A Simple Solid Phase Mass Tagging Approach for Quantitative

Oct 5, 2003 - Department of Chemical Engineering, Yale University, New Haven, ... To whom all correspondence should be addressed: James A. Wilkins,...
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A Simple Solid Phase Mass Tagging Approach for Quantitative Proteomics Yang Shi,† Rong Xiang,† Janet K. Crawford,‡ Christopher M. Colangelo,‡ Csaba Horva´ th,† and James A. Wilkins*,† Department of Chemical Engineering, Yale University, New Haven, Connecticut 06520-8286 and W. M. Keck Foundation Biotechnology Resource Laboratory, Yale University, New Haven, Connecticut 06520-8114 Received October 5, 2003

New mass-tagging reagents for quantitative proteomics measurements have been designed using solid phase peptide synthesis technology. The solid phase mass tags have been used to accurately measure the relative amounts of cysteine-containing peptides in model peptide mixtures as well as in mixtures of tryptic digests in the femtomol range. Measurements were made using both matrix-assisted laser desorption ionization-time-of-flight mass spectrometry (MALDI-TOF MS) and online reversed-phase capillary liquid chromatography coupled through a nanoelectrospray interface to an ion trap mass spectrometer (capillary LC/ESI-MS). Results of mass-tagging experiments obtained from these two mass spectrometry techniques and their relative advantages and disadvantages for identification and quantitation of mass tagged peptides are compared. These reagents provide a simple, rapid and costeffective alternative to currently available mass tagging technologies. Keywords: quantitative proteomics • capillary liquid chromatography • mass spectrometry • solid phase • stable isotope labeling

Introduction An important goal in proteomics is to compare the relative amounts of different proteins in biological samples and to try to correlate these differences with changes in physiological state. Such changes may, for example, reflect transition to a diseased from a normal state and give clues about potential targets for therapeutic intervention.1,2 Relative expression levels of cellular proteins under different conditions, e.g., normal and transformed cells; cells subjected to different growth conditions, etc., have traditionally been measured using differential labeling techniques in which proteins are separated by two-dimensional gel electrophoresis (2-D gels) and stained followed by quantitative or qualitative comparison of the levels of corresponding spots on the gels.3,4 Spots of interest are removed from the gel and proteins are identified by mass spectrometry. More recently, a technique has been introduced in which proteins from two cell populations are differentially labeled with Cy3 and Cy5 fluorescent dyes, mixed and separated on the same 2-D gel followed by quantitative measurement and comparison of fluorescence levels. Although the latter technique, termed difference gel electrophoresis (DIGE),5 vastly improves the reproducibility of comparative studies using 2-D gels, a number of problems still remain, such as the difficulty in detecting low * To whom all correspondence should be addressed: James A. Wilkins, Department of Chemical Engineering, Yale University, P.O. Box 208286, New Haven, CT 06520-8286. Tel: (203) 432-4373. Fax: (203) 432-4360. E-mail: [email protected]. † Department of Chemical Engineering, Yale University. ‡ W. M. Keck Foundation Biotechnology Resource Laboratory, Yale University.

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abundance proteins. The isotope coded affinity tag (ICAT) technique pioneered in Aebersold’s laboratory6,7 takes advantage of differential tagging of cysteine residues in proteins with stable isotopes followed by affinity and ion exchange chromatography. Peptides in individual ion exchange fractions are then identified using online reversed phase liquid chromatography coupled to mass spectrometry. Relative amounts of peptides are determined by comparison of ion currents obtained from peptides labeled with deuterium (heavy) or hydrogen (light) containing mass tags in a mass spectrometer. The original ICAT reagent was designed using d0 (light) or d8 (heavy) tagging reagents to provide an 8 mass unit difference between light and heavy tagged peptides. ICAT therefore allows relative measurement of cysteine-containing peptides from two related samples and offers the potential of reproducible, quantitative comparison and relatively rapid identification of a number of cellular proteins when coupled with liquid chromatography and tandem mass spectrometry methods.7 However, some welldocumented problems were found with the original ICAT reagent including its relatively high mass, fragmentation of the tag itself and chromatographic separation of d0 and d8 labeled peptides.8 These problems led to difficulties with quantitation and to peptide losses during chromatographic purifications done prior to the final reversed phase peptide separation. More recently, a new ICAT reagent was introduced that addresses some of the above problems. Specifically, introduction of 12C or 13C isotopes produces the mass difference in the light and heavy tags (Applied Biosystems, Inc, Foster City, CA; www.appliedbiosystems.com). This eliminates the problem of chromatographic separation of the light and heavy labeled 10.1021/pr034081k CCC: $27.50

