Deuterium Isobaric Amine-Reactive Tags for Quantitative Proteomics

Aug 17, 2010 - This paper demonstrates the applications of a novel isobaric reagent, named deuterium (2H) isobaric amine-reactive tag (DiART), for qua...
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Anal. Chem. 2010, 82, 7588–7595

Deuterium Isobaric Amine-Reactive Tags for Quantitative Proteomics Junxiang Zhang,† Yan Wang,‡ and Shuwei Li*,†,§ Institute of Bioscience and Biotechnology Research, University of Maryland, 9600 Gudelsky Drive, Rockville, Maryland 20850, Proteomics Core Facility, and Department of Chemistry and Biochemistry, University of Maryland, College Park, Maryland 20742 This paper demonstrates the applications of a novel isobaric reagent, named deuterium (2H) isobaric aminereactive tag (DiART), for quantitative proteomics. Peptides labeled with DiART were analyzed using an electrospray ionization (ESI)-based LTQ-Orbitrap mass spectrometer. Our data showed that 2H-associated isotope effects, such as partial loss of 2H labels during tandem mass spectrometry (MS/MS) and 2H-related chromatographic shift, were either not observed or negligible. With the use of a hybrid collision-induced dissociation (CID)-higher energy C-trap dissociation (HCD) acquisition method, we were able to identify DiART-labeled peptides with high confidence and quantify them with high accuracy. Furthermore, we adopted a hybrid electron-transfer dissociation (ETD)HCD acquisition protocol and developed a novel data analysis approach to measure phosphorylation of peptides. Our results showed DiART had excellent performance on LTQ-Orbitrap instruments and provided a cost-effective technique for large-scale quantitative proteomics measurements. Quantitative proteomics, or the large-scale quantitative analysis of proteins in biological samples, has been advancing rapidly. Particularly, mass spectrometry (MS)-based methodologies have become more widely adopted for comprehensive determination of proteins from highly complex biological samples, providing powerful solutions for biomarker discovery and subsequent target validationsfordevelopingnewclinicaldiagnosticsandtherapeutics.1,2 In comparison to other biochemical methods, however, MS itself is not ideal for quantitative analysis because its signal intensity is strongly affected by factors such as matrix effect and physicalchemical properties of analytes. To solve this problem, one needs to adopt more sophisticated approaches by combining MS with other strategies, such as stable isotope labeling or label-free methods based on various statistical models. Label-free quantification, as its name suggests, does not involve stable isotope labeling; thus it requires a highly reproducible * To whom correspondence should be addressed. Fax: 240-314-6225. E-mail: [email protected]. † Institute of Bioscience and Biotechnology Research. ‡ Proteomics Core Facility. § Department of Chemistry and Biochemistry. (1) Ong, S. E.; Mann, M. Nat. Chem. Biol. 2005, 1, 252–262. (2) Yan, W.; Chen, S. S. Briefings Funct. Genomics Proteomics 2005, 4, 27–38.

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chromatography profile and stable MS signals, which, in turn, makes it incompatible with very complex samples.3 In contrast, stable isotope labeling approaches, despite their requirements for additional labeling steps and cost of reagents, can provide more accurate quantitative information and allow multiple samples to be measured concurrently. Therefore, stable isotope labeling strategies have been widely used in quantitative proteomics and numerous approaches have been developed over the past decade.4-8 These methods, based on how peptides are labeled and quantified, can be divided into two main categories, massdifference and isobaric tags. Mass-difference approaches introduce a small mass difference to identical peptides from two or more samples, so they can be distinguished in the MS spectrum, and their peak areas are integrated for quantification. Examples of mass-difference approaches include stable isotope labeling with amino acids in cell culture (SILAC)9,10 and isotope-coded affinity tags (ICAT).11 Isobaric tags, on the other hand, use a different concept for peptide quantification.12,13 Isobaric reagents are usually a set of molecules with identical structures that consist of a reporter, a balancer, and a reactive group. Stable isotopes are incorporated at multiple positions so that the reporter region in each reagent differs in mass. Yet this difference is compensated by the balancer so that the molecular weights of all isobaric reagents in a set are exactly the same. As a result, isobaric tag labeled peptides do not exhibit mass shift in a set of differentially labeled samples. Instead, they appear to be a single peak in the mass spectrum. After parent ions are isolated and fragmented in a subsequent tandem mass spectrometry (MS/MS) measurement, (3) Podwojski, K.; Eisenacher, M.; Kohl, M.; Turewicz, M.; Meyer, H. E.; Rahnenfuhrer, J.; Stephan, C. Expert Rev. Proteomics 2010, 7, 249–261. (4) Julka, S.; Regnier, F. E. Briefings Funct. Genomics Proteomics 2005, 4, 158– 177. (5) Zeng, D.; Li, S. Bioorg. Med. Chem. Lett. 2009, 19, 2059–2061. (6) Li, S.; Zeng, D. Chem. Commun. 2007, 2181–2183. (7) Zhou, H.; Ranish, J. A.; Watts, J. D.; Aebersold, R. Nat. Biotechnol. 2002, 20, 512–515. (8) Schmidt, A.; Kellermann, J.; Lottspeich, F. Proteomics 2005, 5, 4–15. (9) Kruger, M.; Moser, M.; Ussar, S.; Thievessen, I.; Luber, C. A.; Forner, F.; Schmidt, S.; Zanivan, S.; Fassler, R.; Mann, M. Cell 2008, 134, 353–364. (10) Ong, S. E.; Blagoev, B.; Kratchmarova, I.; Kristensen, D. B.; Steen, H.; Pandey, A.; Mann, M. Mol. Cell. Proteomics 2002, 1, 376–386. (11) Gygi, S. P.; Rist, B.; Gerber, S. A.; Turecek, F.; Gelb, M. H.; Aebersold, R. Nat. Biotechnol. 1999, 17, 994–999. (12) Dayon, L.; Hainard, A.; Licker, V.; Turck, N.; Kuhn, K.; Hochstrasser, D. F.; Burkhard, P. R.; Sanchez, J. C. Anal. Chem. 2008, 80, 2921–2931. (13) 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.; Bartlet-Jones, M.; He, F.; Jacobson, A.; Pappin, D. J. Mol. Cell. Proteomics 2004, 3, 1154–1169. 10.1021/ac101306x  2010 American Chemical Society Published on Web 08/17/2010

