OH, CH3CN, CHCl3, CH2Cl2 (DCM), dimethylformamide (DMF), i-PrOH, AcOEt, Et2O, and H2O, HPLC grade), ammonium bicarbonate (NH4HCO3, pure g99.5%), piperidine, trifluoroacetic acid (HPLC grade, pure g99.0%), N,N′-dicyclohexylcarbodiimide (DIC), SDS (pure g99.0%), proteins (BSA, transferrin, and lactotransferrin), Fmoc-Gly-OH, Fmoc-Ala-OH, enzymes (trypsin, pepsin), R-cyano-4-hydroxytrans-cynnamic acid (R-CHCA, pure g99.0%), Tris (2-carboxyethyl)phospine hydrochloride (TCEP), 2-(1H-benzotriazole-1yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU), N,N-diisopropylethylamine (DIPEA), adipic acid, and Tris-HCl were purchased from Sigma Aldrich Fluka (Milano, Italy) and Tentagel S RAM resin from Rapp Polymer GMBH, Tubingen, Germany. Preparation of Standard. Bovine serum albumin (P02769), serotransferrin (P27425), and lactotransferrin (P24627) were used without further purification. The standard solution and standard protein mixtures were prepared in-house. To test the reactivity of the solid phase support with primary amines solutions of free amino acids, a synthetic peptide and protolytic mixture of BSA, serotransferrin, and lactotransferrin were prepared and then reacted. Preparation of Tryptic Digests. Stock solutions of BSA were prepared at concentrations of 1 mg/mL in 10 mM Tris-HCl, pH 8.8, 0.1% (w/v) sodium dodecyl sulfate (SDS) and 5 mM TCEP. This stock solution was heated to 100 °C for 5 min to denature the proteins and incubated with trypsin (1:50 enzyme/substrate ratio) for 18 h at 37 °C; the reaction was stopped by heating for 5 min. The resulting peptides mixture were freeze-dried and stored at -20 °C. Preparation of Peptic-Tryptic Digests. Stock solution of BSA was prepared at concentrations of 1 mg/mL in 0.2 N HCl and was incubated with pepsin (1:50 enzyme/substrate ratio) for 3 h at 37 °C. Then the sample was dried and dissolved in 10 mM Tris-HCl, pH 8.8, 0.1% (w/v) sodium dodecyl sulfate. The resulting solution was mixed with lyophilized trypsin to give 1:50 molar ratios of trypsin to protein. Digestion was performed for 18 h at Analytical Chemistry, Vol. 82, No. 13, July 1, 2010