 2004 American Chemical Society

Simple Solid Phase Mass Tagging Approach

versions of the same peptide and simplifies quantitation. A cleavable linker was also introduced so that the biotin moiety could be removed following avidin affinity chromatography. The commercially available reagents however are expensive and still retain the limitation that ion exchange and affinity chromatography must be performed prior to reversed phase LC/ MS. The initial chromatographic steps in this process are timeconsuming and can lead to sample losses, reducing of sensitivity, however, Gygi et al.9 have shown that low abundance proteins can be identified using a three-dimensional chromatography approach. More recently, solid-phase mass tagging approaches have appeared in the literature.10,11 Solid-phase techniques offer the advantage that upfront affinity and ion exchange chromatographic steps and their associated problems are eliminated and could allow operation in a more high throughput format. Zhou et al.10 introduced the idea of a solid-phase mass tag with an attached amino acid (leucine) and showed that their solidphase reagents were able to detect more labeled peptide pairs in a complex mixture derived from yeast than the traditional ICAT reagent. Qiu et al.11 designed reagents termed ALICE (acid-labile isotope-coded extractants). They were able to accurately measure ratios of proteins in model mixtures. Although it takes advantage of the solid phase approach, the ALICE reagent along with the solid phase tags introduced by Zhou et al. still use deuterium labeling for the heavy tags, therefore the problem of chromatographic separation of light and heavy tagged peptides remains. In addition, the reagent and sample preparation steps for both of these reagents employ synthetic and/or cleavage techniques not commonly available to most laboratories. In both of the above papers, a large excess of solid-phase reactive sites was added compared to the sample size, but this was not highlighted by the authors.10,11 Excess labeling reagent increases the potential for unwanted side reactions with noncysteine containing peptides. This may be particularly troublesome in complex mixtures containing low abundance proteins or in small samples. Our approach to the mass-tagging problem was to develop a reagent that could be easily synthesized, using commonly available techniques and reagents at a reasonable cost. The area of solid phase peptide synthesis provides a rich source of options for creation of such a reagent. In this study, we describe the synthesis of mass tags based on uniformly 12C and 13C labeled tri-alanine peptides iodoacetylated on their N-termini to provide a cysteine-reactive function. The C-terminal ends of the peptides are linked in conventional fashion through an acid-labile handle to a polymethacrylate solid phase support. These mass tags are shown to react specifically with cysteine containing peptides in defined mixtures. After cleavage, these reagents provide a 9 mass unit difference between heavy and light labeled cysteine containing peptides that can be used to accurately measure the ratios of proteins in mixtures using MALDI-TOF MS or LC/MS. The mass tag itself is also shown to be stable during collision induced dissociation (CID) of peptides. These reagents should therefore be useful in proteomics experiments in which quantitative measurements of relative amounts of proteins in samples such as normal and diseased tissues are desired. Several advantages of the current technology are discussed.

Experimental Section Materials. All proteins and a laminin fragment peptide (laminin fragment 925-933; L peptide) used in tagging experi-