they will yield a series of reporter ions with slightly different mass which can be used for relative quantification of the parent ions or the relative abundance of the same peptide in different samples. This isobaric tag strategy offers two major advantages: (1) it facilitates multiplex and high-throughput analysis of multiple samples (up to eight samples can be analyzed in one experiment), and (2) the detection sensitivity is often improved as the same peptides from different samples contribute to a single peak. Currently, there are two commercially available isobaric reagents, tandem mass tag (TMT) from Thermo Scientific and isobaric tag for relative and absolute quantification (iTRAQ) from AB Sciex, which have more or less the same functionality and performance. TMT and iTRAQ reagents, however, are costly because they are coded with 13C and 15N and require rather tedious synthesis. For example, six-plex TMT reagents are synthesized with a 15-step scheme and the overall yield is less than 1%.14 This imposes a significant limiting factor that prevents many researchers from adopting this technique. In addition, these multiple-step syntheses can also introduce undesirable isotope impurity, so each batch of iTRAQ reagents is accompanied with a correction factor table to take isotope impurity into account.15 This, in turn, complicates subsequent data analysis. One possible approach to shorten these lengthy syntheses is using 2 H as one of the heavy isotopes. But 2H is not employed in TMT and iTRAQ tags because 2H often introduces adverse 2H isotopic effects, such as partial loss of 2H labels during MS/ MS and 2H-associated chromatographic shift, which compromise the reliability and reproducibility of quantitative measurement and result in poor quality of data in comparative proteomics analysis.16 Fortunately, 2H isotope effects can be controlled by using minimum number of 2H and placing them on hydrophilic groups.17,18 Following those rules, we previously conceptualized a novel type of 2H-labeled isobaric tags, named deuterium isobaric amine-reactive tag (DiART), in order to provide the proteomics society an affordable isobaric tag reagent with high isotope purity. These reagents use N,N′dimethylleucine as the reporter group and are capable of measuring up to six protein samples concurrently when employed on a matrix-assisted laser desorption ionization tandem mass spectrometer (MALDI-MS/MS).19 More recently, a set of similar four-plex isobaric tags (DiLeu) were reported, which also use 2H labeling and N,N′-dimethylleucine reporter.20 However, DiLeu uses a nontraditional activation chemistry to label peptides and requires a special reaction condition. In contrast, DiART, TMT, and iTRAQ share the same group, N-hydroxysuccimide ester (NHS), for peptide coupling, so their labeling protocols are identical, making it easy for TMT/iTRAQ users to adopt the DiART technique. To broaden DiART applications, we demonstrate here the performance of DiART labeling on an electrospray ionization (ESI)-based MS instru(14) Hamon, C.; Schwarz, J.; Becker, W.; Kienle, S.; Kuhn, K.; Schafer, J. International Patent WO2007/012849, 2007. (15) Dey, S.; Pappin, D. J. C.; Purkayastha, S.; Pillai, S.; Coull, J. M. U.S. Patent US2005/0148774, 2005. (16) Zhang, R.; Regnier, F. E. J. Proteome Res. 2002, 1, 139–147. (17) Zhang, R.; Sioma, C. S.; Thompson, R. A.; Xiong, L.; Regnier, F. E. Anal. Chem. 2002, 74, 3662–3669. (18) Regnier, F. E.; Zhang, R. U.S. Patent US7449170, 2008. (19) Zeng, D.; Li, S. Chem. Commun. 2009, 3369–3371. (20) Xiang, F.; Ye, H.; Chen, R.; Fu, Q.; Li, L. Anal. Chem. 2010, 82, 2817– 2825.