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37 °C before quenching by heating for 5 min. The resulting peptides mixture was freeze-dried and stored at -20 °C. Preparation of Peptic Digests. A stock solution of BSA (serotransferrin and lactotransferrin) was prepared at a concentration of 1 mg/mL in 0.2 N HCl and was incubated with pepsin (1:50 enzyme/substrate ratio) for 18 h at 37 °C. The reaction was stopped by heating for 5 min. The resulting peptides mixture was lyophilized and then dissolved in 1 mL of Tris-HCl buffer (pH 8.8), 0.1% (w/v) SDS. Mixture 1 was BSA (1.66 mg, 24.8 nmol), TRAS (2.39 mg, 31 nmol), and LTF (0.85 mg, 10.9 nmol) dissolved in 6 mL of 0.2 N HCl. Mixture 2 was 1 mL of sample 1 diluted three times. Both samples were incubated with pepsin (1:50 enzyme/substrate ratio) for 18 h at 37 °C, and the reaction was stopped by heating for 5 min. The resulting peptides mixture were freeze-dried and stored at -20 °C. Preparation of Free Amino Acids Solution. A mixture of four amino acids (Ala, Lys, Phe, and Arg) was prepared in 10 mM Tris-HCl, pH 8.8, 0.1% (w/v) sodium dodecyl sulfate, such that each AA was present at 1.5 pmol/µL. Preparation of GSH Solution. A total of 1.25 mg of glutathione (GSH) peptide was dissolved in 1 mL of 10 mM TrisHCl, pH 8.8, 0.1% (w/v) sodium dodecyl sulfate (SDS). It was diluted with Tris buffer to a final concentration of 4 µM. Mass Tagging Protocol. To detect quantitative changes of peptides from two samples using solid-phase mass tagging reagents and mass spectrometry, the solid-phase support ASPIMT (1.5 mg in a 1.5 mL tube) and GSPIMT (1.5 mg in a 1.5 mL tube) were washed three times each with 200 µL of DCM and DMF. The supernatant was removed by spinning at 2 000 rpm for 4 min at each wash cycle. Each solid-phase support A SPIMT/GSPIMT was resuspended in 30 µL of DMF. The lyophilized peptide mixture obtained from the two samples to be compared was dissolved in 1 mL of Tris-HCl buffer (pH 8.8), 0.1% (w/v) SDS and then 200 µL of the resulting solutions were added to solid-phase support ASPIMT and GSPIMT. Then 10 µL of KOH (1 M) was added to each tube. The samples were incubated at room temperature for 3 h. After incubation, the solutions were removed and the beads were washed with DMF (200 µL × 2), CH3CN/H2O, 6:4, (v:v) (200 µL × 2), and DCM (200 µL × 2). The supernatant was removed by spinning at 2 000 rpm for 4 min at each wash. After washing, the beads were cleaved by incubating them with 200 µL of a 30% TFA in DCM solution for 1 h at room temperature (rt). Then the beads were washed with CH3CN/H2O, 6:4, (v:v) (200 µL × 2), and the solutions were collected. The organic solvents were evaporated using a speedvac device, and the resulting solutions were freeze-dried and stored at -20 °C. Peptide Reconstitution. Lyophilized ASPIMT- and GSPIMTlinked BSA (serotransferrin and lactotransferrin) peptic peptides (typically 3 nmol per vial) were suspended in 200 µL of 10% (v/v) acetic acid in water and were agitated on a vortex mixer to aid reconstitution. ASPIMT- and GSPIMT-linked BSA peptic peptide mixtures were mixed in various ratios: [ASPIMT]/[GSPIMT]) 1/1, 2/1, 3/1, 4/1, and 5/1. Each sample was diluted again in 10% (v/v) acetic acid in water to a final concentration of 1.5 pmol/µL. A volume of 1 µL of each sample was mixed with the matrix (R-cyano-4-hydroxycinnamic acid 5554

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(R-CHCA), 0.3% in TFA) in a 1:5 ratio. The sample/matrix mixture (0.5 µL) was loaded on a MALDI target plate (the total amount of BSA per spot was 125 fmol). Lyophilized ASPIMT- and GSPIMT-linked peptides prepared from the protein mixture (mixtures 1 and 2) were suspended in 200 µL of water and were agitated on a vortex mixer to aid reconstitution. Then the resulting solutions were mixed in the ratio 1:1. Sample was diluted again in 10% (v/v) acetic acid in water to a final concentration of 2 pmol/µL. A volume of 1 µL of each sample was mixed with matrix (R-CHCA, 0.3% in TFA) in a 1:5 ratio. The sample/matrix mixture (0.5 µL) was loaded on a MALDI target plate. Lyophilized ASPIMT- and GSPIMT-linked peptides prepared from the protein mixture (mixtures 1 and 2) were suspended in 200 µL of water and were agitated on a vortex mixer to aid reconstitution. Then the resulting solutions were mixed in the ratio of 1:1, and 1 µL was used for the LC-MS experiments. Lyophilized GSPIMT- and ASPIMT-linked GSH peptide (typically 0.4 nmol per vial) were suspended in 100 and 200 µL of 10% (v/v) acetic acid in water, respectively, and were agitated on a vortex mixer to aid reconstitution. ASPIMT- and GSPIMTlinked GSH peptide was mixed with matrix (R-CHCA, 0.3% in TFA) in a 1:5 ratio. The sample/matrix mixture (0.5 µL) was loaded on a MALDI target plate (the total amount of GSPIMTand ASPIMT-GSH per spot was 332 and 166 fmol, respectively). MALDI MS and MS/MS Analysis. A 1 µL portion of a premixed solution of each sample and R-CHCA (0.3% in TFA) was spotted on the matrix target, dried at room temperature, and directly analyzed by MALDI mass spectrometry. MALDI-time-offlight (TOF) analyses were performed using a 4700 Proteomics Analyzer mass spectrometer, from Applied Biosystems (Foster City, CA), equipped with a 200 Hz Nd:YAG laser at 355 nm wavelength. The MS spectra were acquired in reflectron mode (20 keV accelerating voltage), with 400 ns delayed extraction, averaging 2500 laser shots with a mass accuracy of 50 ppm. For MALDI-TOF/TOF analysis, collision induced dissociation (CID) was performed (a) at a collision energy of 1 kV, defined by the potential difference between the source acceleration voltage (8 kV) and the floating collision cell (7-6 kV), while the pressure inside the collision cell was (a) 8 × 10-7 Torr (CID OFF); (b) 1.3 × 10-6 Torr of air (CID ON). The CID spectra were obtained by summing 2500 single laser-shot spectra in both cases. CID was performed on those peaks identified as SPIMT-linked peptides with the signal-to-noise (S/N) ratio >20. The baseline of the CID mass spectra was corrected using ABI-4700 DataExplore software. After baseline correction, the relative intensity of the signal ions (m/z at 168 and 182 Da) were used for relative quantitation. Proteins were identified by searching a comprehensive protein database using Mascot programs (www.matrixscience.com). All MS/MS searches were performed using an initial mass tolerance of 50 ppm with enzyme cleavage specificity. LC-MS and MS/MS Analysis. An aliquot of SPIMT-linked peptides of [mixture 1]/[mixture 2] was injected into RP nanoLC system (LC Paking Ultimate with autosampler Famos WP). The chromatographic analysis was performed using a C18 column (80 Å pore, 75 µm i.d., 150 mm length), injecting a volume of 1 µL. The flow rate was 300 nL min-1 using the following