research articles ments along with HPLC grade water, acetonitrile, (2,5) dihydroxybenzoic acid (DHB) and trypsin (TPCK treated) were obtained from Sigma-Aldrich (St. Louis, MO). The small-scale peptide synthesis group at the Keck Biotechnology Resource Facility at Yale produced other cysteine-containing synthetic peptides (ID numbers 153, 9772, 856, and 876) used in this work. They were supplied at a concentration of 1 mg/mL in 0.02% TFA. BioMac polymethacrylate resin (1000 Å pore size) with attached Fmoc-Rink linker was obtained from Biosearch Technologies, Inc (Novato, CA). Trifluoroacetic acid (TFA) was from American Bioanalytical (Natick, MA). Succinimidyl iodoacetate and Tris (2-carboxyethyl) phosphine hydrochloride (TCEP) were from Pierce Biotechnology (Rockford, IL). 15 µm fused silica spray tips (75 µm i.d.; 360 µm o.d.) for electrospray mass spectrometry were from New Objective (Woburn, MA). Discovery C18 resin (5 µm diameter, 300 Å pore size) used for column fabrication was a gift from Supelco (Sigma Aldrich, St. Louis, MO). Peptide Synthesis and Preparation of Mass Tags. Unless otherwise noted, all chemicals were purchased from American Bioanalytical. The peptides were synthesized by Fmoc chemistry on a Rainin Symphony peptide synthesizer (Rainin Instruments, Oakland, CA). The resin used was Fmoc-Linker-AMBioMac from Biosearch Technologies Inc (Novato, CA). 25 µmol of resin (based on a value of 63 µmol sites/g resin) was used for each synthesis. The 12C peptide was synthesized using standard protocols. Deprotection was done with 20% piperidine/dimethylformamide (DMF), 2 × 5 min each. Amino acid coupling was done at a 4-fold excess of amino acid in 0.4 M N-methylmorpholine/DMF with 2-(1H-benzotriazole-1-yl)1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU), twice for 20 min each. Fmoc-L-Alanine-OH (12C) was obtained from Novabiochem (Laufelfingen, Switzerland). Uniformly labeled Fmoc-L-Alanine-OH (13C) was obtained from Isotec (Miamisburg, OH) and single 1 h coupling cycles were used. After peptide synthesis, the N-terminal Fmoc was removed and the resin was washed with DMF, then dichloromethane (DCM) and dried under nitrogen prior to reaction with succimidyliodoacetate (SIA). Reaction with SIA was carried out at room temperature by addition of 0.5 g of either 12C or 13C trialanine beads to 1 mL of DMF containing a 5-fold excess of SIA over free amino groups (63 µmol/g resin). The reaction was found to be complete after 2 h and beads were washed with 5 volumes of DMF followed by resuspension in 5 mL of 0.02% TFA in H2O. Mass tags were stored in this solution at 4 °C and were stable for at least 1 month. Preparation of Tryptic Digests. Tryptic digests of proteins were made as follows. Solutions of bovine serum albumin (BSA), ovalbumin (OVA) and ribonuclease A (RNase) were prepared at concentrations of 1 mg/mL each in 10 mM Trisacetate, pH 8.3, 0.1% (w/v) sodium dodecyl sulfate (SDS) and 5 mM TCEP. The solutions were heated to 100 °C for 5 min to denature the proteins. Solutions were cooled to room temperature and trypsin was added to each at a ratio of 1:50 (w/w) with respect to protein. Solutions were incubated overnight at room temperature to hydrolyze proteins followed by addition of PMSF to a final concentration of 1 mM to inhibit remaining trypsin activity. TFA was then added to a final concentration of 0.02% to reduce the pH of the solutions to less than 3. Tryptic digests were stored at 4 °C prior to use. Mass Tagging Protocol. For most experiments, 100 µg of beads with either 12C or 13C mass tag was added to peptide solutions, and samples were incubated for 1 h. Peptide soluJournal of Proteome Research • Vol. 3, No. 1, 2004 105

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Figure 1. Schematic diagram of the solid-phase mass tag. Solidphase reagents were synthesized using polymethacrylate/PEG resin with an acid-labile “Rink” linker. The mass tag itself is a trialanine peptide containing either 12C or 13C at the * positions to produce a 9 mass unit difference between the reagents. The reactive iodoacetate group is added to the N-terminus of the trialanine peptides using succinimidyliodoacetate (SIA).

tions were prepared by adding the desired amount of each peptide to a 20-µL aliquot of 0.1 M Tris-acetate, pH 8.5 in a 0.5 mL microfuge tube. Samples were mixed using a vortex (Reax Top; Heidolph Instruments, Kelheim, FRG) set on continuous mixing mode in order to keep the resin in suspension during the incubation. After incubation, samples were centrifuged for 1 min in a microfuge (Eppendorf, Hamburg, FRG), and the supernatant fluid was removed. Beads were mixed together and then washed with 20-µL volumes for 20 min each with 70% ACN, 0.1% (w/v) SDS and again with 70% ACN with mixing as above. After washing, beads were cleaved by incubating them with 20 µL of 50% (v/v) TFA in DCM for 1 h at room temperature. Beads were centrifuged as above and supernatants were removed and vacuum-dried in a Speed Vac (Savant Instruments, Holbrook, NY). Mass tagged peptides were resuspended in 20 µL of 70% ACN containing 0.02% TFA. In the MALDI-TOF MS experiments described in Figures 2-4 and 6, 0.5 µL (1/40th of the total mass tagged sample) was applied to the sample target. In the LC/MS experiments described in Figures 5 and 7, the mass tagged sample in 70% ACN was diluted 1/100 with 0.02% TFA in water prior to loading. The amount loaded in this case represented 1/80th of the original sample. MALDI-TOF Mass Spectrometry. All spectra were recorded using a Voyager DE RP instrument (Applied Biosystems, Foster City, CA). For sample preparation, 1.0 µL of matrix (10 mg/mL DHB in 10% ACN) was spotted onto targets and mixed with 0.5 µL of sample. Samples were air-dried and spectra were obtained in either linear or reflectron mode at an accelerating voltage of 20 kV. Online Liquid Chromatography/Nanoelectrospray Ionization Mass Spectrometry (LC/NanoESI-MS). Reversed-phase LC directly coupled to NanoESI-MS was performed in 75 µm capillaries using linear gradient elution essentially as described12 using an LCQ ion trap mass spectrometer (Thermofinnigan, San Jose, CA) operated in the data dependent mode. Data on the first and second strongest ions above an intensity of 2 × 105 were collected with dynamic exclusion enabled with the collision energy set at 35%.

