Catch-and-Release Reagents for Broadscale Quantitative Proteomics

Catch-and-Release Reagents for Broadscale Quantitative Proteomics Analyses Carlos A. Gartner, Joshua E. Elias, Corey E. Bakalarski, and Steven P. Gygi* Department of Cell Biology, Harvard Medical School, 240 Longwood Ave, Boston, Massachusetts 02115 Received November 16, 2006

The relative quantification of protein expression levels in different cell samples through the utilization of stable isotope dilution has become a standard method in the field of proteomics. We describe here the development of a new reductively cleavable reagent which facilitates the relative quantification of thousands of proteins from only tens of micrograms of starting protein. The ligand features a novel disulfide moiety that links biotin and a thiol-reactive entity. The disulfide is stable to reductive conditions employed during sample labeling but is readily cleaved under mild conditions using tris-(2-carboxyethyl) phosphine (TCEP). This unique chemical property allows for the facile use of immobilized avidin in a manner equivalent to the use of conventional reversible-binding affinity resins. Target peptides are bound to avidin resin, washed rigorously, then cleaved directly from the resin, resulting in simplified sample handling procedures and reduced nonspecific interactions. Here we demonstrate the stability of the linker under two different reducing conditions and show how this “catch-and-release (CAR)” reagent can be used to quantitatively compare protein abundances from two distinct cellular lysates. Starting with only 40 µg protein from each sample, 1840 individual proteins were identified in a single experiment. Using in-house software for automated peak integration, 1620 of these proteins were quantified for differential expression. Keywords: peptide • proteome • isotope • labeling • quantification • cleavable • cysteine • mass spectrometry (MS) • chromatography (HPLC) • automated • parallel • avidin • biotin • purification

Introduction Quantification of protein abundance changes has had a major impact on our understanding of their roles in health and disease. Changes in the expression of a protein resulting from a particular stimulus may indicate a role for that protein, either causal or effectual, in biological responses to that stimulus. A wide variety of factors with the potential to affect protein expression have been studied including the administration of xenobiotics,1 cell cycle progression states,2 and cancer,3,4 to name a few. Typically, the broadest proteome coverage has been sought to maximize identification of gene products that may play specific roles in the biological condition under consideration. Toward this goal, the development of new reagents and strategies that maximize procedural simplicity and expand quantification results are greatly valued by the investigator. The relative quantification of protein expression levels in different cell samples through the utilization of stable isotope dilution has become a standard method in the field of proteomics.5-7 We describe here a new reductively cleavable linker with chemical properties that make it ideal for use with isotope labeling protein quantification techniques. It exploits biotin as an affinity label for cysteine-containing peptide * To whom correspondence should be addressed. [email protected]; tel, (617) 432-3155; FAX, (617) 432-1144.

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fractionation and takes full advantage of conventional avidin’s high affinity while maximizing specificity. The “catch-andrelease (CAR)” reagent contains a novel hindered disulfide moiety incorporated into a linker that is completely stable to reductants employed during alkylation of sample cysteines but is readily cleaved with phosphine reductants. Although reducible disulfide-containing linkers have been employed in conjunction with biotin for target purification in the past,8,9 their labile nature, particularly with respect to the potential for sulfur scrambling with free cysteines, render these reagents far too reactive for use in quantitative proteomics applications. However, the development of a moiety that is inert to disulfide scrambling and is selectively cleavable ensures its utility as a reagent for quantitative applications and allows for the use of disposable immobilized avidin instead of its less desirable monomeric counterpart. Captured peptides can be thoroughly washed and gently eluted with concomitant removal of cleaved biotin-containing byproducts in a single step resulting in enriched target peptides derivitized with small mass tags that can be enriched with heavy isotopes. These tags were shown to provide linear and reproducible relative quantification, over a range of 2 orders of magnitude, of cysteine-containing peptides from a test protein differentially labeled with lightor heavy-labeled CAR reagents. Importantly, the small tag addition did not have a negative impact upon the quality of any MS/MS spectrum examined. Finally, as a proof-of-principle 10.1021/pr060605f CCC: $37.00

 2007 American Chemical Society

CAR for Quantitative Proteomics Analyses

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Figure 1. Catch-and-release (CAR) reagents and strategy for their utilization in protein profiling. (a) Structure of the reagent showing its principle structural features. The unique disulfide moiety imparts upon the linker a marked stability to reduction by DTT, but maintains sensitivity to TCEP-induced cleavage. The jagged line adjacent to the disulfide group denotes a mixture of isopropyl stereoisomers. The asterisks represent carbon atoms that carry isotopic density (13C) in the reagent’s heavy version. (b) General procedure for CAR analysis. Samples are fractionated and trypsinized after labeling protein cysteine residues. Capture of biotinylated peptides on conventional immobilized avidin and removal of impurities are followed by elution of target analytes directly from the resin by reduction with TCEP. The masses added to the peptides are very small (145 and 150 amu in light and heavy reagents, respectively).

experiment, we applied the method to compare the steadystate protein expression differences of two widely studied and disparate cancer cell lines (HeLa and HEK-293 cells). Forty micrograms of starting material from each cell type were used to identify more than 2000 proteins, over 85% of which were quantified. A number of these ratios were confirmed by Western blot analysis.

