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Originally published In Press as doi:10.1074/mcp.M700057-MCP200 on July 11, 2007.
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Molecular & Cellular Proteomics 6:1771-1777, 2007.
© 2007 by The American Society for Biochemistry and Molecular Biology, Inc.

Research

Assessing Enzyme Activities Using Stable Isotope Labeling and Mass Spectrometry *,S

Patrick A. Everley{ddagger},§, Carlos A. Gartner{ddagger}, Wilhelm Haas{ddagger}, Alan Saghatelian, Joshua E. Elias{ddagger}, Benjamin F. Cravatt, Bruce R. Zetter§ and Steven P. Gygi{ddagger},||

From the {ddagger} Department of Cell Biology, Harvard Medical School, Boston, Massachusetts 02115, § Program in Vascular Biology, Department of Surgery, Children's Hospital, Boston, Massachusetts 02115, and The Skaggs Institute for Chemical Biology and Department of Cell Biology, The Scripps Research Institute, La Jolla, California 92037


   ABSTRACT
TOP
 ABSTRACT
EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
Activity-based protein profiling has emerged as a valuable technology for labeling, enriching, and assessing protein activities from complex mixtures. This is primarily accomplished via a two-step identification and quantification process. Here we show a highly quantitative and streamlined method, termed catch-and-release activity profiling of enzymes (CAPE), which reduces this procedure to a single step. Furthermore the CAPE approach has the ability to detect small quantitative changes that may have been missed by alternative mass spectrometry-based techniques.

Mass spectrometry and stable isotopes have recently surfaced as fundamental tools for large scale proteomics analyses. Among these techniques are ICAT (1) and stable isotope labeling by amino acids in cell culture (SILAC)1 (2), which have recently been used to study an array of biological questions (3–7). These technologies utilize differential labeling of proteins with carbon and nitrogen stable isotopes, giving rise to isotopically "light" and "heavy" samples. The major strength of these methods is their direct application to mass spectrometry, which can yield rapid protein identification. Quantification can also be achieved by comparing intensities of light and heavy peptide peaks. Although these methods are useful for obtaining quantitative measurements of protein levels, they are strongly biased toward the most abundant proteins.

To go beyond a mere listing of general protein levels, alternative methods must be used for gaining better sensitivity and readout of protein activity. One of the most elegant among these procedures is activity-based protein profiling (ABPP) (8), which includes the use of functional tags that specifically bind to and covalently modify active enzymes but not their endogenously inhibited or inactive zymogen forms. By only labeling functional proteins within a specific enzyme class, catalytically active proteins can be selectively enriched and subsequently quantified to yield relative activity levels between biological samples. Functional probes for many classes of enzymes have already been described, including those specific for serine hydrolases, metalloproteases, kinases, and others (9, 10). To date, however, characterization of proteins by ABPP has been inefficient by requiring two steps. Qualitative information is acquired from mass spectrometry, whereas accurate quantitative data must be obtained from the relative in-gel fluorescence intensities of the labeled enzymes (11). Recent gel-free studies have attempted to simplify this process but were limited by the semiquantitative nature of the analysis (12) or were still reliant on a two-step procedure using capillary electrophoresis and mass spectrometry (13). A highly quantitative, streamlined approach that takes advantage of stable isotope labeling and mass spectrometry would greatly benefit the field of functional proteomics.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
 REFERENCES
 
Synthesis of Cleavable Fluorophosphonate Catch-and-release (FP-CAR) Probe—
Reaction of isotopically light intermediate reactant (Reaction 1, Y) from Gartner et al. (14) and an alkyl chain-linked fluorophosphonate (Reaction 1, X) was carried out in N,N-dimethylformamide (Reaction 1, DMF) for 4 h as described previously (15) to yield final FP-CAR reagent (Reaction 1, Z). High resolution mass spectrometry was performed on the final product to confirm the expected mass (Supplemental Fig. 1).