Results and Discussion To design an efficient, cost-effective mass-tagging reagent, we turned to the methods used for solid-phase peptide synthesis. Figure 1 shows a schematic diagram of the mass tags. For this purpose, we chose a polymethacrylate resin (1000 Å pore size) supplied with an acid-cleavable “rink” linker. Fmoc 106

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chemistry was used to synthesize trialanine peptides with either Fmoc 12C alanine for the “light” reagent or Fmoc 13C alanine for the “heavy” reagent (see the Experimental Section). This provides a 9 mass unit difference between light and heavy tagged peptides or a 4.5 mass unit difference between doubly charged, mass tagged tryptic peptide pairs. The technology also allows facile addition, for instance, of another alanine residue to provide a 12, 6 or 4 mass unit differences for singly, doubly or triply charged tryptic peptide pairs. The current design thus provides a flexible platform for customization of mass tagging reagents depending upon specific requirements. After synthesis of the trialanine peptides, beads were dried under nitrogen and stored in a desiccator. The cysteine-reactive iodoacetyl group was then added onto the N-terminus of the peptides by reaction with succinimidyl iodoacetate. The manufacturer reported a value of 63 µmol of sites/g resin that was in good agreement with the value obtained by measurement of the dibenzofulvene-piperidine adduct released from the rink linker prior to peptide synthesis using an extinction coefficient of 7800 M-1 cm-1 at 301 nm.13 We first examined the reaction of a single cysteine containing peptide with the mass tags. The reaction product formed by incubation of 1 pmol of a laminin-derived peptide (L peptide) with the 12C mass tag was acid cleaved and prepared as described in the Experimental Section. An aliquot of this mass tagged peptide corresponding to 25 femtomol of the original L peptide sample was analyzed by MALDI-TOF MS as shown in Figure 2A. The 12C mass tag adds 270 Da to the parent mass of the laminin peptide (967.6 Da) to give a signal at m/z 1237.6 in the MALDI-TOF mass spectrum (panel A). In Figure 2B, we examined the reaction of increasing amounts (up to 4 nmol) of L peptide with 100 µg of the 12 C mass tag. As described above, aliquots of each sample corresponding to 1/40th of the original samples were analyzed (see the Experimental Section). Thus, sensitivity in this experiment extended to the femtomol level of L peptide bound to the mass tag. With this amount of beads, saturation was reached at a level of approximately 2 nmol of L peptide. On the basis of the value from measurement of the Fmoc linker reported above (63 µmol/g beads), this indicates that approximately 1/3 of the theoretical sites are available during 1 h incubation. The time course of the L peptide reaction with the mass tag is shown in Figure 2C. In this experiment, 100 µg of the 12C mass tag was added to an excess of L peptide and aliquots were removed at the desired intervals. After cleavage, mass tagged L peptide peak intensities at m/z 1237.6 were measured using MALDI-TOF MS. The initial reaction attains a maximum level between 30 min and 1 h and remains stable for at least 24 h. Reaction kinetics is expected to be fast using the 1000 Å pore size methacrylate resin, which should exhibit efficient mass transfer characteristics for most peptides. We selected an incubation time of 1 h for all subsequent experiments. In Figure 3, quantitation was demonstrated by measuring the 12C/13C tagged peptide ratios at a range of L peptide concentrations. In this experiment, we incubated several 0.1 mg aliquots of 12C beads, each with a fixed amount of L peptide (10 pmol), while 0.1 mg aliquots of 13C beads were incubated with various amounts of the same peptide, ranging from 5 pmol to 100 pmol. Samples at each ratio were prepared in triplicate. The beads were incubated with peptide and treated as outlined in the Experimental Section. The assay was shown to respondto-different ratios of 12C/13C peptides in a linear way. The results in Figure 3 thus show that mass tags are able to accurately