eluents: solvent A (H2O/ACN/TFA 95%/5%/0.1%), solvent B (H2O/ACN/TFA 2%/98%/0.1%). The [A]/[B] gradient was started from 100/0, changed 5/95 between 5 and 50 min, maintained at 5/95 between 50 and 60 min, than changed to 100/0 between 60 and 65 min and keep at 100/0 between 65 and 90 min. The online LC-MS MS/MS analysis was carried out with a hybrid Q-Star Pulsar-i (MSD Sciex Applied Biosystem, Toronto, Canada) mass spectrometer. As peptides elute from the nanocolumn into the mass spectrometer, scans are acquired in a datadependent manner. In this way, the mass spectrometer acquires one conventional scan over the m/z range 300-1600. Ions detected above a preset ion current threshold are automatically selected and subject to tandem mass spectrometry (MS/MS). Typically, two scans of MS/MS are recorded for each conventional scan; the first MS/MS scan represents the most intense multicharged ion (charge states 2-5) in the conventional scan mode, the second scan represents the second most intense multicharged ion (charge states 2-5). Dynamic exclusion is applied during the data-dependent acquisition, which prevents an abundant ion from being continually selected for MS/MS. Once selected, the peptide ion is not further selected for a period of time (60 s) so that other, less intense ions can be analyzed. RESULTS AND DISCUSSION SPIMT Reagents. The synthesis of small molecules on polymeric supports is a powerful method for the development of new molecules with a predetermined profile of properties.26 Solidphase peptide synthesis approach was adopted to devise efficient, cost-effective mass-tagging reagents. For this purpose, TentaGel S RAM resin supplied with an acid-cleavable “Rink amide” linker was chosen. The hydrophilic properties of TentaGel resins make them suitable for amine specific coupling reactions running in aqueous solution. A two-plex cost-effective mass-tagging reagent must be label free rather then labeled with low and/or high expensive isotopes and shaped by isobaric members such that all derivatized peptides are indistinguishable allowing the signal mass window tuning. In attempting to realize these reagents, we reasoned that simple regioisomeric dipeptides might provide a straightforward solution since they yield specific sequence ions following CID. Moreover, these dipeptides can easily be prepared in the laboratory by a routine Fmoc solid phase peptide synthesis. Vital to this kind of solid-phase mass tagging reagents is the anchoring of suitable groups that facilitate the attachment of the proteolytic peptides to the solid phase support. The use of an adipic acid derivative provides a smart solution since the intramolecular formation of a seven-membered ring is deprived by entropic effect27 and does not compete with the coupling reaction of the peptide. NHydroxysuccinimidyl ester was chosen as the carboxylic acid function activating group according to literature procedures.10,11 Clearly, this reactive functionality can easily react with different biological nucleophiles to be tagged. Among the different couples of SPIMTs, a suitable one was prepared from commercially available Fmoc-glycine and Fmoc-alanine avoiding any side-chain (26) Holmes, C. P.; Jones, D. G. J. Org. Chem. 1995, 60, 2318–2319. (27) Stieber, F.; Grether, U.; Waldmann, H. Angew. Chem., Int. Ed. 1999, 38, 1073–1077.