ments, such as an LTQ-Orbitrap, which is now predominantly used in the proteomics community. EXPERIMENTAL SECTION Materials. All proteins and chemicals were purchased from Sigma-Aldrich (St. Louis, MO). DiART reagents were synthesized and prepared with slight modifications (data not shown) from what we previously described.19 Labeling of a Three-Protein Mixture. Bovine catalase (100 µg), horse myoglobin (10 µg), and bovine serum albumin (1 µg) were dissolved in 1 mL of denaturing/reducing solution (8 M urea, 50 mM sodium borate buffer, pH ) 8.3, 5 mM tris(2-carboxyethyl)phosphine (TCEP)) and incubated at 37 °C for 30 min. Then, bromoacetamide (20 mM) was added to alkylate free cysteine residues. The proteins were precipitated with acetone, dissolved again in 200 µL of buffer (200 mM sodium borate buffer, pH ) 8.3, 0.8 M urea), and then digested with trypsin (10 µg) at 37 °C overnight. Six aliquots of the protein mixture (10 µL each) were mixed with each of the six DiART reagents (10 µL of 10 mM in acetonitrile), respectively, and the reaction was incubated at room temperature for 2 h. The reaction was stopped by adding 10 µL of 5% ethanolamine solution. All of six samples were then mixed together by a predetermined ratio (1:1:1:1:1:1, 5:2:5:2:1:1, or 5:1: 2:20:50:100), dried in a SpeedVac, and then dissolved in 5% acetonitrile containing 0.1% formic acid. Labeling of β-Casein and Enrichment of Phospho-Containing Peptides. β-Casein (100 µg) was dissolved in 100 µL of sodium borate buffer (200 mM, pH ) 8.3) and digested by trypsin (2 µg) at 37 °C overnight. Six aliquots of the digest (15 µL each) were mixed with each of the six DiART reagents (15 µL of 10 mM in acetonitrile), and the reaction mixture was incubated at room temperature for 2 h. The reaction was stopped by adding 15 µL of 5% ethanolamine solution. All of six samples were then mixed together by a predefined ratio (1:0.5:2:1:0.25:7). This peptide mixture was acidified to pH 3 by adding 1 M HCl solution and incubated with 50 µL PHOS-Select iron affinity gel for 30 min. After the supernatant was removed and the gel was washed with 100 mM acetic acid solution containing 30% acetonitrile 500 µL × 3 times, the bound peptides were eluted with 400 mM ammonia solution, immediately acidified with 1 M HCl solution to pH 2, and stored at -20 °C until MS analysis. LC-MS/MS. Labeled peptides were separated by reversedphase chromatography on a Shimadzu Prominence nano highperformance liquid chromatograph (HPLC) that is coupled to a Thermo LTQ-Orbitrap MS analyzer. After injection, peptides were loaded into a LC Packings (Sunnyville, CA) C18 PepMap precolumn (0.3 mm × 5 mm) at 10 µL/min. After 15 min of loading and desalting with 100% solvent A (0.1% formic acid and 5% acetonitrile), peptides were then eluted into a Michrom (Auburn, CA) Magic C18AQ column (3 µm, 200 Å, 0.1 mm × 150 mm) and chromatographically separated using a binary solvent system consisting of A and B (0.1% formic acid and 95% acetonitrile) at a flow rate of 400 nL/min. A gradient was run from 10% B to 45% B over 30, 60, or 120 min, followed by a 5 min linear gradient to 80% B and a 5 min wash step with 80% B. The column is then equilibrated at 5% B for 10 min before the next sample was injected. The LTQ-Orbitrap XL mass spectrometer (Thermo Electron, San Jose, CA) was operated in positive ion mode with dataAnalytical Chemistry, Vol. 82, No. 18, September 15, 2010

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dependent MS/MS acquisition. The instrument was set to complete a full mass scan from mass-to-charge ratio (m/z) range of 400-2000 with resolution of 60 000 (m/z ) 400) in the Orbitrap followed by data-dependent MS/MS analysis of up to three most intense ions with higher energy C-trap dissociation (HCD) in the Orbitrap with resolution of 7500 at m/z 400, and the same three ions with collision-induced dissociation (CID) in the linear ion trap at unit mass resolution. Peaks eluting from the LC column that have ions above 10 000 arbitrary intensity units and charges higher than 1 triggered the ion trap to isolate the ion and perform an MS/MS experiment scan after the MS full scan. HCD collision energy was set at 42% and CID at 35%. For standard proteomics experiments that require high sensitivity of peptide detection, dynamic exclusion was turned on to exclude ions from being selected again for MS/MS analysis for 30 s with a m/z window of -0.5 to +1.5. For evaluation of potential chromatography separation of the same peptide labeled with different reagents caused by isotopic effect from 2H labeling, dynamic exclusion was turned off so that peptides would be selected for MS/MS analysis repeatedly as long as it was one of the three most intense ions in the fullscan MS. In addition, two peptides were selected from the initial analysis and added to parent list so that they were selected for MS/MS analysis as long as they were detected in the full-scan MS spectrum, regardless of their relative intensity. The two peptides were selected to represent an early eluting peptide and a late eluting peptide with low relative intensity. Data Analysis. To make a Mascot server (version 2.3) compatible with DiART labeling, two configuration files (mod.xml and quantitatio.xml) on the server were modified by treating DiART tags as a special posttranslational modification. For the hybrid CID-HCD data set, MS acquisition files were processed with Proteome Discoverer 1.1 (Thermo Scientific) to combine the HCD and CID spectra of the same parent ion. The m/z and intensity of all parent ions and their associated fragment ions were saved as a mascot generic format file (suffix .mgf). This file was used by the customized Mascot server for peptide identification and quantification. For the hybrid ETD-HCD data set, electron-transfer dissociation (ETD) and HCD spectra were extracted separately and saved as two separate mascot generic format files by Proteome Discoverer 1.1. A Perl script was written and used to process and combine these two files into a single file as described in the Result and Discussion, which was then submitted to the customized Mascot server for peptide identification and quantification. RESULTS AND DISCUSSION Design of DiART Reagents and Their Compatibility with ESI-Based Instruments. DiART reagents are a set of six chemically identical molecules (Figure 1) sharing a reporter, a balancer, and an amine-reactive group but containing different stable isotopes at multiple positions. As a result, the mass of the reporter in each reagent differs, yet this difference is offset by the balancer to keep the total molecular weight of all six reagents the same. When up to six samples are labeled with DiART reagents and analyzed by MS, differentially labeled peptides would not display any mass difference. Only after a parent ion is fragmented, it yields a series of reporter ions within the m/z range of 114-119 for peptide quantification, in addition to the commonly 7590