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Figure 3. Detection of L peptide ratios by mass tags. Several 0.1 mg aliquots of the 12C mass tag were incubated with a fixed amount (10 pmol) of L peptide; an equal number of aliquots of 13C mass tag were incubated with different amounts of L peptide (5-100 pmol). After incubation, samples at different ratios (in triplicate) were processed as described in the Experimental Section and aliquots of cleaved products were subjected to MALDI-TOF MS analysis. L peptide concentration ratios over a 20-fold range gave a linear response. Table 1. Mass-Tagged Peptides Identified and Measured by MALDI MS and LC/MS

method

MALDI MS

LC/MS

a

Figure 2. MALDI-TOF MS analysis of the reaction product of the 12C mass tag with L peptide. (A) The spectrum of the cleaved product obtained after a 1h reaction, showing the m/z value expected by the addition of the mass tag (270 Da) to the parent mass of L peptide (967.6 Da). (B) Reaction of increasing amounts of L peptide with 0.1 mg of the 12C mass tag. Samples were incubated for 1 h and an aliquot of the cleaved product was analyzed by MALDI-TOF MS. Saturation was reached at approximately 2 nmol of L peptide. (C) Time course of the reaction. An excess of L peptide was incubated for various times with 0.1 mg of the 12C mass tag and the cleaved product was analyzed as above. The inset shows the early time course of the reaction, which is complete within 1 h.

detect relative concentration differences of L peptide in two solutions over a 20-fold concentration range. The linear response over this range suggests that MT reagents should be useful for measuring such ratios in more complex mixtures. To test the ability of the mass tags to recognize cysteines in sample mixtures, we made two peptide mixtures, each with a different amount of four cysteine-containing synthetic peptides (see Table 1). Mixture 1 contained 5 pmol of peptide 153, 5 pmol of peptide 9772, 25 pmol of peptide 956 and 12.5 pmol

peptide ID

peptide mass

exptl ratio mass mass expected 12 13 12 w/ Ctag w/ C tag ( C/13C) ratio

153 9772 956 876 153 9772 956 876

937.20 1403.70 1464.00 1566.50 937.20 1403.70 1464.00 1566.50

1207.93 1673.08 1733.91 1836.19 1207.50 837.20a 1733.80 918.50a

1216.96 1682.12 1742.90 1845.19 1216.50 841.70a 1742.70 923.40a

0.10 0.19 4.46 2.01 0.09 0.11 7.74 2.78

0.10 0.10 5.0 2.5 0.10 0.10 5.0 2.5

Doubly charged peptide ions detected.

of peptide 876. Mixture 2 contained 50 pmol, 25 pmol, 5 pmol and 5 pmol of the same peptides, respectively. We then added 0.1 mg of 12C mass tag to Mixture 1 and the same amount of 13C mass tag to Mixture 2 and incubated for 1 h at room temperature with mixing. The samples were centrifuged and supernatants were removed. Beads were then mixed together, washed, and peptides were cleaved as described in the Experimental Section. Aliquots of samples were first analyzed by MALDI-TOF MS. As shown in Figure 4, pairs of mass tagged peptides were detected, differing by 9 mass units, as expected. The inserts in Figure 4 highlight the pairs of tagged peptides showing the observed ratios. These ratios were in good agreement with the ratios expected for the peptide mixtures as shown in Table 1. An aliquot of the same sample (corresponding to 1/80th of the total mass tagged sample) was analyzed by online reversed phase capillary LC/NanoESI-MS. Figure 5A,B shows ion chromatograms for two of the four 12C and 13C labeled peptides. MS spectra collected in Figure 5C,D show spectra from individual scans obtained from the peaks of the other two pairs of mass tagged peptides. The masses of the 12C and 13C labeled pairs are shown in bold for emphasis and ratio data were in good agreement with the expected ratios (Figure 5C,D). Quantitative data obtained from this experiment are also shown in Table 1 and are comparable to those from MALDI-TOF MS. Absolute ratios observed for individual peptides sometimes vary widely from the expected ratios (e.g., see values from MALDI-TOF MS for peptide 972 in Table 1); this Journal of Proteome Research • Vol. 3, No. 1, 2004 107

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Figure 4. Mass spectrum of the mixture of mass tagged cysteine-containing peptides. Two peptide mixtures, each containing different amounts of 4 cysteine-containing synthetic peptides were prepared and incubated with 12C or 13C solid-phase mass tag. Ratios of 12C and 13C labeled peptides were measured by MALDI-TOF MS as shown in the spectrum. Insets show the individual pairs of mass tagged peptides.