masking step (Scheme S1 in the Supporting Information). It is clear that using this strategy more than two tags can be generated choosing commercially available lower adipic homologues (i.e., C3, C2) or the 1,6-13C2 adipic acid, allowing this way the comparison of additional samples. A diagram showing the components of the two-plexed isobaric tagging chemistry is displayed in Scheme S2 in the Supporting Information. The complete tag-molecule consists of a reporter group (based on the couple of homologues 6-oxo-N-(1-oxopropan2-yl)hexanamide and 6-oxo-N-(2-oxoethyl)hexanamide), a mass balance group (based on the homologues 2-aminoacetamide and 2-aminopropanamide), and a peptide-reactive group (OSu ester). Regioisomeric dipetide moieties keep the overall mass of the reporter and balance components constant. The reporter groups lead to ions of m/z 168.2 and 182.2 Da, while the balance groups generate species of 86.0 and 72.0 Da, such that the combined mass remains constant for each one of the two reagents (Scheme S2 and Figure S10 in the Supporting Information). The tag after reaction with a peptide forms an amide linkage with any N-terminal peptide amine group; these amide linkages give similar fragmentation to backbone peptide bonds when subjected to CID. Because of the fragmentation of the tag amide bond, the balance moiety is lost as a neutral, while the charge is retained by the reporter group fragment. A mixture of two identical peptides, each one tagged with one SPIMT reagent of the two-plex set, appears as a single, unresolved precursor ion in MS (identical m/z). Upon CID conditions, the two reporter group ions appear as distinct masses (168.2-182.2 Da). All the other sequence-informative fragment ions (b-, y-, etc.) remain isobaric, and their individual ion current signals (signal intensities) are additive. The adipamide residue can be considered as pseudo amino acid to perform de novo sequencing, therefore each CID spectrum was analyzed by introducing “J” as a “friendly” form for the adipic acid linker. The relative concentration of the peptides was thus deduced from the relative intensities of the corresponding reporter ions and the quantification was thus performed at the MS/MS stage. The SPIMT approach is similar in principle to other peptide labeling techniques and enjoys the same features as these other approaches, with some additional advantages. Pairs of SPIMT tagged peptides are chemically identical, but unlike other isobaricisotope tags, the SPIMTs are also isobaric and lead to a more accurate quantification. The novelty of the SPIMT strategy relies on a CID-based differentiation of regioisomeric species useful for quantification of tagged analytes. The untagged materials are removed by simple washing of the solid-phase support, remarkably improving data quality. Furthermore, the samples collected after solid-phase release can be analyzed by MALDI MS and MS/MS without any additional chromatographic step. Performance of SPIMTs. The reactivity of SPIMT reagents was initially tested with a mixture of free amino acids (lysine, arginine, alanine, and phenylalanine) under different experimental conditions. The reactivity of amino acid succinimidyl esters toward side-chain and N-terminal amino functions is pH dependent.28 Thus the reaction conditions for SPIMT-OSu coupling in aqueous buffers were optimized. At pH 8.8 (Tris-buffer), the coupling reaction occurred on the side-chain only of the amino group of (28) Mao, S.-Y. The Protein Protocols Handbook, 2nd ed.; Walker, J. M., Ed.; Humana Press: Totowa, NJ, 2002; pp 351-352.