Results Stability of the Hindered Disulfide Linker to Reductive Conditions. The CAR reagent was designed with a unique disulfide bridge adjacent to a polyethoxyether linker that connects biotin and a cysteine-reactive entity (Figure 1a). It is sterically hindered by a carbon framework making it inert to cross-reactivity with other cysteine thiols or to reduction by dithiothreitol (DTT) under alkylating conditions. Tagged cysteine-containing peptides can be bound to conventional immobilized avidin, washed thoroughly, and readily eluted using tris-(2-carboxyethyl) phosphine (TCEP) as the reductant (Figure 1b). For the described reagent to be effective, the disulfide must be stable under conditions employed during reduction of sample disulfides as premature reduction would result in poor target recovery and irreproducible quantitative measurements. To demonstrate the stability of the linker under these condi-

tions, the iodoacetyl portion of the molecule was first quenched with D-penicillamine (D-Pen), a sterically hindered cysteine analog that does not react with the reagent’s disulfide moiety, even under extreme conditions (data not shown). This was done to avoid complications inherent to observation of multiple potentially reactive centers (the disulfide and the iodoacetyl functionalities) and their interactions with reductants. In each of these experiments, 5 pmol of the light CAR reagent/D-Pen conjugate were diluted in buffer alone (control) or incubated with varying amounts of DTT or TCEP under multiple conditions varying reaction temperature, duration, and reductant (Figure 2a). After stopping the reactions with formic acid, 5 pmol of heavy CAR reagent/D-Pen conjugate (containing five 13 C atoms) were added as an internal standard. An amount of sample corresponding to approximately 300 fmol of internal standard was subjected to capillary LC/MS analysis on a linear ion trap mass spectrometer. Each analysis was extracted as three separate chromatograms: One corresponded to the mass of the singly charged uncleaved heavy D-Pen conjugate (MH+ ) 832.4 amu) as the internal standard, and the second corresponded to the singly charged uncleaved light D-Pen conjugate being tested for stability to reduction (MH+ ) 827.4 amu). The third chromatogram corresponded to the mass of the singly charged cleaved biotin-containing product derived from the light D-Pen conjugate (MH+ ) 535.3 amu). Journal of Proteome Research • Vol. 6, No. 4, 2007 1483

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Figure 2. Measurement of CAR reagent stability to reductive conditions. (a) Conditions were tested for cleavage of a CAR reagent/Dpenicillamine conjugate. (b) Control incubation (no reductant) yielded equal amounts of target (light conjugate, red trace) and internal standard (heavy conjugate, black trace). (Inset) Mass spectrum of parent ions pertaining to both light and heavy conjugates. (c) Incubation with 10 mM DTT at room temperature shows no degradation of analyte relative to internal standard after 1 h. (d) Incubation at 50 °C with 10 mM TCEP cleaved over 97% of the conjugate in 1 h. The cleaved biotin-containing product eluted as a single peak just before 20 min.

Relative to control (Figure 2b), DTT in protein alkylation buffer containing 6 M urea and 0.05% SDS was completely ineffective in reducing the disulfide moiety at room temperature, even at 10 mM reductant concentration (Figure 2c). In this case, the light D-Pen conjugate abundance was indistinguishable from that of the heavy internal standard and no trace of cleaved product was observed. In contrast, 10 mM TCEP in buffer containing 20% methanol reduced >97% of the light D-Pen conjugate starting material within 60 min at 50 °C (Figure 2d). The appearance of cleaved product occurred concurrently with reduction in this case. In each example shown, the inset 1484

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depicts the mass spectrum derived from the CAR/D-Pen conjugates eluting just after 20 min in the chromatograms. Results of all reductive cleavage studies are summarized to include other conditions tested (Figure 3). Validation of CAR Reagent Use for Relative Quantification of a Test Protein: Linearity Studies with R-Lactalbumin. Bovine R-lactalbumin was used as a test protein to demonstrate the utility of CAR reagents for relative quantification of a protein by analysis of its cysteine-containing peptides. Equal amounts of the protein were alkylated with heavy- and light-labeled CAR reagents and mixed in differing proportions of heavy- to light-


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Figure 3. Summary of stability study for the CAR reagent/D-Pen conjugate. The disulfide moiety was stable to reduction by DTT in buffer A (alkylation buffer containing urea and SDS) under conditions used for protein derivatization. Reduction was achieved under mild conditions in buffer B (ammonium bicarbonate containing 20% methanol) using TCEP. Data shown are the average of triplicate experiments. Error bars denote standard deviation.