Figure 5. Analysis of the mass tagged synthetic peptide mixture by online reversed phase capillary LC/NanoESI-MS. An aliquot of the sample prepared as described in Figure 4 and in the text was analyzed by capillary LC coupled to NanoESI-MS. Panels A1 and A2 with B1 and B2 show individual ion chromatograms obtained for mass tagged pairs of peptides (peptides 153 and 956, respectively). A1 and B1 are 12C tagged, whereas A2 and B2 are 13C tagged. Panel C and D show individual MS scans for peptide 9772 and 876, respectively. Masses of tagged peptides are in bold numerals.

highlights the advantage of detecting multiple peptides per protein in order to obtain more accurate quantitation. These 108

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data indicate that the mass tagging reagents can accurately label and report ratios of peptides in an artificial mixture of

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Figure 6. MALDI-TOF MS analysis of mass tagged peptides from two mixtures of protein digests. Two mixtures of protein digests were prepared containing different amounts of tryptic digests of BSA, OVA, and RNase. MALDI-TOF MS analysis of an aliquot of one of the crude mixtures (panel B) is shown along with a spectrum obtained from an aliquot of mass tagged peptides after reaction and cleavage (panel A). Panels C1 to C3 show spectra with an expanded scale (see panel A) obtained for tagged peptide pairs from OVA (panel C1), BSA (panel C2), and RNase (panel C3).

cysteine containing peptides. In order for them to be useful for quantitative measurements in more complex mixtures such as cell extracts, the mass tags must specifically label cysteinecontaining peptides mixed with noncysteine peptides. We therefore, tested the ability of the mass tags to react specifically with cysteine-containing peptides and to accurately detect relative concentrations from two mixtures of peptide digests. The mixtures contained tryptic digests of bovine serum albumin (BSA), ovalbumin (OVA) and ribonuclease A (RNase). Samples were prepared as follows. Sample 1 contained 50 pmol BSA, 250 pmol OVA, and 100 pmol RNase, whereas sample 2 contained 50 pmol BSA, 50 pmol OVA, and 200 pmol RNase. Sample 1 was mixed with 0.1 mg of 12C beads and sample 2 was mixed with the same amount of 13C beads in Tris-acetate buffer, pH 8.5. The suspensions were incubated separately for 1 h at room temperature and then further processed as described in the Experimental Section. Figure 6A demonstrates a MALDI-TOF mass spectrum after mass tagging showing pairs of tagged peptides from the protein digest mixture, whereas Figure 6B shows a MALDI-TOF mass spectrum of the complete peptide mixture before mass tagging, showing numerous peptide peaks. The comparison shows that the complexity of mass spectrum of the peptide mixture is greatly reduced after mass tagging. Only cysteine-containing peptide pairs are shown in the tagged mass spectrum. Major mass tagged peptide pairs seen in the spectrum shown in Figure 6A are listed in Table 2 along with the intensity ratios expected and observed in a single spectrum. Figure 6C(1-3) shows the mass spectra of detected ratios of pairs of peptides indicated as peaks 1, 2, and 3 in panel

A. In general, we found good agreement between expected and observed ratios for mass tagged peptides, especially in the case of BSA. A total of 13 unique cysteine containing peptides from BSA were identified out of a possible 35 along with 50% of the cysteine-containing peptides from OVA and RNase. These data, obtained from sample aliquots corresponding to low pmol levels of the protein digests, demonstrate that the solid phase mass tags are able to effectively recognize cysteine-containing peptides in this complex mixture. To positively identify the mass tagged peptides, we also separated them using online capillary LC/NanoESI-MS. Mass tagged peptides were eluted with a linear gradient of acetonitrile as described in the Experimental Section and data dependent MS/MS analysis was used to verify the identification of heavy and light peptide pairs. In this experiment we also examined the fragmentation pattern of the mass tag itself. Figures 7A-C illustrate 3 pairs of mass tagged peptides from BSA, OVA, and RNase, respectively from this LC/MS run. Intensities of the peaks corresponding to the 12C and 13C labeled peptides were again in good agreement with the expected ratios. Figure 7D illustrates an MS/MS spectrum derived from collision induced dissociation (CID) of the peptide shown in panel A. Many of the fragment ions from this mass tagged peptide were identified in the latter spectrum. Of particular interest is the tagged mass of cysteine-containing ions. In this peptide (MPC*TEDYLSLILNR), each of the b and y series ions retained the mass tag, (270 mass units) which remained intact throughout the CID procedure. This result and other data (not shown) show that the mass tag was not fragmented during the CID procedure. This eliminates comJournal of Proteome Research • Vol. 3, No. 1, 2004 109