Figure 4
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  		REACTION 1
 
FP-CAR Reactions with Trypsin and Trypsinogen—
Bovine trypsin (acetylated) and trypsinogen were purchased from Sigma. Trypsin and trypsinogen were incubated at 10 µg/ml (UV-visible) in 50 mm Tris-HCl (pH 8.0) with 5 µm FP-CAR reagent for 1 h at room temperature. FP-CAR reaction conditions (5 µm for 1 h at room temperature) were consistent throughout this study. For silver staining, equal amounts of protein (225 ng) were run on a 1-mm 4–12% gradient gel (Invitrogen) and stained with silver. For avidin blotting, equal molar amounts of protein (100 fmol) were resolved by SDS-PAGE as described above and transferred to PVDF. The membrane was blocked in casein (Vector Laboratories, Burlingame, CA) followed by incubation with avidin-horseradish peroxidase (1:10,000 dilution, Vector Laboratories) for 1 h at room temperature. After five washes in PBS-Tween 20, the membrane was treated with Western Lightning Chemiluminescence Reagent Plus (PerkinElmer Life Sciences) and exposed to Hyblot CL autoradiography film. Trypsin (10 µg/ml) was treated with a 3 m excess (26 µg) of soybean trypsin inhibitor for 2 h at room temperature (compared with untreated control) followed by incubation with FP-CAR reagent. The reaction was carried out at pH 7.0 because of soybean trypsin inhibitor instability under basic conditions. Silver staining and avidin blotting were carried out as described above. Trypsinogen (1 µm) was treated with enteropeptidase (1 nm, Sigma) for 30 min (compared with untreated control) followed by incubation with FP-CAR reagent. The enteropeptidase reaction was carried out in sodium citrate buffer at pH 5.6 to prevent autoactivation of trypsinogen (16). Protein was subjected to silver staining and avidin blotting as described above with 1 pmol of protein used for avidin blotting.

For mass spectrometry analysis of the trypsin active site peptide, unmodified bovine trypsin (Sigma) was incubated at 10 µg/ml in 50 mm Tris-HCl (pH 8.0) with 5 µm FP-CAR reagent for 1 h at room temperature. Protein was separated by SDS-PAGE and stained with Coomassie Blue. After destaining, the band was excised and processed for reduction/alkylation. Briefly gel pieces were reduced with tris(2-carboxyethyl)phosphine for 90 min at 50 °C followed by alkylation with iodoacetamide. The sample was incubated with trypsin overnight and prepared for tandem mass spectrometry by passage through a C18 STAGE tip (17).

Cell Culture and Protein Preparation—
LNCaP and LNCaP-LN3 cells were plated at 1 x 106 cells/15-cm dish in eight dishes each and cultured as described previously (18). Briefly cells were grown in RPMI 1640 medium (Invitrogen) with dialyzed serum (Invitrogen). LNCaP cells were cultured with [12C,14N]lysine and [12C,14N]arginine (light condition), whereas LNCaP-LN3 medium contained [13C6,15N2]lysine and [13C6,15N4]arginine (Isotec, Sigma) (heavy condition). Media were supplemented with 1 nm dihydrotestosterone to promote optimal growth of testosterone-sensitive prostate cancer cells (19). Cells were cultured for 6–7 days, changing media every 2–3 days until 70–80% confluency was reached. (Note that cells grown under dialyzed conditions showed similar growth rates and no noticeable differences in morphology compared with cells cultured with non-dialyzed serum.) For whole-cell lysate (WCL) protein preparation, plates were washed three times in ice-cold PBS and collected in 50 mm Tris-HCl (pH 8.0) and 1% Triton X-100. Cells were homogenized with a Dounce tissue grinder 20 times on ice and ultracentrifuged at 20,000 x g for 5 min at 4 °C. Pellets were discarded, and supernatant protein concentrations were determined by BCA assay (Pierce).