labeled material. Samples were run on a gradient SDS-PAGE gel and the bands excised. After in-gel tryptic digestion and purification on immobilized avidin beads, three cysteinecontaining peptides adducted with light or heavy cleaved CAR tags were quanitified by LC/MS analysis (Supplementary Figure 1, Supporting Information). The heavy-to-light ratios accurately reflected expected ratios within 10% based on relative amounts of material mixed at the protein level, and linearity was maintained over 2 orders of magnitude. The identities of recovered peptides were confirmed by sequencing in a separate LC-MS/MS analysis. Comparison of Protein Expression in HeLa and HEK-293 Cell Lines using CAR Reagents. Two widely studied and distinct cell lines were chosen for comparison by CAR analysis in order to ensure a broad diversity of relative protein expression levels (Figure 4a). Identical amounts of protein (40 µg) extracted from HeLa and HEK-293 cells were labeled with light and heavy CAR reagents, respectively. Labeled cell extracts were then combined and fractionated on a gradient SDS-PAGE gel. The entire lane was divided into 10 fractions of equal size, and each section was subjected to in-gel tryptic digestion. Recovered peptides were enriched for those containing cysteine using immobilized avidin beads. Fractions were treated in parallel by transferring peptide-bound avidin beads to 1 mL polypropylene filtration tubes (Supelco, Bellefonte, PA) after binding. All washing and recovery steps were performed in these tubes using a tabletop centrifuge (200 × g) to collect effluent in plastic culture tubes (13 × 100 mm). Resulting peptides were freed of polar impurities using the Stage tip method,10 and subjected to LC-MS/MS analysis on a hybrid linear ion trap Fourier transform ion cyclotron resonance (FTICR) mass spectrometer using a TOP10 method as described.11 The vast majority of peptide ions from all 10 fractions were detected as signals split into pairs of light- and heavy-labeled material in the FTICR survey scans. Labeled peptides were identified from their MS/MS spectra throughout each chromatographic run. More than 62 000 MS/ MS spectra were acquired during the analysis of the ten gel fractions. Tandem mass spectra were searched with the SE-

QUEST algorithm12 using a 100 ppm precursor ion tolerance against a composite database containing both the forward and reversed IPI human sequences.13 The use of this target/decoy database allowed us to determine an estimated false-positive rate for our dataset as described.14 A final list of peptide identifications was selected based on several factors including mass accuracy, tryptic state, Xcorr, ∆Cn, internal tryptic sites, and the presence of cysteine. The combined use of these factors resulted in an estimated false positive rate of 10 fold, were identified. (c) Examples of protein classes that are differentially expressed relative to overall protein ratio distribution. Of 1620 protein identifications for which quantitative data was recorded, 1396 mapped to biological processes according to the PANTHER classification system (https://panther.appliedbiosystems.com). Four of these processes were selected to represent classes that suggest differential expression between 293 and HeLa cells. Each data point represents a protein’s measured ratio as a function of the fraction of proteins with ratios less than (lower axis) this value within the depicted classification. Points that lie above zero on the y-axis indicate greater expression in 293 cells, whereas points that lie below indicate greater expression in HeLa cells.

In addition to the examples of EF2 and filamin A, the relative expression levels of five other proteins were confirmed by Western blot analysis (Supplementary Figure 2, Supporting Information). All examples were chosen based solely on the availability of appropriate antibodies.

Discussion Catch-and-release reagents represent an important technological advance in relative quantification of protein expression levels. Here we demonstrate the effectiveness and simplicity of the CAR method yielding abundant data from small amounts 1486


of starting material. The key chemical feature of this new tool is a cleavable disulfide moiety that was designed to be extraordinarily inert to reduction by DTT under conditions of cysteine alkylation, yet maintain a susceptibility to cleavage by phosphines (Figure 2). Steric hindrance provided by the disulfide-flanking carbon framework on the linker imparts upon the reagent the ability to easily withstand the reductive conditions employed during labeling of protein cysteine residues by the iodoacetyl terminus of the reagent. The two methyl carbons on the terminal side of the disulfide bridge also serve to carry isotopic label in the heavy CAR reagent counterpart. No


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Figure 5. Analysis of peptides recovered from immobilized avidin by reduction with TCEP as demonstrated for peptide E581-K594 from human elongation factor 2. All steps shown were performed via automated procedures as described in the text. (a) Base peak chromatogram showing the peak corresponding to the tagged peptide. (b) Extracted ion chromatograms corresponding to light (HeLa) and heavy (HEK-293) tagged peptides. Peak integration results agree with Western blot data (Inset, left). The mass spectrum of the chromatographic peak is included (Inset, right). (c) Peptide identity is derived from its tandem mass spectrum. The MS/MS spectrum of the heavy labeled peptide is shown. Journal of Proteome Research • Vol. 6, No. 4, 2007 1487

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Andover, MA) to bypass a synthetic step. The result was a heavy CAR reagent with five 13C labels instead of six. However, a total of seven positions are available for isotopic substitution if one also considers the amide nitrogen derived from nitromethane. (iii) Use of disposable conventional immobilized avidin- Lysis can be performed in biocompatible buffers under mild conditions allowing for the linker to be cleaved liberating labeled targets directly from immobilized avidin. Separate steps for removal of cleaved biotin are therefore not necessary since this portion of the molecule remains with the avidin resin and is discarded. The use of a disposable avidin source also allows for easy parallel sample handling and broadens the potential for high-throughput techniques. (iv) Small mass tags remaining with cysteine-containing peptides leading to easily interpretable MS/MS spectra- In no case has ligand-induced peptide fragmentation been observed to degrade the quality of CID spectra. This leads to MS/MS spectral data that are highly amenable to computer-based peptide sequence identification (for examples, see Supplementary Figure 2, Supporting Information).