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Figure 1. MS/MS spectra of SPIMT-GSH reagent (A) ASPIMT-GSH and (B) GSPIMT-GSH.

Lys (ε-ΝΗ2 pKa ) 10.6); whereas Phe, Ala, and Arg (pKa 9.24, 9.87, 9.04, respectively) were not anchored to the supports under this condition. Conversely, at pH 10.0 only the N-terminal function of all the four amino acids was tagged. The reactivity of an amino group strongly depends on its ionization state. The protonation decreases the nucleophilicity of a species, and the pH of the medium affects the rate of many nucleophilic reactions. The relationship between protonation and pH depends on the pKa of the nucleophile. At fixed pH, the most reactive group is usually the one with the lowest pKa,29 because the residue (Lys) possessing a higher pKa value is prone to undergo protonation and Schiff base formation more easily.29,30 Therefore pH 10.0 represents the condition for the optimal balance between deprotonation of the amino group, nucleophilic attack, and hydrolysis reaction of the NHS ester with water. Running the coupling reaction at pH 10.0 prevents the unwanted reactions for peptides containing amino acid residues with side-chain amino function, allowing for a site specific N-terminal tagging. These experiments were performed with a large excess of solid-phase reactive sites compared to the sample dimension. Although using an excess of the tagging reagent increases the potential for unwanted side reactions with cysteine and/or lysine containing peptides, the simple test of the free amino acids, which are small molecules and can be easily attached to the solid-phase (29) Wong, S. S. Chemistry of Protein Conjugation and Cross-Linking; CRC Press: Boca Raton, FL, 1991; pp 12-14. (30) Weber, H. K.; Zuegg, J.; Faber, K.; Pleiss, J. J. Mol. Catal., B: Enzym. 1997, 3, 131–138.

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support, showed as that the unwanted lysine side-chain reactions are pH dependent. Furthermore, in order to test the ability of the mass tags to selectively recognize N-terminal amino function at pH 10.0, a sample of GSH, a cysteine-containing natural peptide, was checked by mass spectrometry. The reaction product, formed by incubation of 0.4 nmol of GSH with the SPIMT reagents, was cleaved by acidolysis and prepared for the analysis as described in the Experimental Section. An aliquot of each mass tagged peptide corresponding to 166 and 332 fmol of the original GSH peptide sample was analyzed by MALDI MS/MS as shown in Figure 1A,B. SPIMTs add 255.1 Da to the parent mass of the GSH peptide to give a signal of m/z 563.2 in the MALDI-TOF mass spectrum. A few experiments were performed in order to determine the time needed for the coupling reaction and showed that the complete conversion of peptide was obtained in 1 h. To ensure a completeness of the functionalization on solid phase-support, an incubation time of 3 h was selected for all subsequent experiments. The experiments on the fragmentation of the SPIMT-GSH derivatives in the MALDI TOF/TOF instrument showed that the coupling reaction occurred only at the N-termini. Comparison of the CID spectra (1 keV) from GSH tagged with SPIMTs showed clearly that the introduction of the AGJ/GAJ residue at the N-termini led to the expected fragmentation pattern. The relative intensities of y- and b-type sequence ions indicated that the fragmentation pattern of GSH was changed slightly by SPIMT conjugation. More importantly, a number of new peaks appeared

Figure 2. MS/MS spectra of SPIMT-linked peptic/tryptic peptide of m/z 1274.52 from BSA acquired at (A) low collision energy (1 kV, 8 × 10-7 Torr) and (B) high collision energy (1 kV, 1.3 × 10-6 Torr).

