Targeting Proteases in Cardiovascular Diseases by Mass Spectrometry-Based Proteomics

Abstract

HomeCirculation: Cardiovascular GeneticsVol. 5, No. 2Targeting Proteases in Cardiovascular Diseases by Mass Spectrometry-Based Proteomics Free AccessResearch ArticlePDF/EPUBAboutView PDFSections ToolsAdd to favoritesDownload citationsTrack citationsPermissions ShareShare onFacebookTwitterLinked InMendeleyReddit Jump toFree AccessResearch ArticlePDF/EPUBTargeting Proteases in Cardiovascular Diseases by Mass Spectrometry-Based Proteomics Diana Klingler, PhD and Markus Hardt, PhD Diana KlinglerDiana Klingler From the Boston Biomedical Research Institute, Watertown, MA. Search for more papers by this author and Markus HardtMarkus Hardt From the Boston Biomedical Research Institute, Watertown, MA. Search for more papers by this author Originally published1 Apr 2012https://doi.org/10.1161/CIRCGENETICS.110.957811Circulation: Cardiovascular Genetics. 2012;5:265Proteases catalyze the hydrolysis of peptide bonds, which results in the cleavage of protein and peptide chains, and thereby lead to an irreversible change of protein structure. The fundamental nature of this process makes proteolysis a powerful post-translational modification that can control protein function and abundance. Digestive proteases in the gastrointestinal tract break down proteins fairly indiscriminately, while peptidases involved in cell signaling catalyze very specific cleavage reactions to regulate the abundances of bioactive peptides. Uncontrolled proteolysis could have disastrous physiological consequences; therefore, a multitude of mechanisms exist to tightly regulate proteolytic processing. One of the more basic regulatory principles is substrate specificity, in which the 3-dimensional structure of the protease determines which substrates are accessible to the active site. Other regulatory mechanisms include the activation of proteases from inactive precursors (ie, zymogens) and the limitation of protease activities to specific pH ranges and compartments (eg, lysosomal proteases). Another regulatory element is the spatial and temporal interplay of proteolytic network components; devoid of all required factors (proteases, substrates, and their respective activators and inhibitors) present, reactions may not occur (Figure 1).Download figureDownload PowerPointFigure 1. A simplified view of the essential functional relationships between members of a proteolytic network. Activators and inhibitors can modulate the activity state of proteases. For cleavage reactions to occur, the spatial and temporal distributions of activated proteases and substrates need to overlap.The renin-angiotensin system can serve as an example of how a proteolytic network regulates a physiological process:1 briefly, angiotensinogen is cleaved by renin to produce angiotensin I, which in turn is cleaved by angiotensin-converting enzyme (ACE) to form the effector peptide angiotensin II. Binding of angiotensin II to the angiotensin II receptor type 1 (AT1) mediates vasoconstriction, while binding to AT2 results in vasodilation (Figure 2). The renin-angiotensin system tightly controls arterial blood pressure and assures the uninterrupted perfusion of vital organs with oxygen and nutrients. Recent discoveries of alternative angiotensin processing pathways illustrate that the renin-angiotensin system is more complex than previously thought.2 Disregulation of the renin-angiotensin system can lead to pathological conditions such as hypertension, which is a major risk factor for congestive heart failure, stroke, and myocardial infarction. Due to their pivotal roles, proteolytic enzymes have emerged as prime targets for the pharmaceutical treatment of cardiovascular diseases.3 The increasing awareness of the modulatory effect of the biological context mandates that protease function is analyzed on a system wide level. Mass spectrometry (MS)-based proteomics provides a unique technological platform to capture the complexity and dynamic nature of proteolytic networks. Here, we review the current state of proteomics approaches used in protease biology and highlight strategies particularly suited to characterize protease activities in the context of cardiovascular disorders.Download figureDownload PowerPointFigure 2. Processing of angiotensin peptides by angiotensin-converting enzyme (ACE), ACE2, and neprilysin (NEP) as