Figure 6. Relative quantification of filamin A (FLNA) in HeLa and HEK-293 cells by two independent methods. A total of 22 unique cysteine-containing peptides served to confidently identify the protein, while 14 of these were used for quantification by the CAR method. Western blot analysis strongly supported the results.

evidence of disulfide reduction or cysteine-mediated sulfur exchange has been observed under alkylation conditions. Naturally, excessive stability to any reduction would render the reagent ineffective for its intended purpose, the recovery of tagged target molecules directly from conventional immobilized avidin beads. We found that the disulfide linker was readily cleaved, however, using TCEP as reductant in the presence of 20% methanol (Figure 2d). The critical importance of cosolvent or denaturant during reduction was discovered when 10 mM TCEP failed to cleave any detectable amount of CAR reagent/ D-Pen conjugate in buffer lacking methanol, even after 30 min at 50 °C (data not shown). Key features of the CAR strategy are summarized below. (i) Chromatographic equivalence of labeled peptides- The chromatographic coelution of light- and heavy-labeled peptides on LC-MS/MS analysis was achieved by the use of 13C labels in the heavy counterpart. (ii) Relatively low financial expense for isotope incorporation- In the current synthetic scheme, every position in the molecule that carries isotopic density is derived from readily available and relatively inexpensive 13C-labeled starting materials: acetone, nitromethane, and acetic acid (Supplementary Figure 3, Supporting Information). Although labeling of all six terminal carbon positions is preferred, we used a commercially available monoisotopically substituted iodoacetic acid source (Cambridge Isoptope Laboratories, 1488


To assess the scale of experiments that might be performed using the reagents, we compared protein expression ratios from two widely studied human cancer cell lines, HeLa and HEK293 cells. These samples are derived from vastly different cell types, epithelial and embryonic kidney cells, respectively, and were expected to show great differences in abundance of a large number of proteins. Just 40 µg of starting material from each cell type were utilized. After labeling in the presence of DTT, the protein samples were mixed and fractionated via SDSPAGE. Ten gel regions were digested with trypsin followed by avidin capture of cysteine-containing peptides. In parallel, all samples were simultaneously and vigorously washed, followed by gentle elution of target peptides with TCEP which allowed for greatly reduced levels of polymers commonly found in affinity isolation experiments. Nearly 2000 proteins were identified and more than 1600 of these quantified from a small amount of starting material suggesting that the CAR strategy will be applicable to proteome studies involving samples derived from limited amounts of primary tissue. Multiple ongoing investigations in our laboratory are already highlighting the value of this technique in tissue studies. To ensure correct identifications, researchers often exclude all proteins identified by a single peptide from further consideration,15 because most incorrect protein identifications fall into this category. As we have previously shown,13 the majority of these identifications are correct and represent a large fraction of all protein identifications. One strength of the CAR strategy is its ability to simplify complex peptide mixtures making a greater number of proteins available for identification. However, this necessarily reduces the number of tryptic peptides one can expect from any given protein, and renders a large portion of the data unusable if a multiple-peptides-per-protein standard is applied. Rather than dismissing this potentially valuable subset of proteins, we applied several lines of evidence in addition to algorithm-assigned scoring to adjust confidence in protein identification. These additional measurements include specific protease cleavage, ppm mass accuracy, the number of unique peptide spectral matches per proteins, and the signal-to-noise ratio of the ionized peptide in the precursor MS spectrum. Guided by the distribution of decoy database hits, we designed filtering constraints that excluded nearly all false positive protein identifications while retaining the majority of estimated true positives including those identified by single peptides. Ultimately, 1840 proteins were selected from 3427