in the spectra including b3 (tag, m/z 256.1) and y6 ions. The quantitation signals appeared at 168.2 and 182.2 Da, respectively. The b3 ion at m/z 256.1 Da served as the signature for SPIMT conjugation, and the concomitant appearance of the b5 and y6 ions indicated further evidence of the coupling. Thus, SPIMTs offered the quantitation signals without losing sequence information. The intensity of the desired SPIMT fragments, reporter ions, was independent from the amino acid sequence of the peptide, and at 1 keV collision energy the SPIMT fragments reflected the abundances of the tagged peptides accurately. These data indicated that the mass tagging reagents can accurately tag and report peptides ratios in an artificial peptide mixture. To ensure the suitability of the SPIMTs for quantitative measurements in more complex mixtures, the ability of the mass tags to react specifically with the N-terminal amino function of peptides and to accurately detect relative concentrations from the two mixtures of peptide digests, we therefore tested the reactivity of SPIMT reagents toward tryptic digests of BSA. The samples were prepared as follows. The same amount of BSA tryptic digest (200 µL, 3 nmol of the original sample) in Tris buffer (pH 10.0) was mixed with 1.5 mg of ASPIMT and GSPIMT beads, respectively. The suspensions were incubated separately for 3 h at room temperature and then further processed as previously described. The MALDI-TOF mass spectrum after mass tagging showed only a few ions of tagged peptides obtained from the protein digest mixture, whereas MALDI-TOF mass spectrum of the complete peptide mixture before mass tagging displayed numerous peptide peaks. The comparison of the two MALDI spectra suggested that the complexity of mass spectrum of the peptide mixture was greatly reduced after mass tagging, only six singly tagged tryptic peptides were detected. Furthermore, the absence of protonated species differing from each other of

255.1 Da suggests that no double tagging occurred in the above-mentioned experimental conditions. The untagged peptides were collected during the workup of the coupling reaction and analyzed by MS. The ratio of the number of tagged and untagged tryptic peptides indicated that SPIMT reagents did not work efficiently. This may be particularly troublesome in complex mixtures containing low-abundance proteins. Furthermore the number of tagged peptides may not be enough for complete protein identification. In attempting to overcome this problem, we tested the reactivity of SPIMT reagents toward proteolytic peptides of BSA obtained by two consecutive digestions. Therefore, a peptide mixture prepared by pepsin/ trypsin digestion (PT-BSA) (see the Experimental Section) was treated with SPIMT reagents. The same amount of BSA peptic/ tryptic digest (200 µL, 3 nmol of the original sample) in Tris buffer (pH 10.0) was mixed with 1.5 mg of ASPIMT and GSPIMT beads, respectively. The complexity of the mass spectrum of the peptide mixture was reduced after mass tagging. Only Nterminal tagged peptide are shown in the mass spectra (Figure S1A,B in the Supporting Information). The obtained spectra from two members of the two-plex are superimposable, suggesting that the reactivity of the two solid-phase support is identical. Moreover, these data are indicative for a highly specific coupling reaction toward lypophilic peptide mixtures. The performance of SPIMTs on the relative quantitation was then evaluated. The mixtures of ASPIMT- and GSPIMT-PT-BSA were prepared in various ratios, and 125 fmol was loaded on a MALDI spot for CID. In Figure 2A,B are reported the MALDI MS/MS spectra of m/z 1274.52 acquired at low and high collision energy, respectively. Figure 2A exhibits the ion at 256.1 Da used as a signature for SPIMT conjugation, and the concomitant appearance of the ion of m/z 1019.3 ([M-SPIMT + H]+, yn-3) confirmed again the Analytical Chemistry, Vol. 82, No. 13, July 1, 2010

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Figure 3. (A-E) MS/MS spectra of ASPIMT IARRHPYF/GSPIMT-IARRHPYF from the BSA peptic digest premixed in various ratios. The region of quantitative signals is displayed. (F) Plot of the measured vs expected [ASPIMT]/[GSPIMT] ratio (y ) 0.8902x + 0.4185; R2 ) 0.994) obtained from MS/MS spectra of ASPIMT-IARRHPYF/GSPIMT-IARRHPYF of the BSA peptic digest premixed in various ratios.