FP-CAR Reaction with Cell Lysates—
WCLs from LNCaP and LNCaP-LN3 were incubated with 5 µm FP-CAR reagent for 1 h at room temperature. After addition of protease inhibitors (Roche Applied Science) and phosphatase inhibitors (25 mm NaF, 10 mm Na4P2O7, and 1 mm Na3VO4), protein was mixed 1:1 and dialyzed in buffer containing 10% methanol, 50 mm ammonium bicarbonate, 5 mm EDTA, and 0.05% SDS for 4 h at room temperature to remove excess FP-CAR reagent. Three-hundred microliters of avidin solution (50% slurry of immobilized avidin, Pierce) were added to the protein sample and rocked overnight at 4 °C. The protein sample was transferred to a column with nonspecific avidin binding washed with 50 ml of Wash Buffer 1 (100 mm ammonium bicarbonate, 2 m urea, 0.1% SDS, 150 mm NaCl, 0.5% Triton X-100, and 20% glycerol) followed by 50 ml of Wash Buffer 2 (100 mm ammonium bicarbonate, 10% methanol, and 0.5% octyl glucoside (w/v)). Beads were then transferred to Eppendorf tubes and reduced in tris(2-carboxyethyl)phosphine (10 mm in Wash Buffer 2) for 90 min at 50 °C to cleave FP-CAR reagent and release proteins bound to avidin. Overlays were removed and saved, and avidin resin was washed twice in Wash Buffer 2 for 15 min with shaking. Washes were combined with overlays followed by TCA precipitation, reduction, and alkylation. In-solution trypsin digestion was performed overnight, and the sample was acidified with acetic acid and evaporated in vacuo. Peptides were purified with C18 STAGE tips before they were subjected to LC-MS/MS analyses.

LC-MS/MS—
Microcapillary liquid chromatography electrospray ionization tandem mass spectrometry (LC-MS/MS) was performed on a hybrid linear quadrupole ion trap/Orbitrap (LTQ Orbitrap) (20) or a hybrid linear quadrupole ion trap/Fourier transform ion cyclotron resonance (LTQ FT) (21) mass spectrometer (Thermo Electron, San Jose, CA) equipped with an Agilent 1100 Series binary HPLC pump (Agilent Technologies, Palo Alto, CA) and a Famos autosampler (LC Packings, Sunnyvale, CA). Peptides were separated on fused silica microcapillary columns (inner diameter, 125 µm; approximately 5 µm at the tip) that were in-house packed to a length of 18 cm with a C18 reversed phase resin (Magic C18AQ, 5-µm particle size, 200-Å pore size; Michrom BioResources, Auburn, CA). Separation was achieved by applying a gradient from 5 to 30% acetonitrile in 0.125% formic acid at a flow rate of 300 nl/min for 60 min (catch-and-release activity profiling of enzymes (CAPE) experiments, LTQ Orbitrap), 90 min (SILAC experiments, LTQ Orbitrap), or 35 min (characterization of trypsin active site peptide, LTQ FT, gradient from 5 to 70% acetonitrile). MS and MS/MS data were acquired in a data-dependent fashion: 10 LTQ MS/MS spectra were acquired on the most abundant ions detected in a low resolution master spectrum, whereas high resolution MS scans were collected in which 1 x 106 ions were measured over an m/z range of 375–1800 (resolution settings were 60,000 for the Orbitrap and 100,000 for the FT).

Database Searching of MS/MS Data—
MS/MS spectra were searched with a fully tryptic enzyme constraint (two tryptic termini and up to two internal missed cleavages) against a concatenated target (forward orientation) and decoy (reversed orientation) human International Protein Index (IPI) protein database (ftp.ebi.ac.uk/pub/databases/IPI/current/) using the SEQUEST algorithm (22) and a mass tolerance of ±200 ppm. Searches were performed with dynamic modifications on methionine (oxidation), lysine, and arginine with masses 15.9949, 8.0142, and 10.0083 Da, respectively, along with a static modification of 57.0214 Da on cysteines (alkylation). Using in-house software similar in principle to that developed by Kislinger et al. (23), {Delta}Cn, XCorr, and ppm thresholds were optimized for peptide charge states of 1, 2, 3, and 4 to yield an estimated 1% false positive rate (24) with maximum sensitivity.