unique peptide sequences with estimated false-positive rates of 0.4 and 0.2%, respectively. Results of automated protein quantification measurements using the VISTA algorithm are summarized (Figures 4b and 4c). Figure 4b shows the number of proteins identified from all fractions as a function of their 293-to-HeLa cell abundance ratios expressed in log(2). Large differences in protein expression were identified (for example, see Supplementary Figure 2, Supporting Information). The same data are expressed in Figure 4c via an alternative reprentation in which the y-axis represents the ratio of HEK-293 to HeLa cell derived proteins in log(2). The x-axis denotes the fraction of total quantified proteins with HEK-293:HeLa ratios lower than an individual protein being considered. For example, the protein with the highest HEK-293:HeLa ratio (y-value of approximately 6 in log(2)) was found at an x-value of 1.0, because its HEK-293:HeLa ratio was larger than 100% of other proteins quantified. Data corresponding to all proteins quantified are plotted in this manner as the sigmoidal black curve in Figure 4c. A few attributes of this data are worthy of note. For example, the y-value corresponding to the median lies just above 0.0, as described for Figure 4b. Upon careful consideration, it is not a surprise that the average ratio of quantified proteins lies closer to 1.2:1.0 than to the expected 1:1 ratio, as the two cell lines were purposely chosen for their expected marked protein expression differences. Samples were carefully combined after CAR labeling in a 1:1 ratio according to measurement of total protein performed in triplicate for each sample (mg protein/ mL). However, relative quantification reflects relative abundance (µmol protein/mL) such that small deviations from a 1:1 ratio at the abscissa median could be expected in samples as different as these. For example, a single protein that is very highly expressed in one cell line, but less so in the other, would be expected to shift the median significantly. Further support for the precision of the determined ratios lies in the very close agreement of ratios determined from different peptides within the same protein (Supplementary Table 2, Supporting Information). Still more evidence for this assessment rests in the finding that ratios derived for a subset of proteins annotated according to the PANTHER classification system (https://panther.appliedbiosystems.com) form general trends in Figure 4c indicating that certain pathways are more abundant in one cell line than the other. Four such subsets are included on the graph demonstrating that DNA repair proteins and those involved in oxidative phosphorylation are expressed to a higher degree in HEK-293 cells. Conversely, cell adhesion proteins and surface receptor mediated signal transduction components are expressed to a higher degree in HeLa cells. Analysis of such trends will certainly be facilitated in the future when coupled with CAR analysis. In conclusion, we have presented a novel reductively cleavable reagent for large-scale quantitative proteomics experiments. We demonstrated its use by identifying nearly 2000 proteins from HeLa and HEK-293 cells and determining relative expression levels for more than 1600 proteins in a matter of days. With its utility and simplicity, it is anticipated that this technique will facilitate quantitative proteomics experiments on a large scale.

Methods Preparation of CAR Reagents. General steps to the chemical synthesis of CAR reagents are shown in Supplementary Figure 3 (see Supporting Information).

research articles Disulfide Stability Testing. The iodoacetyl moiety of the reagents was first quenched with D-Pen in order to study the reactivity of the disulfide itself. In separate tubes, 1.0 mM lightor heavy-labeled CAR reagents were prepared in 50 mM ammonium bicarbonate buffer (pH ) 8.3) from a 35 mM stock in storage solution (50% acetonitrile in 0.01% TFA). In a separate tube, a 10 mM stock of D-Pen was prepared in 50 mM ammonium bicarbonate (pH ) 8.3). The CAR reagent solutions were combined 1:1 with the D-Pen solution in separate tubes and incubated for 2 h in the dark at room temperature. The final concentration of each CAR/D-Pen conjugate stock solution was 0.5 mM. Conjugate stability to reduction by DTT was tested by diluting light- and heavy-labeled CAR/D-Pen to 10 µM in 100 mM Tris, (pH ) 8.3) containing 6 M urea, 5 mM EDTA, and 0.05% SDS. A stock solution of DTT was prepared in this buffer at double its target concentration. To a separate 0.5 mL tube was transferred 5 µL of light-labeled conjugate (25 pmol) measured with a syringe. An equal volume of DTT solution freshly prepared at double its target concentration in the alkylation buffer was then added to initiate reaction. Reactions were stopped at the appropriate time points using 185 µL of 7.5% acetonitrile containing 5% formic acid. The heavy-labeled CAR/D-Pen conjugate was then added (5 µL measured with a syringe, 25 pmol) as an internal standard. The entire sample was then shaken for 10 min with 10 µL MonoQ resin previously washed with 7.5% acetonitrile containing 5% formic acid to remove SDS. A 30 µL portion of the sample was removed and dried on a centrifugal evaporator. Nonvolatile buffer components were removed by the Stage tip method10 and the remaining material resuspended in 30 µL 7.5% acetonitrile containing 5% formic acid. Analysis by microcapillary LC/MS was performed on a LTQ mass spectrometer using 2 µL of each sample (approximately 300 fmol internal standard). The same procedure was employed to test cleavage conditions using TCEP as reductant, except that 50 mM ammonium bicarbonate (pH ) 8.3) containing 20% methanol was used as solvent. In cases where the reaction was performed at 50 °C, both reaction components were preincubated at that temperature for 5 min prior to reaction initiation. As neither SDS nor urea were used in the cleavage experiments, the MonoQ and Stage tip purification steps were omitted and 2 µL sample were used directly for LC/MS analysis immediately after reaction quenching and internal standard addition. It is important to note that the highest purity of TCEP (BioVectra dcl, Oxford CT) was necessary to avoid the appearance of excessive polymeric impurities upon mass spectral analysis. In every case described here, TCEP stock solutions were freshly prepared at a concentration of 100 mM in 350 mM ammonium hydroxide and diluted to its target concentration in an appropriate buffer. The pH of each solution was tested with pH strips to ensure it was within the proper range. Validation of Relative Protein Abundance Measurements using Bovine R-Lactalbumin. Bovine R-lactalbumin, 40 µg, was dissolved in 50 µL of a cysteine alkylation buffer previously described (100 mM Tris, pH ) 8.3, 6 M urea, 5 mM EDTA, 0.05% SDS).16 The sample was degassed at reduced pressure (0.3 Torr) on a centrifugal evaporator for 30 s. It was then brought to 1 mM DTT from a 25 mM stock freshly prepared in water and divided in half. After reducing at 50 °C for 30 min, the samples were brought to room temperature and light or heavy CAR reagent added to 5 mM final concentration from a 35 mM stock (stored in 50% acetonitrile containing 0.01% TFA). Journal of Proteome Research • Vol. 6, No. 4, 2007 1489