conjugation. Figure 2B exhibits variations in the relative intensity of the quantitation signals at m/z 168.2 and 182.2 Da as a function of the fixed 2:1 premixing. The relative intensities of 168.2 and 182.2 linearly increased with the relative amount of the SPIMTPT-BSA mixture. Furthermore, we tested the ability of the mass tags to react specifically with N-terminal amino function of lipophilic peptides and to accurately detect relative concentrations from two mixtures of peptide digests (see Figure S2A,B in the Supporting Information). To further demonstrate the sensitivity and the specificity of this approach for real proteomics applications, two samples containing different amounts of BSA peptic digests were mixed with a large excess of ASPIMT and GSPIMT beads, respectively. So 3 nmol of peptic BSA digest in Tris buffer (pH 10.0) was treated with 6.73 mg of GSPIMT bead, whereas 6 nmol of peptic BSA digest in Tris buffer (pH 10.0) was treated with 3.65 mg of ASPIMT bead. The suspensions were incubated separately for 3 h at room temperature and then further processed. Relative quantitatation was performed loading 1 µL of the premixed tagged peptides under CID conditions of the precursor ion of m/z 1314.72. The data obtained are shown in Table S1 in the Supporting Information. By application of the relative intensities of the two reporter ions (168.2 and 182.2 Da), the deduced composition of the mixture was found to be ASPIMT (168)/GSPIMT (182) 70:30 (%). This result suggests that an experimental error of 3% can be performed using a 4.5-fold excess of beads. Furthermore, the mixtures of ASPIMT- and GSPIMT-peptic BSA digest were prepared in various ratios, and 125 fmol was loaded on a MALDI spot for CID. Samples at each ratio were prepared in triplicate. Figure 3A-E exhibits variations of the reporter signals (168.2 and 182.2 Da) on going from 1:1 to 5:1 relative functionalized 5558

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peptide ratios. Relative quantitation was performed following CID of the precursor ion of m/z 1314.72. As the relative amount of A SPIMT-BSA is increased, the relative intensities of reporter signal increase linearly. The results are reported in Figure 3F and show that the mass tags are able to accurately detect any relative concentration differences of peptide in two solutions over a 5-fold concentration range. The linear response over this range suggests that SPIMT reagents should be useful for measuring such ratios in more complex mixtures. Furthermore, SPIMTs worked well in the peptide concentration range of 100-300 fmol with good quantitation linearity, without losing sequencing information. Quantitation and Identification. Since this method ensures that all peptides were labeled prior to MS analysis, the direct submission of MS and MS/MS data to the database for protein identification and automatic sequencing should not be a critical step. The SPIMT reagents adds 255.1 Da to the parent mass of each proteolitic peptide, and this conjunction occurs only on the N-terminal amino function. Introducing “AGJ” as a “friendly tag form” for the N-terminal- and lysine variable-modification on the Mascot Search Database Form, the direct submission of MALDI MS data for protein identification can be easily performed. MALDI MS spectra were evaluated using MASCOT database searching. Searches were performed against the SWISS PROT or NCBI database, with the restricted taxonomy and pepsin enzyme cleavage specificity. Accordingly, the direct submission of MALDI MS data allowed the identification of serotransferrin, BSA, and lactotransferrin with the highest MOWSE score (see Figures S4-S6 in the Supporting Information). By considering that the MS/MS spectra contained additional peaks arising from the fragmentation of the tag, some extra instructions were introduced in the Mascot “modification file”, e.g., N-termini SPIMT modification must be allowed, and that the fragments arising from the fragmentation of the tag itself must be ignored (see section B in

Figure 4. MS/MS spectrum of ASPIMT-RYYGYTGA peptide of m/z 1205.6 from serotransferrin, acquired at low collision energy (1 keV, 8 × 10-7 Torr). Immonium ions are denoted in parentheses, and the internal fragments are indicated with the asterisks (*).

Figure 5. MS/MS spectrum of GSPIMT-RYYGYTGA peptide of m/z 1314.7 from BSA, acquired at low collision energy (1 keV, 8 × 10-7 Torr). Immonium ions are denoted in parentheses, and the internal fragments are indicated with the asterisks (*).