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Figure 7. Analysis of mass tagged tryptic digest mixture by online reversed phase capillary LC/NanoESI-MS. The mixtures of BSA, OVA, and RNase tryptic digests prepared as in Figure 6 were analyzed by capillary LC/NanoESI-MS and pairs of tagged peptides from BSA (panel A), OVA (panel B), and RNase (panel C) are shown in the mass spectra. Masses of 12C and 13C labeled peptides are shown in bold. Panel D shows a CID MS/MS spectrum of a 12C tagged, BSA-derived peptide (m/z ) 968.7; panel A). The fragmentation pattern shows y and b ions that retained the intact mass of the 12C mass tag (270 Da). Table 2. Mass-Tagged Peptides Identified from Protein Digest Mixture by MALDI MS peptide sequence

peptide mass

mass w/12Ctag

mass w/13C tag

experimental ratio (12C/13C)

expected ratio

BSA

(R) LC*VLHEK (T) (K) QNC*DQFEK (L) (K) C*C*TESLVNR (R) (K) SLHTLFGDELC*K (V) (K) YIC*DNQDTISSK (L) (K) LKPDPNTLC*DEFK (A) (R) MPC*TEDYLSLILNR (L) (R)mPC*TEDYLSLILNR(L) (K) DDPHAC*YSTVFDKLK (H) (R) RPC*FSALTPDETYVPK (A) (K) LFTFHADIC*TLPDTEK (Q) (K) LFTFHADIC*TLPDTEKQIKK (Q) (K) GLVLIAFSQYLQQC*PFDEHVK (L) (R)FKDLGEEHFKGLVLIAFSQYLQQC*PF DEHVK (L)

841.9 1011.8 1024.9 1362.8 1387.7 1519.7 1667.9 1683.8 1738.8 1823.9 1850.9 2349.0 2435.1 3665.5

1111.9 1281.8 1564.9 1632.8 1657.7 1789.7 1937.9 1953.8 2008.8 2093.9 2120.9 2619.0 2705.1 3935.5

(R) ADHPFLFC*IK (H) (R) YPILPEYLQC*VK (E) (-)GSIGAASMEFC*FDVFK(E)

1190.9 1465.8 1750.8

1460.9 1735.8 2020.8

RNase

(R) ETGSSKYPNC*AYK (T) (K) HIIVAC*EGNPYVPVHFDASV (-) (K)NGQTNC*YQSYSTMSITDCR(E) (K) TTQANKHIIVAC*EGNPYVPVH FDASV (-)

1447.7 2166.8 2189.9 2810.8

1717.7 2436.8 2459.9 3080.8

0.82 1.10 1.00 1.13 1.08 1.06 0.95 1.05 1.60 1.02 1.12 1.12 0.89 0.93 1.06 5.12 2.86 2.25 3.41 1.18 0.42 0.78 0.85 0.81

1.00

OVA

1120.9 1290.8 1582.8 1641.8 1666.7 1798.8 1946.9 1962.8 2017.8 2102.9 2129.9 2628.0 2714.0 3944.5 average: 1469.8 1744.8 2029.7 average: 1726.7 2445.8 2468.8 3089.9 average:

protein

plications in the interpretation of MS/MS spectra and is particularly important during database search procedures such as Sequest commonly used in proteomics experiments. This experiment demonstrated the potential of this mass tagging approach for use in relative quantitation of proteins in complex mixtures. To demonstrate further the sensitivity of this approach for “real world” proteomics applications, another pair of mixtures of the protein digests of BSA, OVA and RNase was made. Mixture A contained 1 pmol of BSA, 5 pmol of OVA, and 2 pmol 110

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5.00

0.50

of RNase, whereas Mixture B contained 1 pmol, 1 pmol, and 4 pmol of the same proteins, respectively. Mixture A was tagged with the 12C and Mixture B with the 13C reagent. Samples were prepared as described above for the experiment shown in Figure 6 and also in the Experimental Section. Aliquots corresponding to 1/8th of the original samples (corresponding to 125 femtomol of BSA in Mixture A and 125 femtomol of both BSA and OVA in Mixture B) were analyzed by MALDI-TOF MS as shown in Figure 8. Ratios of peptides measured in this experiment mirrored those shown in Figure 6. The results are

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Figure 8. MALDI-TOF MS analysis of mass tagged peptides from low level mixtures of protein digests. Mixtures of tryptic digests of BSA, OVA, and RNase were prepared containing no more than 5 pmol of each digest and were mass tagged (see Results and Discussion). The tagged mixtures were prepared as described in Experimental and an aliquot containing 1/8th of the cleaved peptides, corresponding to the amounts of proteins indicated in the figure, was analyzed by MALDI-TOF MS. Insets show spectra with an expanded scale obtained for tagged peptide pairs from OVA (left), BSA (center) and RNase (right). These results compare favorably with those presented in Figure 6.

therefore consistent with the idea that low levels of peptides, such as those that are often encountered in proteomics research can be accurately assessed using this technique.