Peptide and Protein Quantification—
Peptides were quantified with the VISTA program as described previously (18). Although CAPE led to the identification of 25 hydrolase proteins (Supplemental Table 1), only peptides with the highest VISTA confidence scores were used for quantification, and only proteins detected with at least two quantified peptides in each of the four experiments (control and experimental with biological replicates for each) were considered for analysis. We note that additional non-serine hydrolase proteins were identified in our study predominantly due to the LTQ instrument sensitivity. We also note that the vast majority of these proteins in our raw dataset were single hit identifications and thus excluded from analysis. Peptide ratios were determined by dividing area under the curve of the extracted chromatograms for the heavy and light peptides, respectively. Ratios were then subjected to log2 transformation, and redundant peptides were excluded from analysis based on confidence measurements including VISTA score, XCorr, and signal to noise ratio values. For purposes of identification and quantification, peptides were considered unique if they had different charges or different states of methionine oxidation. Non-redundant peptide ratios were averaged from each protein to calculate an average protein ratio.

SILAC Analyses of LNCaP and LNCaP-LN3 Cells—
For SILAC analyses, WCLs collected from LNCaP and LNCaP-LN3 were mixed at a 1:1 ratio by protein concentration (200 µg from each, 400 µg total), boiled for 3 min in sample buffer, and resolved with a NuPAGE 4–12% gradient gel (Invitrogen). The gel was stained with Coomassie Blue and processed into 10 samples as described previously (18). Peptides were clarified with Sep-Pak C18 cartridges (Waters Corp.) prior to mass spectrometry analysis. LC-MS/MS was carried out as described above.

Spectral Counting—
All redundant MS/MS spectra from serine hydrolases shown in Supplemental Table 2 were divided into light and heavy isotopic populations. C-terminal peptides not containing either lysine or arginine were excluded from analysis. Spectra were summed for proteins from each of the two control and two experimental samples, and averages were calculated. Protein ratios were determined by dividing the average number of heavy and light spectra for each protein followed by log2 transformation.


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
REFERENCES
 
As discussed for cleavable labeling reagents (14), smaller tags are more amenable to mass spectrometry because there are fewer potential sites of fragmentation during collision-induced dissociation, simplifying the analysis of the fragment ion spectrum. Drawing on the advantages of the newly developed catch-and-release (CAR) tags, we sought to apply this cleavable technology to the analysis of enzyme activity and to facilitate simultaneous protein identification and quantification by mass spectrometry. The use of fluorophosphonate (FP) tags to profile serine hydrolases by ABPP has been described in a number of studies and was chosen here as a proof-of-principle application. Thus, our scheme makes use of an FP catch-and-release probe (FP-CAR reagent, Fig. 1A and Supplemental Fig. 1). This reagent consists of (i) an FP group that will react with the active site nucleophile in serine hydrolases, (ii) a disulfide moiety that is selectively reducible due to steric hindrance, (iii) a biotin tag for the binding and enrichment of active hydrolases, and (iv) long linkers on each side of the cleavable site to allow for the binding and recovery of full-length proteins following avidin capture.


Figure 1
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  		FIG. 1. Structural features and MS analysis of FP-CAR reagent. A, FP-CAR reagent noting key structural and functional components such as fluorophosphonate (i), phosphine-cleavable disulfide linker with sterically hindering dimethyl and isopropyl groups (ii), biotin tag (iii), and long linkers (iv) to allow entry and flexibility during initial capture and subsequent isolation/release of full-length serine hydrolases. B, high mass accuracy MS scans of peptide DSCQGDSGGPVVCSGK from trypsin containing the active site serine residue (denoted by * in right panel). The doubly charged unlabeled peptide has a monoisotopic m/z ratio of 805.33 (left panel), whereas the peptide labeled with the cleaved reagent portion has a monoisotopic m/z ratio shifted to 1023.44 (right panel). C, assignment of b (red) and y (blue) ion series in MS/MS scan from the 1023.44 precursor ion confirms peptide identification and modified site localization. T denotes the mass of the cleaved FP-CAR reagent (436.2 Da) plus a water molecule (18.0 Da). The large m/z shift between diagnostic ions y9 and y10 due to FP-CAR derivatization shows correct labeling at the active site serine residue. Doubly charged species from y13 and y14 were observed at 842.5 and 922.6, respectively. All peak intensities under x2"umbrella" were doubled for visual clarity. Seq, sequence.