research articles Samples were alkylated in the dark for 2.5 h at room temperature and quenched with 30 mM D-Pen from a 300 mM stock freshly prepared in water. After 1 h of quenching in the dark at room temperature, the samples could be stored at 4 °C overnight if desired. Light- and heavy-labeled proteins were then mixed in appropriate ratios and 2 µg total protein per lane were applied to a 1 mm 12% SDS-PAGE gel. No DTT was used in the sample buffer as all cysteines were considered alkylated. After Coomassie staining and band excision, in-gel tryptic digestion was performed as previously described17 and the peptides extracted. After drying each sample by vacuum centrifugation, peptides were redissolved in 10 µL of 7.5% acetonitrile containing 5% formic acid. They were then diluted with 150 µL of avidin binding buffer (250 mM sodium phosphate [pH ) 6.5] containing 10% glycerol). A blank sample was also prepared in parallel to ensure that the resulting sample pH was between 5 and 6. In a separate tube, immobilized avidin resin (Pierce, Rockford, IL) was washed twice with excess avidin wash buffer (50 mM sodium phosphate [pH ) 6.0] containing 10% glycerol) and the resin distributed into 0.5 mL tubes, 20 µL packed resin per sample. The wash buffer was removed from each avidin-containing tube with a syringe and peptide samples added. Tubes were shaken at room temperature for 1 h, at which point they were centrifuged and buffer removed with a syringe. The resin was washed twice with constant shaking using 250 µL avidin wash buffer each time. Each sample was then washed three times with 250 µL avidin final wash buffer (50 mM ammonium bicarbonate [pH ) 8.3] containing 5 mM EDTA and 20% methanol) before incubating the beads with 50 µL biotin cleavage buffer (avidin final wash buffer containing 5 mM TCEP). The beads were then incubated at 50 °C for 90 min in cleavage buffer. Samples were agitated briefly every 30 min to ensure accessibility of immobilized peptides to the TCEP. After cooling to room temperature, the overlays were removed with a syringe and saved in fresh tubes. Resin from each sample was washed twice with 75 µL of avidin final wash buffer at room temperature for 15 min with shaking. Washes were combined with their corresponding recovered samples and acidified using 10 µL acetic acid. After evaporation to dryness on a vacuum centrifuge, peptides were freed of contaminants using the Stage tip method10 and evaporated again before analysis by LC-MS/MS using the LTQ FT mass spectrometer. Comparison of Protein Expression Levels from Two Cell Lines using CAR Reagents. E1A-transformed Human Embryonic Kidney (HEK) 293 cells (gift from Edward Harlow, Massachusetts General Hospital, Boston, MA) and HeLa cells (American Type Culture Collection, Manassas, VA) were maintained in Dulbecco’s Modified Eagle’s Medium (Mediatech, Herdon, VA) supplemented with 10% fetal bovine serum (Hyclone, Logan, UT), penicillin (20 inhibitory units/mL) and streptomycin (20 µg/mL). Upon reaching confluence cells were washed once with ice-cold PBS and lysed to approximately 2 mg/mL protein in 50 mM Tris buffer (pH ) 7.4) containing 150 mM sodium chloride, 2 mM EDTA, 1% octyl glucoside, 0.2% SDS, 0.5% cholate, and EDTA-free Complete Mini protease inhibitor cocktail (Roche Diagnostics, Mannheim, Germany) included according to manufacturer’s instructions. Protein concentration was measured in triplicate by the BCA assay (Pierce, Rockford, IL). Samples were diluted 1:1 with cysteine alkylation buffer (100 mM Tris [pH ) 8.3] containing 6 M urea, 5 mM EDTA, and 0.05% SDS) and dialyzed into 1 L of that buffer for 3 h using Slide-A-Lyzer MINI dialysis cups (Pierce, 1490