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Figure 1. Structure of DiART reagents. When peptides labeled with these tags are fragmented in MS/MS, they yield strong reporter ions ranging from 114 to 119. Positions containing heavy stable isotopes are illustrated.

observed b- and y-ions that can be used for peptide sequence identification. Therefore, DiART reagents are similar to iTRAQ and TMT in principle. However, the main advantage of DiART is that they can be prepared from readily available starting materials within five synthetic steps at high isotopic purity (>97%) and yield (∼40%, overall yield), greatly reducing their production cost. And therefore, DiART reagents will make it affordable for the scientific society to work on large sample sets to obtain quantitative proteomics analysis with statistically relevant replicates. Like other commercially available isobaric tags that rely on reporters generated from MS/MS for quantification, DiART reagents must satisfy two requirements for practical uses: (1) the incorporation of DiART tags should not impair the fragmentation pattern of the labeled peptides, and (2) the production of reporter ions should be efficient and show high signal-to-noise ratio (S/ N). To meet these criteria, we designed the DiART reagents based on a well-established chemical reaction, N,N′-dimethylation of amine using formaldehyde and sodium cyanoborohydride.21 This modification on N-terminus and Lys of a peptide is known to have little effect on the fragmentation of the labeled peptides, yet a strong characteristic a1 ion from dimethylated N-terminal amino acid can be generated.22 Because DiART reagents are in fact the conjugates of N,N′-dimethylleucine (the reporter) and β-alanine (the balancer), DiART-labeled peptides should perform similarly as those treated under dimethylation conditions. Indeed, we were able to demonstrate the application of the DiART technique for protein quantification on a MALDI time-of-flight/time-of-flight (MALDI-TOF/TOF) mass spectrometer.19 However, because the reporter ions are in the low-mass range, DiART reagents are not compatible with every type of MS instrument. For instance, peptide fragmentation in ion-trap mass spectrometers by CID, albeit highly efficient, is subject to the so-called “one-third rule”, meaning that fragments below 25-30% of their precursor massto-charge cannot be trapped and detected.23 This is clearly demonstrated in Figure 2A, where the CID-MS/MS spectrum of a peptide (m/z ) 850.94 with double charge) displayed no peaks below m/z ) 365. Although the sequence of this peptide (FSTVAGESGSADTVR) could be easily determined by other (21) Hsu, J. L.; Huang, S. Y.; Chow, N. H.; Chen, S. H. Anal. Chem. 2003, 75, 6843–6852. (22) Hsu, J. L.; Huang, S. Y.; Shiea, J. T.; Huang, W. Y.; Chen, S. H. J. Proteome Res. 2005, 4, 101–108. (23) Yang, Y. H.; Lee, K.; Jang, K. S.; Kim, Y. G.; Park, S. H.; Lee, C. S.; Kim, B. G. Anal. Biochem. 2009, 387, 133–135.

Figure 2. CID (A) and HCD (B) MS/MS spectra of a representative precursor ion (m/z ) 850.94 with double charge). (A) The sequence of this peptide was identified with high confidence (Mascot score ) 136). An asterisk (*) indicates the DiART labeling site. Both b- and y-series fragments are marked in the CID spectrum. Peaks marked with # are parent ions after losing a reporter. CSBO ) bovine catalase. (B) The inset is the expanded HCD spectrum within the m/z range of 113-120. Relative ratios of this peptide in different samples are shown and compared to their predefined value (in parentheses).

larger peaks, relative quantitation information is lost because of the missing low-mass region. Recent advancements in mass spectrometers have resolved this low-mass cutoff limitation by introducing different fragmentation mechanisms like pulsed-Q dissociation (PQD)24 and HCD.25,26 When the same peptide was fragmented by HCD (Figure 2B), which has much better lowmass coverage than CID, the low-mass reporter ions (m/z ) 114-119) became the most abundant peaks and their S/N ratios were high enough to allow this differentially labeled peptide to be accurately quantified. 2 H-Associated Isotope Effects. Because DiART reagents use 2H as one of the heavy isotopes for labeling, the synthesis of DiART compounds is much easier than their counterparts that only use 13C and 15N. Interestingly, even though many 2Hlabeled reagents were widely used in the earlier days and are more cost-effective than 13C- or 15N-only tags, they have been largely phased out in proteomic analysis because of a few adverse properties. For example, it has been observed that 2H can exchange with 1H during gas-phase MS/MS fragmentation, even if 2H is linked to a carbon that is normally nonexchangeable.27 This 2H-1H exchange will cause partial loss of 2H labels and compromise the accuracy of protein quantification. In addition, because 2H is slightly more hydrophilic than 1H, 2Hlabeled peptides tend to elute faster than unlabeled counterparts in reversed-phase HPLC (RP-HPLC). This chromatographic resolution of differentially labeled peptides can cause a continuous change of their abundance ratio across their entire chromatographic peak and make it difficult to determine their ratio reliably.16 Therefore, 2H-containing labeling reagents, such as DiART, require careful designs to prevent these deleterious 2 H-related isotopic effects. (24) Bantscheff, M.; Boesche, M.; Eberhard, D.; Matthieson, T.; Sweetman, G.; Kuster, B. Mol. Cell. Proteomics 2008, 7, 1702–1713. (25) Kocher, T.; Pichler, P.; Schutzbier, M.; Stingl, C.; Kaul, A.; Teucher, N.; Hasenfuss, G.; Penninger, J. M.; Mechtler, K. J. Proteome Res. 2009, 8, 4743–4752. (26) Boja, E. S.; Phillips, D.; French, S. A.; Harris, R. A.; Balaban, R. S. J. Proteome Res. 2009, 8, 4665–4675. (27) Burinsky, D. J.; Sides, S. L. J. Am. Soc. Mass Spectrom. 2004, 15, 1300– 1314.