Conclusion This paper introduces a simple new solid phase mass tagging technique that offers several advantages over currently available technologies. The synthetic methods used for the tri-alanine peptides are straightforward and commonly available. Addition of a cysteine-reactive group on the N-terminus of the peptides is also accomplished by a straightforward reaction utilizing an activated ester of iodoacetic acid. In this context, we observe that several other potential cysteine-reactive (or other) groups could be added onto the N-terminus to give different reactivity and/or specificity to the tags. The flexible approach used here will also allow for addition of more alanine group(s) in order to create mass tags with larger mass differences between heavy and light reagents, if desired. The polymethacrylate resin used in these experiments provides a relatively inert surface with a large pore size that provides good mass transfer characteristics and cleavage is accomplished by treatment with TFA. Another advantage of this solid phase approach is the rapid sample preparation time and good sensitivity. The total time for sample preparation in our experiments was approximately 2 h, without including the time for preparation of peptide digests. In contrast, some mass tagging protocols requiring multiple chromatographic steps that require days to complete. We show here that the current mass tagging approach can accurately report peptide ratios either in mixtures of synthetic, cysteinecontaining peptides or in mixtures of tryptic peptide digests from several proteins. Sensitivity of the technique was found to be in the femtomol range, a level consistent with the requirements of proteomics research. Future experiments will

focus on the detection of protein expression differences from more complex biological samples.

Acknowledgment. This work was supported by Grant No. GM 20993 from the National Institutes of Health, U.S. Department of Health and Human Services and by an ongoing research collaboration with Thermofinnigan, Inc. This project was also funded in part with Federal funds from the National Heart, Lung, and Blood Institute, National Institutes of Health, under Contract No. N01-HV-28186. References (1) Hanash, S. Nature 2003, 422, 226-232. (2) Jeffery, D. A.; Bogyo, M. Cur. Opin. Biotechnol. 2003, 14, 87-95. (3) Chen, G.; Gharib, T. G.; Thomas, D. G.; Huang, C. C.; Misek, D. E.; Kuick, R. D.; Giordano, T. J.; Iannettoni, M. D.; Orringer, M. B.; Hanash, S. M.; Beer, D. G. Proteomics 2003, 3, 496-504. (4) Poirier, F.; Pontet, M.; Labas, V.; Le Caer, J. P.; Sghiouar-Imam, N.; Raphael, M.; Caron, M.; Joubert-Caron, R. Electrophoresis 2001, 22, 1867-1877. (5) Unlu, M.; Morgan, M. E.; Minden, J. S. Electrophoresis 1997, 18, 2071-2077. (6) Tao, W. A.; Aebersold, R. Curr. Opin. Biotechnol. 2003, 14, 110118. (7) Gygi, S. P.; Rist, B.; Gerber, S. A.; Turecek, F.; Gelb, M. H.; Aebersold, R. Nat. Biotechnol. 1999, 17, 994-999. (8) Zhang, R. J.; Sioma, C. S.; Wang, S. H.; Regnier, F. E. Anal. Chem. 2001, 73, 5142-5149. (9) Gygi, S. P.; Rist, B.; Griffin, T. J.; Eng, J.; Aebersold, R. J. Proteome Res. 2002, 1, 47-54. (10) Zhou, H. L.; Ranish, J. A.; Watts, J. D.; Aebersold, R. Nat. Biotechnol. 2002, 20, 512-515. (11) Qiu, Y. C.; Sousa, E. A.; Hewick, R. M.; Wang, J. H. Anal. Chem. 2002, 74, 4969-4979. (12) Xiang, R.; Horvath, C.; Wilkins, J. A. Anal. Chem. 2003, 75, 18191827. (13) Forns, P.; Fields, G. B. In Solid-phase synthesis; Kates, S. A., Albericio, F., Eds.; Marcel Dekker: New York, 2000, pp 1-78.

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