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Rockford, IL). Protein concentration was again measured and 40 µg of each sample placed into fresh tubes. After a 30 s degassing period on a vacuum centrifuge (0.3 Torr), samples were brought to 1 mM DTT from a 25 mM stock freshly prepared in water. Reduction was carried out for 30 min at 50 °C and the samples cooled to room temperature. Light-labeled CAR reagent was added to the HeLa cell lysate at 5 mM final concentration from its 35 mM stock in storage solvent, whereas HEK 293 lysate was alkylated with the heavy-labeled reagent. Reaction was allowed to proceed for 2.5 h in the dark at room temperature before quenching with 30 mM D-Pen added from a 500 mM stock freshly prepared in water. After 1 h of quenching at room temperature, samples were stored at 4 °C overnight. Samples were combined and fractionated by SDS-PAGE on a 1.5 mm gradient gel (NuPage 4-12% Bis-Tris, Invitrogen, Carlsbad, CA) and stained with Coomassie Brilliant Blue. The entire lane was divided into 10 sections of equal size and ingel trypsin digestion performed on each fraction. After peptide extraction and drying, biotinylated fragments were isolated and purified on immobilized avidin resin exactly as described for bovine R-lactalbumin analysis. However, all washing and elution steps were done in parallel using empty solid phase extraction tubes (Supelco, Bellefonte, PA). The entire portion of each sample recovered was then analyzed by microcapillary LC-FTICR-MS/MS analysis as described below. LC-MS and LC-MS/MS Analyses. Liquid chromatography tandem mass spectrometry (LC-MS/MS) was performed using the LTQ FT, which is a hybrid linear (2-D) ion trap-Fourier transform ion cyclotron resonance (FTICR) mass spectrometer (7 T, ThermoElectron, San Jose, CA) as described.11 Briefly, the entire volume of each reconstituted sample was loaded onto a 125 µm × 18 cm fused silica C18 (Magic C18-AQ, 200 Å pore size, 5 µm diameter, Michrom BioResources, Auburn, CA) microcapillary column using a FAMOS capillary autosampler (LC Packings, Sunnyvale, CA) and an Agilent 1100 series binary HPLC pump (Agilent Corporation, Palo Alto, CA) with an inline flow splitter. Peptides were transferred from the autosampler directly to the resolving column for 20 min at a pressure of 120 bar in Buffer A (3% acetonitrile, 0.15% formic acid), followed by gradient elution at 60 bar from 7 to 33% Buffer B (97% acetonitrile, 0.15% formic acid) over 55 min. Effluent was directed into a nanospray source of the mass spectrometer operating at a 2.1 kV source potential. During gradient elution, ten ion-trap MS/MS spectra were acquired per data-dependent cycle from a high-resolution (R > 70 000 @ 500 m/z) FTICR survey spectrum. Database Searching and Data Processing. Accurate precursor ion masses, MS/MS spectra, and chromatographic information were extracted from the raw file output of the LTQ FT with in-house software as described.18 Tandem mass spectra, represented in the dta file format, were searched with the SEQUEST algorithm (version 27, revision 12)12 against a concatenated target (forward) and decoy (reversed) human IPI protein database (ftp.ebi.ac.uk/pub/databases/IPI/current, downloaded on July 4, 2005) with the following restrictions: mass tolerance of (100 parts-per-million; at least one tryptic terminus per peptide; up to two internal cleavage sites per peptide; dynamic modifications of 15.99492 for methionine to account for oxidation; static modifications of 145.05613 on all cysteines to account for the mass addition of the cleaved light reagent; dynamic modifications of 5.01677 on cysteines to account for the potential mass difference of the heavy reagent. Search

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results and chromatographic data were stored in a PostgreSQL database. The SEQUEST-derived scores XCorr, ∆Cn as well as partsper-million accuracy were used to derive filtering cutoffs of non-redundant top-ranked peptide-spectral matches. An inhouse algorithm similar in principle to one previously described19 adjusted threshold values for these measurements to maximize sensitivity while maintaining a < 1% false positive rate, as estimated by the number of decoy database hits.13,14 All peptide matches were secondarily required to have two tryptic termini, and to contain at least one cysteine residue. This score-independent method for enriching for correct identifications allowed for the use of fairly non-stringent scorebased filtering criteria13: Charge ) 2+: XCorr g 1.3, ∆Cn g 0.01; Charge ) 3+: XCorr ) 1.7, ∆Cn g 0.03; Charge ) 4+: XCorr g 1.8, ∆Cn g 0.01. Analysis was performed on data collected from each gel fraction independently. Protein identifications were assembled from their constituent identified peptides in Microsoft Excel. A given protein was selected as being confidently assigned if at least one uniquely identified peptide from the protein exceeded filtering criteria described above, had zero internal cleavage sites, and had two tryptic ends. All top-ranked cysteine-containing peptides returned by SEQUEST were submitted for quantitative analysis by an automated software suite, VISTA, which we have used previously11,20,21 (Bakalarski et al., manuscript in preparation). Briefly, the theoretical masses of the heavy- and light-labeled peptides were calculated from their sequence composition information and these masses used to extract precursor ion chromatogram intensities of each variant from FTICR MS spectra. Extracted candidate peaks were filtered for a mass accuracy of better than (20 ppm and for the presence of the predicted isotopic distribution for the peptide. For each isotopic variant, the area under the curve was separately determined as a function of elution time, and the relative peptide abundance between the two samples reported as a ratio of their respective areas. Quantified peptides were scored for quality using empirically determined test conditions. Proteins were considered successfully quantified if they met the identification criteria described above, met minimal ratio quality test conditions, and agreed with measured ratios across independent identifications of the protein. Manual validation of many hundreds of chromatograms was also performed.