 
Although the reactivity properties of our cleavable FP are in accordance with previously published FP tags (Supplemental Fig. 2), the active site serine residue of trypsin was also identified using our cleavable reagent coupled with MS/MS. High mass accuracy MS spectra for the unmodified and FP-modified versions of the trypsin active site peptide (Fig. 1B) as well as a corroborating MS/MS fragmentation pattern (Fig. 1C) confirm both the binding of our cleavable reagent to the trypsin active site nucleophile and its compatibility with mass spectrometry.

Having established that the FP-CAR reagent behaves as expected with single proteins, we turned our attention to its use in complex mixtures from a cellular model of tumor metastasis. A general analysis of serine hydrolase activity has been explored in various models of tumorigenicity (11) but has yet to be performed in prostate cancer. The LNCaP prostate cancer line and a highly metastatic variant, LNCaP-LN3 (25), were chosen for this purpose. A genomic profile examination of prostate cancer showed that commercially available LNCaP cells cluster with clinically localized prostate cancer specimens (26), indicating that LNCaP represents a biologically relevant cellular model. LNCaP cells are also androgen-sensitive, suggesting that they represent an early stage of prostate cancer.

By coupling this labeling approach to metabolic stable isotopic labeling and mass spectrometry, we were able to obtain quantitative data for a list of proteins reflecting relative functional activity. The major advantage of this method is that using differential isotopic labeling coupled with mass spectrometry will result in more accurate quantification of enzyme activity. A schematic for comparing LNCaP and LNCaP-LN3 cells using our procedure, termed CAPE, is shown (Fig. 2A).


Figure 2
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  		FIG. 2. CAPE procedure applied to complex mixtures using a model of tumor metastasis. CAPE was used to assess differential serine hydrolase activity between low (LNCaP) and high (LNCaP-LN3) metastatic prostate cancer cells. A, CAPE combines multiple existing proteomics methods. Many steps in this schematic are common to SILAC (red), ABPP (blue), and/or CAR (green). Low and high metastatic prostate cancer cells were cultured with light and heavy amino acids, respectively. WCLs were collected, labeled with FP-CAR reagent, and mixed at a 1:1 ratio according to measured protein concentration. Serine hydrolases were captured by incubation with avidin, and nonspecific binding was reduced by washing. Protein was trypsin-digested in solution and analyzed by tandem mass spectrometry. MS and MS/MS scans provided quantitative and qualitative data, respectively. B, red and blue bars correspond to the average serine hydrolase activity levels from control (LNCaP cells cultured under both light and heavy amino acid conditions to confirm an expected 1:1 ratio) and experimental (comparing light LNCaP and heavy LNCaP-LN3 cells to determine differential serine hydrolase activity in prostate cancer metastasis) conditions, respectively, with error bars representing high/low values between biological replicates. Ratios greater than 0 represent up-regulated activity in the high metastatic cells, whereas values less than 0 correspond to decreased activity. The black dashed line shows the mean protein ratio from the control samples with red dashed lines corresponding to 3 standard deviations from the mean. Student's t test was performed on all peptides used for quantification by combining replicate experiments and comparing intraprotein variability between experimental and control peptide ratios with * representing p values <0.00001. Protein abbreviations were taken from Swiss-Prot gene names (LYPLA1, acyl-protein thioesterase 1; WBSCR21, Williams-Beuren syndrome critical region protein 21 form D; APEH, acylamino-acid-releasing enzyme; PAFAH1B3, platelet-activating factor acetylhydrolase IB {gamma} subunit; ESD, esterase D; FASN, fatty-acid synthase; also see Footnote 1 and Supplemental Table 2). Note that only proteins detected with at least two quantified peptides in each of the four experiments (control and experimental with biological replicates for each) were considered for analysis.

 
To assess which proteins exhibited significant quantitative changes, CAPE was repeated using LNCaP cells cultured under both heavy and light conditions as a control. This experiment confirmed that an expected ratio of 1:1 (no change) could be maintained throughout the labeling and purification process.

CAPE experiments led to the confident identification of 25 hydrolase proteins (Supplemental Table 1) from LNCaP and LNCaP-LN3 WCLs. Using stringent identification and quantification criteria, nine hydrolases from this list were chosen for further analysis. The hydrolase activities for these proteins displayed accurate reproducibility among biological replicates for both experimental and control conditions (Fig. 2B).