Figure 3. HCD MS/MS spectra of each DiART reagent.

To test whether 2H isotopes in DiART reagents are exchangeable with 1H during MS/MS fragmentation, we infused each of the six DiART reagents into an LTQ-Orbitrap instrument and fragmented them with HCD. As expected, each reagent produced a strong singly charged reporter ion with expected m/z while no other satellite peaks were observed (Figure 3). This result not only demonstrated that the partial loss of 2H labels during MS/MS was not a concern for DiART reagents, but also suggested that these reagents were of high isotopic purity (>97%). 2 H-associated chromatographic shifts, however, can be a more challenging problem for 2H-labeling approaches. Because 2 H is more hydrophilic than 1H, inclusion of 2H could significantly affect interaction between the analyte and stationary phase of reversed-phase HPLC columns and result in altered retention time. As a matter of fact, baseline separation between an analyte and its 2H-labeled analogues is routinely observed in small-molecule HPLC analysis. In our attempt to obtain relative quantitation of peptides using an isobaric mass tag, this kind of chromatography shift will be detrimental. Even a slight difference in chromatography retention time will result in inconsistent ratios between peptides labeled with different tags over a chromatography peak. In general, a higher number of 2H used for labeling leads to more separation between Analytical Chemistry, Vol. 82, No. 18, September 15, 2010

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labeled and unlabeled analyte, and 2H isotopes on hydrophobic positions contribute more to this resolution than those close to hydrophilic groups. As a result, a few methods have been developed to control this 2H isotope effect, e.g., using the minimum number of 2H atoms in labeling tags and placing 2H next to hydrophilic groups.17 This concept has been proven by reductive methylation, a reliable and cost-effective approach for large-scale quantitative proteomics studies.28,29 In this application, tryptic peptide mixtures are treated with formaldehyde and sodium cyanoborohydride to modify N-terminal and lysine amine to N,N′-dimethylamine. An Arg- or Lys-containing tryptic peptide will consequently contain two or four methyl groups, respectively. When a tryptic sample is labeled with 2H-formaldehyde, an Arg-containing peptide will exhibit a +4 mass shift from its counterpart reacted with unlabeled formaldehyde, whereas a Lys-containing peptide will show a +8 mass difference. Because 2H atoms in these peptides are close to the hydrophilic amine and only interact weakly with a C18 reversedphase column, differentially labeled peptides can still coelute in RP-HPLC, allowing them to be quantified by MS. The development of DiART reagents adopted the same principless keeping the number of 2H atoms in each reagent minimum (up to four per molecule) and positioning the 2H around polar amine groupssto minimize potential 2H-related chromatographic shift. We evaluated the performance of the DiART design using a three-protein sample containing 100 µg of bovine catalase (CSBO), 10 µg of horse myoglobin (MYG_EQU), and 1 µg of bovine serum albumin (BSA). After trypsin digestion, we divided the resulting peptide mixture into six tubes and let each of them react with one of the six DiART reagents, respectively. Once labeling was completed, we mixed these six samples together at a predefined ratio (114:115:116:117:118:119 ) 5:2:5:2:1:1), then carried out LC-MS/MS analysis. During data acquisition, dynamic exclusion was turned off so that a peptide would be selected for MS/MS analysis repeatedly as long as it was one of the three most intense ions in the full-scan MS. In addition, two peptides were selected from the initial analysis and added to a parent list so that they would be selected for MS/MS analysis as long as they were detected in the full-scan MS spectrum, regardless of their relative intensity. These two peptides were selected to represent an early eluting peptide and a late eluting peptide with low relative intensity. HPLC separation was carried out for 3 h (120 min for 5-35% B, when most peptides elutes) to represent the most rigorous separation that might be used in practical sample analysis. After LC-MS/MS analysis, computer-reconstructed chromatograms for each of the six reporter ions were created in six channels and normalized (Figure 4). The peaks of both peptides, whose sequences were identified as VLNEEQR and TFYLK, respectively, overlaid perfectly in all channels. It is worth noting that TFYLK is a short and hydrophobic peptide that has two labeling sites, so its interaction with the RP-HPLC stationary phase is expected to be affected chromatographically more by DiART tags than larger and more hydrophilic peptides. However, even in this extreme case, chromatographic shift was still negligible. This suggests that the performance of DiART tags met the expectation of their design in minimizing 2H isotope effects. (28) Lemeer, S.; Jopling, C.; Gouw, J.; Mohammed, S.; Heck, A. J.; Slijper, M.; den Hertog, J. Mol. Cell. Proteomics 2008, 7, 2176–2187. (29) Melanson, J. E.; Avery, S. L.; Pinto, D. M. Proteomics 2006, 6, 4466–4474.