Acknowledgment. Support for this work was provided by NIH grants (HG003456 and GM67945). We thank members of the Gygi lab for helpful discussions and S. Gerber for technical assistance. Supporting Information Available: Supplementary Figures 1-3 and Tables 1 and 2. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Welch, K. D.; et al. Proteomic identification of potential susceptibility factors in drug-induced liver disease. Chem. Res. Toxicol. 2005, 18, 924-933.

(2) Erhardt, N. M.; Haines, L. R.; Pearson, T. W.; Sherwood, N. M. Inhibition of PACAP Activity by a Receptor Antagonist Results in Changes in Cell Cycle and Apoptotic Proteins in Chick Neuroblasts. J. Mol. Neurosci. 2005, 27, 107-124. (3) Barnidge, D. R.; Jelinek, D. F.; Muddiman, D. C.; Kay, N. E. Quantitative protein expression analysis of CLL B cells from mutated and unmutated IgV(H) subgroups using acid-cleavable isotope-coded affinity tag reagents. J. Proteome Res. 2005, 4, 1310-1317. (4) Qi, Y.; Chiu, J. F.; Wang, L.; Kwong, D. L.; He, Q. Y. Comparative proteomic analysis of esophageal squamous cell carcinoma. Proteomics 2005, 5, 2960-2971. (5) Ornstein, D. K.; Petricoin, E. F., 3rd. Proteomics to diagnose human tumors and provide prognostic information. Oncology (Williston Park) 2004, 18, 521-529; discussion 529-532. (6) Ciordia, S.; de Los Rios, V.; Albar, J. P. Contributions of advanced proteomics technologies to cancer diagnosis. Clin. Transl. Oncol. 2006, 8, 566-580. (7) Freeman, W. M.; Hemby, S. E. Proteomics for protein expression profiling in neuroscience. Neurochem. Res. 2004, 29, 1065-1081. (8) Trotti, D.; et al. Amyotrophic lateral sclerosis-linked glutamate transporter mutant has impaired glutamate clearance capacity. J. Biol. Chem. 2001, 276, 576-582. (9) Huh, K. H.; Wenthold, R. J. Turnover analysis of glutamate receptors identifies a rapidly degraded pool of the N-methyl-Daspartate receptor subunit, NR1, in cultured cerebellar granule cells. J. Biol. Chem. 1999; 274, 151-157. (10) Rappsilber, J.; Ishihama, Y.; Mann, M. Stop and go extraction tips for matrix-assisted laser desorption/ionization, nanoelectrospray, and LC/MS sample pretreatment in proteomics. Anal. Chem. 2003, 75, 663-670. (11) Haas, W. et al. Optimization and use of peptide mass measurement accuracy in shotgun proteomics. Mol. Cell. Proteomics 2006, 5, 1326-1337. (12) Eng, J. K.; McCormack, A. L.; Yates, J. R., III. An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 1994, 5, 976-989. (13) Elias, J. E.; Haas, W.; Faherty, B. K.; Gygi, S. P. Comparative evaluation of mass spectrometry platforms used in large-scale proteomics investigations. Nat. Methods 2005, 2, 667-675. (14) Peng, J.; et al. A proteomics approach to understanding protein ubiquitination. Nat. Biotechnol. 2003, 21, 921-926. (15) Kratchmarova, I.; Blagoev, B.; Haack-Sorensen, M.; Kassem, M.; Mann, M. Mechanism of divergent growth factor effects in mesenchymal stem cell differentiation. Science 2005, 308, 14721477. (16) Smolka, M. B.; Zhou, H.; Purkayastha, S.; Aebersold, R. Optimization of the isotope-coded affinity tag-labeling procedure for quantitative proteome analysis. Anal. Biochem. 2001, 297, 2531. (17) Shevchenko, A.; Wilm, M.; Vorm, O.; Mann, M. Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 1996, 68, 850-858. (18) Beausoleil, S. A.; Villen, J.; Gerber, S. A.; Rush, J.; Gygi, S. P. A probability-based approach for high-throughput protein phosphorylation analysis and site localization. Nat. Biotechnol. 2006, 24, 1285-1292. (19) Kislinger, T.; et al. PRISM, a generic large scale proteomic investigation strategy for mammals. Mol. Cell. Proteomics 2003, 2, 96-106. (20) Everley, P. A.; et al. Enhanced analysis of metastatic prostate cancer using stable isotopes and high mass accuracy instrumentation. J. Proteome Res. 2006, 5, 1224-1231. (21) Neher, S. B.; et al. Proteomic profiling of ClpXP substrates after DNA damage reveals extensive instability within SOS regulon. Mol. Cell 2006, 22, 193-204.

PR060605F

Journal of Proteome Research • Vol. 6, No. 4, 2007 1491

Catch-and-Release Reagents for Broadscale Quantitative Proteomics

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