The average control log2 ratio of –0.086 with a standard deviation of 0.18 (13%) shows that the relative activities of most of the serine hydrolases remained unchanged (Supplemental Table 2). A significant change was defined as increasing or decreasing more than 3 standard deviations from the mean as determined from the average control protein ratio. Thus, log2-transformed ratios less than –0.63 or greater than 0.46 represent significant experimental changes as confirmed by Student's t test. Unexpectedly none of the enzymes showed up-regulated activity, whereas roughly half exhibited a significant decrease in activity (Fig. 2B). This suggests that a down-regulation of specific serine hydrolases may contribute to or result from the progression of metastasis in these cells.

To validate our data, we chose a number of additional control experiments to show that the labeling of enzymatic activity was specific. As with most enzymes, serine hydrolase activity is dependent on three-dimensional structure integrity. Boiling the WCLs prior to labeling with FP-CAR reagent and using the same criteria for peptide identification and quantification led to an overall reduction in activity with only one of the nine hydrolases found (fatty-acid synthase). Only a small number of peptides were identified for fatty-acid synthase in this control, indicating that this identification was likely due to minor amounts of protein refolding and reconstituted activity after sample boiling. Moreover to ensure that hydrolase identification was dependent on FP-CAR labeling and not simply due to nonspecific binding to avidin beads, the CAPE procedure was repeated with no FP-CAR reagent as a background control. None of the nine measured hydrolases were identified in this control experiment.

Although CAPE ratios showed clear reproducibility at the protein level, a quantitative analysis at the peptide level was required to determine intraprotein variability. As expected, the peptides used for protein quantification from control samples showed predominantly no change (Supplemental Fig. 3).

Optimized by Mann and co-workers (2), SILAC has been widely used to quantitatively evaluate protein levels using mass spectrometry. To assess the utility of CAPE in comparison with existing methods, the same WCLs from LNCaP and LNCaP-LN3 were mixed at a 1:1 ratio by protein concentration and subjected to SILAC analysis. The consistency of our method is shown in Supplemental Table 2 in which similar quantitative protein levels were observed between SILAC and CAPE. Due to WCL complexity, proteins analyzed by SILAC were first resolved by SDS-PAGE, fractionated 10-fold, and subjected to a longer HPLC elution gradient in an attempt to identify and quantify low abundance proteins. Supplemental Table 2 reveals that this increase in resolution failed to quantify three of the nine serine hydrolases, including two of the most significantly dysregulated enzymes, peroxisomal acyl-coenzyme A thioester hydrolase 2A (ACOT2) and sialate O-acetylesterase precursor (SIAE). Although no differences were observed in quantitative levels for serine hydrolases overlapping in both SILAC and CAPE experiments, we note that CAPE directly enriched serine hydrolases and has the potential for characterizing protein activities that may be missed by SILAC alone. Alternatively because the same metabolically labeled samples can be analyzed by both methods, thereby minimizing experimental variability, we suggest that SILAC and CAPE could be used as complementary techniques for gaining insight into post-translational mechanisms regulating protein activity in a variety of biological models.

CAPE was also evaluated against spectral counting as a means for relative quantification between low and high metastatic cells. As others have suggested, spectral counting is ideal for detecting large quantitative changes (27) and better suited to the analysis of more abundant proteins (12). However, LNCaP and LNCaP-LN3 exhibited small differences in serine hydrolase activity as assessed by CAPE. Many of these proteins have low spectral count numbers as well, suggesting they are present in comparatively low concentrations (Fig. 3). The three most dysregulated enzymes showed relatively no change as measured by spectral counting, highlighting the utility of CAPE and stable isotope labeling for detecting small quantitative changes that may have otherwise been missed by alternative MS-based methods.