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Figure 4. HPLC chromatograms of a peptide mixture differentially labeled with DiART reagents. Computer-reconstructed chromatograms for each of the reporter ions were created in six channels (top). Two parent ions at different retention times (m/z ) 552.81 with double charge at 41.0-43.0 min, m/z ) 553.36 with double charge at 73.2-74.5 min whose parent ion was only 15% base peak in the full-scan spectrum) were chosen as representative peptides for determining the 2H-related chromatographic shift. The normalized chromatograms of six reporter ions from each parent ion were overlaid (middle). The sequences of these peptides are listed with asterisks indicating DiART labeling sites. Five intensity ratios among these reporter ions (115/114, 116/114, 117/114, 118/114, 119/114) are shown at each time point during this period (bottom). The means and CVs (in parentheses) of these ratios were calculated (from 50 and 32 time points for parent ions 552.81 and 553.37, respectively) and color coded (115/114, dark red; 116/114, green; 117/114, blue; 118/ 114, dark yellow; 119/114, dark pink).

During this MS acquisition, we deliberately turned off dynamic exclusion so that we could produce fragments from a few abundant ions at multiple time points and obtain their MS/MS spectra during their entire period when they passed through an HPLC column. This information is critical to demonstrate whether DiART-labeled peptides exhibit 2H-related chromatographic shifts. In a practical application using a complex biological sample, however, the dynamic exclusion must be turned on to prevent the same parent ion from being acquired again for a specified period of time so that the detection of low-abundance peptides can be achieved. Therefore, most peptides would only have one chance to be detected and fragmented after they are eluted off the HPLC column. Although we can set up the MS instrument to acquire MS/MS when a precursor ion is at its maximum intensity, practically it can be difficult to control due to the intrinsic instability of the jet stream during electrospray. It is thereby of great importance that the ratios among differently labeled peptides remain constant throughout their chromatographic peaks so that they could be accurately quantified at any given point of time during elution. To test the performance of DiART-labeled samples, we calculated the ratios of two peptides (VLNEEQR and TFYLK) at each time point covering their entire chromatographic peaks (Figure 4). We then calculated the mean and the coefficient of variation (CV)

Table 1. Mascot Report for DiART-Based Protein Quantification protein name

114/119 0.05

CSBO MYG_EQU ABBOS

0.059b (25c/1.088d) 0.054 (6/1.141) 0.063 (8/1.090)

a

115/119 0.01

116/119 0.02

117/119 0.20

118/119 0.50

protein score

0.000 0.000 0.000

0.035 (22/1.038) 0.038 (4/1.063) 0.035 (10/1.101)

0.192 (41/1.037) 0.193 (9/1.097) 0.178 (13/1.050)

0.546 (49/1.040) 0.551 (10/1.048) 0.525 (15/1.034)

1165 249 396

sequence coverage (%) 58 60 39

a Predefined protein ratio. b Measured protein ratio. c Number of peptides used for calculating the measured protein ratio. d GSD of the measured protein ratio. All values are reported by Mascot.

of these values. Clearly, even though the second peptide (TFYLK) had slightly higher CVs than those of the first one, for the reasons that are discussed above, their CVs were within an acceptable level, indicating that relative abundance of a peptide from up to six different samples could be quantified reliably at any time point during its elution. In addition, we should point out that tryptic peptides with one missed cleavage site are usually included in Mascot searching. These peptides, as well as those containing the Lys-Pro (KP) sequence, may have three or more primary amine groups, so the 114/115-labeled and 118/119-labeled peptides would differ in the number of 2H/1H by 12 or more. Interestingly, even these peptides could be quantified rather accurately by DiART. For instance, the ratios (118/114 and 119/114) of a KP-bearing peptide from CSBO, YNEEKPK, were 0.216 and 0.146, respectively (see the Supporting Information), which were still close to their predefined values of 0.200. Finally, to further improve quantification quality, we could optimize conditions for trypsin digestion, such as adding more trypsin, incubating a longer time, and using microwave reactors, to minimize the number of missed cleavages. Alternatively, we can edit the quantitation method file so that peptides with more than one missed cleavage will not be used for quantitation. Hybrid HCD-CID for Protein Identification and Quantification. Currently, all of the peptide identification methods are based on statistical algorithms, which compare experimentally obtained peptide fragments with those calculated from all proteins in a database to find the best match.30,31 To obtain a large number of peptide identifications, high quality of MS/MS peptide fragmentation is critical for obtaining accurate mass and reliable peptide assignments. Despite its capability to cover the low-mass range, HCD usually results in less optimum MS/MS spectra for peptide identification than the more traditional CID method. By simply relying on HCD data, one may misidentify or ignore many low-abundant peptides, leading to a high level of false positives and negatives. Fortunately, on an LTQ-Orbitrap instrument, both CID and HCD of the same precursor ion can be acquired in parallel, with CID in the linear ion trap and HCD in the Orbitrap, and then combined together for further analysis.25 Because CID and HCD are carried out and detected in two separate mass analyzers, this acquisition of CID spectra does not involve increased cycle time. We adopted this scheme and performed an LC-MS/MS analysis on a peptide sample that was also prepared from the three-protein mixture, but mixed at a different predefined ratio (114:115:116:117:118:119 (30) Perkins, D. N.; Pappin, D. J.; Creasy, D. M.; Cottrell, J. S. Electrophoresis 1999, 20, 3551–3567. (31) Eng, J. K.; McCormack, A. L.; Yates, J. R. J. Am. Soc. Mass Spectrom. 1994, 5, 976–989.