the Supporting Information). Consequently, the direct submission of MS/MS data to the database allowed the identification of proteins in good protein scores even if the fragmentation pattern of each peptide is slightly changed by SPIMT conjugation (Figure S3 in the Supporting Information and Figures 4 and 5). Several serotransferrin, BSA, and lactotransferrin peptides were identified with the highest MOWSE score (Figures S7-S9 in the Supporting Information). The β-cleavage promoted by the N-terminal amide residue yielded regioisomeric y- and b-type ions. The adipamide residue was considered as a “pseudo” amino acid. Accordingly it was introduced in the data dictionary as a “pseudo Glu” J with the elemental composition C6H9NO2 (exact mass, 127.06333) and side-chain formula H. In this fashion, Data Explorer software was used to automatically confirm the sequences of SPIMT-

linked peptides by the Peptide Ion Fragmentation Tool box (Figures S12-S14 in the Supporting Information). The performance of SPIMTs for protein quantitation was further tested with two mixtures of peptide digests. The mixtures contained peptic digests of BSA, serotransferrin, and lactotransferrin. The samples were prepared as follows. Mixture 1, containing 826 pmol of BSA, 1.0 nmol of serotransferrin, and 363 pmol of lactotransferrin, was mixed with 1.5 mg of ASPIMT bead in 200 µL of Tris buffer, pH 10.0. Mixture 2 was prepared by diluting mixture 1 three times and was treated with 1.5 mg of G SPIMT bead in 200 µL of Tris buffer, pH 10.0. The suspensions were incubated separately for 3 h at room temperature and then further processed as described. After coupling with SPIMT, the total amount of the 3:1 mixture loaded was 167 fmol. MALDI-TOF mass spectrum showing only Analytical Chemistry, Vol. 82, No. 13, July 1, 2010

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Figure 6. MALDI MS spectrum of peptide mixtures resulting from pepsin digestion of mixture 1/mixture 2 after SPIMT labeling. The peaks labeled with 9, 2, b represent identified peptide from serotransferrin, BSA, and lactotransferrin, respectively.

tagged peptides from the protein digest mixture is reported in Figure 6. The relative quantitation results are presented in Figure S11 in the Supporting Information (see Table S2 in the Supporting Information). The relative average ratio determined by SPIMT was 3.1 ± 0.04 for [mixture 1]/[mixture 2] agreed well with the composition of the prepared mixture. Although, SPIMT methodology was designed for LC-MALDI TOF/TOF experiments, the newly introduced quantitation method was also tested in RP nano LC-MS and MS/MS, using a 1:1 mixture of [mixture 1] and [mixture 2]. The MS/MS spectrum of the triple charged ion parent of m/z 438.91 Da (SPIMTIARRHPYF) is reported in Figure S15 in the Supporting Information. The concomitant presence of the reporter signals (168.2 and 182.2 Da) and y11-NH3 ion confirmed the N-terminal coupling. Furthermore, the appearance of two ion couples, of m/z 414.2 (3+), 620.8 (2+) and of m/z 409.5 (3+), 613.8 (2+), that differ by 14 Da from each other and arise from the loss of the balance group (72.0 and 86.0 Da) suggest the presence of the two regioisomeric SPIMT-linked peptides. It can be confidentially assumed that the SPIMT approach to simultaneous protein identification and assay is of general use. CONCLUSIONS SPIMTs are unlabeled cost-effective mass-tagging reagents able to form an amide linkage with any N-terminal peptide amine group. These amide linkages fragment in a similar fashion to the backbone peptide bonds under CID conditions, leading also to two signature ions. A mixture of two identical peptides, each one 5560

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tagged with one SPIMT reagent of the two-plex set, appears as a single unresolved precursor ion in MS. Upon CID conditions, the two reporter group ions appear at distinct masses, and all the other sequence-informative fragment ions remain isobaric. SPIMTs provide specific quantitation signals in a region of the MS/MS spectra usually free from the other ions at the intensity level comparable to the peptide sequence ions. Furthermore, the direct submission of MS and MS/MS data to the database for protein identification and automatic sequencing is not a critical step. Accordingly, protein identification is easily performed by Mascot Database Search using peptide mass fingerprint and MS/MS data. The novelty of the SPIMT strategy relies on a CID-based differentiation of regioisomeric species useful for quantification of tagged analytes. ACKNOWLEDGMENT This paper was supported by the University of Calabria. SUPPORTING INFORMATION AVAILABLE Results from Mascot Database Search Peptide Mass Fingerprint and MS/MS data, quantitation results, and MS/MS spectra of SPIMT-linked peptic peptides. This material is available free of charge via the Internet at http://pubs.acs.org.

Received for review February 16, 2010. Accepted May 25, 2010. AC1004212