Figure 3
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  		FIG. 3. Comparison of CAPE and spectral counting for quantification. All redundant spectra for each serine hydrolase were divided into light and heavy isotopic populations. A, bars represent the average number of light (solid colors) and heavy (striped colors) spectral counts obtained from the two control (red) and two experimental (blue) samples for each protein. Note that a nearly equal number of light and heavy spectra (236 and 248, respectively) were obtained from the control samples, indicating no bias toward the correct identification of light or heavy spectra. B, spectral count ratios from A were calculated and log2-transformed. Displayed are the average ratios from spectral counting from the control (open red squares) and experimental (open blue squares) samples as well as CAPE ratios for control (solid red squares) and experimental (solid blue squares) samples as shown in Fig. 2B. Ratios greater than 0 represent up-regulated activity in the high metastatic cells, whereas values less than 0 correspond to decreased activity. C, representative peptide from ABHD10 shows relatively no quantitative change in the control sample (left) as determined by CAPE, whereas the same peptide identified in the experimental sample (right) exhibits an approximate 2.5-fold decrease in activity. Control (red) and experimental (blue) monoisotopic [M + 2H]2+ peaks from light and heavy species are highlighted. LYPLA1, acyl-protein thioesterase 1; WBSCR21, Williams-Beuren syndrome critical region protein 21 form D; APEH, acylamino-acid-releasing enzyme; PAFAH1B3, platelet-activating factor acetylhydrolase IB {gamma} subunit; ESD, esterase D; FASN, fatty-acid synthase; Ctrl, control; Exp, experimental.

 
In summary, we have described a streamlined approach for identification and quantification of serine hydrolase activities using mass spectrometry with our results pointing to some potentially interesting metastatic biomarkers. Although others have suggested a tumorigenic role for decreased SIAE activity (28), ACOT2 and hypothetical protein FLJ11342 (ABHD10) exhibited markedly decreased expression in the present study but have been poorly characterized in other cancer analyses. Additional experiments are underway to determine the role these enzymes play in metastasis.

Recent work has shown the utility of combined chemical and computational approaches for identifying novel serine hydrolases (29). The use of a highly quantitative method such as CAPE in conjunction with a computational technique that pinpoints specific hydrolases could be valuable because such an approach could generate additional enzymes whose function may be important in the progression of disease.

Furthermore the CAR technology could be applied to other activity-based probes to explore functions with high specificity and sensitivity of additional enzyme classes. Considering the milieu of endogenous enzyme inhibitors and post-translational mechanisms that affect protein function, CAPE has the potential for measuring differences in protein activity in a single step that alternative mass spectrometry-based approaches such as SILAC and ICAT could not.

Addendum—
After submission of this work, two studies describing cleavable probes for functional proteomics were published (30, 31).


   ACKNOWLEDGMENTS
 
We thank C. Pettaway (M. D. Anderson Cancer Center, Houston, TX) for use of LNCaP-LN3 cell lines and M. Jedrychowski (Boston University Medical Center, Boston, MA) for critical reading of the manuscript. We also thank members of the Cravatt, Zetter, and Gygi laboratories for helpful discussions during manuscript preparations, notably N. Jessani (Scripps Research Institute), C. Bakalarski (Harvard Medical School), and S. Beausoleil (Harvard Medical School) for technical assistance.


  FOOTNOTES
 
Received, February 9, 2007, and in revised form, July 10, 2007.

Published, MCP Papers in Press, July 11, 2007, DOI 10.1074/mcp.M700057-MCP200

1 The abbreviations used are: SILAC, stable isotope labeling by amino acids in cell culture; ABPP, activity-based protein profiling; CAPE, catch-and-release activity profiling of enzymes, CAR, catch-and-release; FP, fluorophosphonate; FP-CAR, fluorophosphonate catch-and-release; ACOT2, peroxisomal acyl-coenzyme A thioester hydrolase 2A; ABHD10, hypothetical protein FLJ11342; SIAE, sialate O-acetylesterase precursor; WCL, whole-cell lysate; STAGE, stop and go extraction. Back

* This work was supported by National Institutes of Health Grants CA087660 (to B. F. C.), CA37393 (to B. R. Z.), and HG003456 (to S. P. G.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

S The on-line version of this article (available at http://www.mcponline.org) contains supplemental material. Back

|| To whom correspondence should be addressed. Tel.: 617-432-3155; Fax: 617-432-1144; E-mail: steven_gygi{at}hms.harvard.edu


   REFERENCES
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 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 

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