) 5:1:2:20:50:100). This ratio would allow us to estimate the linear range of DiART reagents since peptides labeled with 119 was 100fold over those labeled with 115. To increase the sensitivity of peptide detection during acquisition, dynamic exclusion was turned on to exclude ions from being selected again for MS/MS analysis for 30 s. This condition is necessary to prevent the MS instrument from acquiring a few high-abundant precursor ions repetitively and to increase the chance of detecting low-abundant peptides. For instance, when dynamic exclusion was turned off, only peptides from CSBO and MYG_EQU were identified, whereas none from BSA, whose concentration was 100-fold lower than that of CSBO, were detected. In contrast, when the dynamic exclusion was turned on, we were able to identify all three proteins with high Mascot scores (Table 1). CSBO, the most abundant protein in this three-protein mixture, had an overall score of 1165 with 58% sequence coverage, and the least abundant BSA also exhibited an overall score of 396 with 39% sequence coverage, suggesting DiART labeling could detect proteins with a broad dynamic range of concentration. For any stable isotope labeling based protein quantification, it is critical to determine not only the ratio of a protein from different samples, which is defined in Mascot as the geometric mean of all identified peptides from this protein, but also the geometric standard deviation (GSD), which reflects the variation and reliability of this ratio. DiART reagents could indeed provide an accurate quantification of proteins. As shown in Table 1, for example, the concentration of CSBO in the sample labeled with 118 over that labeled with 119 was determined at 0.546, which is very close to predefined 0.500. This value is the geometric mean of a group of ratios obtained from 49 peptides with a GSD of 1.040, indicating that the confidence interval of this value is from 0.525 (0.546/1.040) to 0.567 (0.546 × 1.040). We must emphasize that these 49 peptides from CSBO included some repetitive peptides because they were detected and fragmented independently in MS/ MS. However, only high-quality peptides were used for quantification and those peptides with poor scores were excluded (see the Supporting Information for ratios of individual peptides). From the same experiment, we also found that DiART reagents had a linear range between 1 and 2 orders of magnitude. For instance, all three proteins had ratios very close to predefined 0.05 when 114/119 values were calculated. The 116/119 values were slightly off, but were still quite consistent, so we could not determine whether this deviation was introduced when we mixed the peptide samples or was caused by limited linear range. We did not observe any reporter ion at 115, suggesting 100-fold was the upper limit that DiART reagents could measure. Other labeling reagents like iTRAQ also have similar problemsthe least abundant reporter ions are usually not observed if the sample concentration Analytical Chemistry, Vol. 82, No. 18, September 15, 2010

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differs in more than 100-fold. This is partly because the number of parent ions used for MS/MS fragmentation is limited and the level of the respective reporter ions is too low to be detectable. However, this will not be a critical issue because the same proteins varying more than 10-fold in abundance among various biological samples can be simply considered as expressed or nonexpressed proteins. To make DiART useful for large-scale proteomics applications, it is necessary to characterize DiART reagents with real biological samples. Since this technique is conceptually similar to iTRAQ/ TMT, lessons learned from iTRAQ/TMT should also be applicable on DiART. Recently, a detailed study on the good and bad of iTRAQ has been carried out, providing valuable information that can predict how DiART will perform on a complex sample.32 For instance, when a complex sample labeled by iTRAQ is analyzed by LC-MS/MS, it is not unusual to have two coeluting peptides with close m/z to be isolated together as parent ions, resulting in an MS/MS spectrum with fragments from both peptides. As a result, reporter ion ratios will reflect rations of the sum of the two peptides. This mixed MS/MS background will suppress potentially large intensity differences, with more underestimation on low-abundance proteins than high-abundance proteins. DiART reagents are unlikely immune from this effect. However, this background interference can be minimized by improving the HPLC separation and, less ideally, by reducing the MS selection window. Interestingly, in the same study, it was found that correction factors for isotope impurity of iTRAQ reagents also play a critical role in accuracy and precision of quantification and must be employed when analyzing complex samples. A benefit offered by DiART, therefore, is that DiART reagents will not require this correction, thanks to their high isotope purity. Hybrid ETD-HCD for Phosphorylated Peptide Detection and Quantification. Besides HCD and CID, ETD, a different fragmentation method, has recently become very popular in determination of posttranslational modification (PTM) sites due to its ability in keeping labile PTM groups (e.g., phosphorylation and glycosylation) intact during MS/MS fragmentation.33,34 ETD also has broad applications in top-down proteomics because it can fragment large peptides evenly at almost every peptide bond without sequence bias, providing a nearly complete set of ions for protein assignment.35 However, unlike CID and HCD that generate b- and y-series fragments, ETD fragmentation produces c- and z-series ions. As a result, DiART-labeled peptides would not display the same signature ions at 114-119 when fragmented by ETD as their fragmentation patterns are completely different. To take advantage of the benefits offered by ETD for PTM detection and by HCD for accurate peptide quantification, we developed a hybrid ETD-HCD scheme, in which the ETD and HCD spectra of the same precursor ion were acquired in parallel. These data can then be processed in two different ways for the identification and quantification of PTM-containing peptides (Figure 5). One way is to simply combine spectra of the same precursor as we did with a standard analysis; then a Mascot server (32) Ow, S. Y.; Salim, M.; Noirel, J.; Evans, C.; Rehman, I.; Wright, P. C. J. Proteome Res. 2009, 8, 5347–5355. (33) Wiesner, J.; Premsler, T.; Sickmann, A. Proteomics 2008, 8, 4466–4483. (34) Mikesh, L. M.; Ueberheide, B.; Chi, A.; Coon, J. J.; Syka, J. E.; Shabanowitz, J.; Hunt, D. F. Biochim. Biophys. Acta 2006, 1764, 1811–1822. (35) Bunger, M. K.; Cargile, B. J.; Ngunjiri, A.; Bundy, J. L.; Stephenson, J. L., Jr. Anal. Chem. 2008, 80, 1459–1467.

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Figure 5. Flowchart of two methods for hybrid ETD-HCD data processing. In method A, data extraction and data merge were performed with Proteome Discoverer. In method B, data extraction was performed with Proteome Discoverer, whereas data processing and data merge were completed by an in-house Perl script.

is configured so that it is compatible with database search by adding a hypothetical type of MS instrument that can generate all four-series ions (b-, y-, c-, z-ions) in a single MS/MS scan. To test this method, we digested a pSer-containing protein, bovine β-casein, and labeled them with each DiART reagent, respectively. We then mixed the differentially labeled peptides at a predefined ratio (1:0.5:2:1:0.25:7), enriched phosphorylated peptides using a PHOS-Select iron affinity gel,36 and analyzed them using the LC-ETD-HCD scheme. We identified a pSer-containing peptide (FQpSEEQQQTEDELQDK) with a score of 56 (Expect value ) 5.6 × 10-3). Nevertheless, this direct approach appears to have certain limitations in a large-scale phosphoproteome project. Because ETD is a less efficient fragmentation technique than CID or HCD, the precursor ions and their charge-reduced forms are usually the only abundant peaks in MS/MS spectra with c- and z-fragment ions below 25%. Consequently, the database search algorithms will sometimes fail to report correct phosphopeptides, yielding high rate of false identification. In addition, because the neutral loss of phosphate group from pSer or pThr residues is frequently observed in CID/HCD, mixing HCD- and ETD-generated fragments together will make it more difficult for a search algorithm to locate phosphorylation sites, especially in peptides containing multiple modifications. To address these issues, we developed a more robust approach for data analysis (Figure 5B). First, the ETD MS/MS spectrum of a peptide was processed to remove any precursor-related peaks and ions below m/z 250 as these small fragments are usually generated from DiART tags. Second, the reporter ions in the HCD spectrum of the same peptide were extracted and their intensities were normalized to ion counts comparable to ETD fragment ions. This normalization is necessary as the ion intensities of HCD fragments is usually much higher than those of ETD, which can lower the score of identified peptides. Finally, the identity and quantification of this peptide were determined using the combined peaks from ETD and HCD when searching a Mascot server configured with a regular ETD-TRAP instrument. This approach used ETD data for peptide identification that preserves labile PTM groups and HCD data for peptide quantification that offers strong (36) Zhou, W.; Merrick, B. A.; Khaledi, M. G.; Tomer, K. B. J. Am. Soc. Mass Spectrom. 2000, 11, 273–282.

Figure 6. ETD (A) and HCD (B) MS/MS spectra of a phosphorylated peptide (m/z ) 832.8 with triple charge). (A) The sequence of this peptide was identified with high confidence (Mascot score ) 95), and the asterisk (*) indicates DiART labeling sites. Both c- and z-series fragments are marked in the ETD spectrum. Peak m/z ) 1248.6, with double charge, was a charge-reduced parent ion, and peak m/z ) 1226.7, with double charge, was unidentified. Cas ) bovine β-casein. (B) The expanded HCD spectrum within the m/z range of 113-120. Relative ratios of this peptide were calculated and compared to their predefined value (in parentheses).

signature reporter ions with high S/N ratios. By using this new protocol and the same MS data set, we were able to identify the pSer-bearing peptide (FQpSEEQQQTEDELQDK) with a score of 95 (Expect value ) 9.8 × 10-7) (Figure 6) and quantify the ratio at 1.00:0.51:2.12:1.23:0.29:7.40, representing a significant improvement on the reliability of peptide assignment. This example demonstrated that data analysis was equally important as data acquisition in MS-based proteomics research. It is also worth noting that we observed a few other pSer-containing peptides (EQLSTpSEENSK and TVDMEpSTEVFTK) that were apparently R-casein contaminants in the tested β-casein sample. CONCLUSION Stable isotope labeling approaches, especially iTRAQ, TMT, and other tags that allow for multiplex measurements, remain a mainstay of quantitative proteomics and have considerable advantages over label-free methods in terms of throughput, accuracy, and precision. Certainly, high cost associated with these reagents is a limiting factor that prevents them from clinical proteomics applications, in which a large number of samples needs to be processed. The DiART technique, thanks to its excellent performance, streamlined protocols, and, more importantly, better affordability, would provide a robust option for those who are sensitive to the cost-effectiveness of labeling strategies. Furthermore, it would now be feasible to design more sophisticated

DiART derivatives that can fit various needs for broad applications. Finally, we should note that DiART and other stable isotope labeling approaches complement to label-free platforms as labeling methods can provide more accurate quantification on small changes, whereas the latter are more suitable for identifying proteins with large difference under various conditions. ACKNOWLEDGMENT We thank Dr. Feng Tao for his valuable discussion and comments. This project was supported in part by the NSF IIP No. 0945037 Grant. NOTE ADDED AFTER ASAP PUBLICATION This paper was published on the Web on August 17, 2010 with a typographical error in the data analysis section of the Experimental Section and a typographical error in the discussion of the 2H-associated isotope effects. The corrected version was reposted on August 20, 2010. SUPPORTING INFORMATION AVAILABLE Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org. Received for review May 19, 2010. Accepted August 6, 2010. AC101306X

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