part of the renin-angiotensin system. Renin cleaves angiotensinogen to produce angiotensin I. ACE converts angiotensin I to angiotensin II. In a second processing axis, angiotensin I is cut by ACE2, resulting in angiotensin (Ang) [1–9], which is cleaved by either ACE or neprilysin to produce Ang [1–7] (bracketed numbers refer to the amino acid positions within the peptide sequences). Ang [1–7] also can result from the processing of angiotensin I by NEP or angiotensin II by ACE2. Binding of angiotensin II to the angiotensin II receptor type 1 (AT1) activates vasoconstriction. In contrast, binding to the AT2 receptor mediates vasodilation, which also can be initiated by the binding of Ang [1–7] to the Mas receptor.Mass Spectrometry-Based Methods to Unravel Proteolytic NetworksThe renin-angiotensin system demonstrates how MS-based proteomics can help to fill the need for a detailed understanding of proteolytic networks. While renin and ACE were the original proteases of the renin-angiotensin system, the updated view includes additional proteases such as ACE2, chymase, neprilysin, and aminopeptidases (A and N).4–7 Likewise, additional bioactive peptides, such as the angiotensin (Ang) metabolites Ang III [2–8]), Ang IV [3–8], and Ang [1–7], which were previously considered functionally inactive, are now included.8 MS-based technologies played a leading role in the identification of these novel system constituents.Historically, protease function is assessed on an individual basis by in vitro enzyme assays after biochemical purification. The advent of new analytic technologies makes it now feasible to investigate protease activity and function in complex environments.9–13 Generally, these approaches can be categorized based on whether they are designed to (1) identify constituents of the proteolytic network (proteases, substrates), (2) screen for protease inhibitors/activators/modulators, or (3) characterize the dynamics of proteolytic processing. In the Table we provide an overview of the most commonly applied strategies in the field and which aspects of proteolysis they address. To choose the appropriate method, it is essential to define what aspect of protease biology needs to be addressed. Figure 1 depicts the functional relationships between members of proteolytic networks and can serve as a guide to develop research strategies. Is the protease of interest catalytically active? How does the zymogen differ from the active form of the protease? Which molecules can modulate the activity and substrate specificity of the protease? What are the endogenous substrates of the protease? What are the spatial and temporal distributions of the protease and other components of the proteolytic network? Many of these questions can be answered readily by online resources; the MEROPS database (http://merops.sanger.ac.uk) curates published information about peptidases, their substrates, and inhibitors, and offers indications about overall substrate specificity;14 the Proteolysis MAP (PMAP) (http://www.proteolysis.org) combines 5 databases (ProteaseDB, SubstrateDB, CutDB, ProfileDB, and PathwayDB) and a computational toolkit (including cleavage site predictions) to create an integrated reasoning environment to analyze proteolytic networks.15Table. Commonly Applied Techniques to Study Proteolytic ReactionsMethodDescriptionApplicationProteaseSubstrateInhibitorProcessGeneral techniques for protease research Yeast 2-hybrid systemsYeast libraries carrying cloned ORF*Identification of protein interactions on a large scale•••Sequencing of positive PCR amplification productsTwo target ORFs are analyzed per experiment Phage-displayAffinity selection of clones and screening of peptide libraries†High throughput screening of protein interactions and inhibitors•••DNA sequencing of remaining amplification productsOne target protein or peptide is analyzed per experiment 2D DIGEGel-based protein separation of fluorescent labeled samples‡Quantitation of proteins and post-translational modifications••Proteins are imaged by fluorescence and identified by MSComparison of 2 samples with internal standards MudPITPeptide products are separated by SCX and RP-HPLC§Identification of peptide products by MS••Comparison of an unlimited No. of samples ICATLabeling of cysteine-residues in proteins followed by digestion∥Relative quantitation of peptide products by MS•Comparison of 2 samples iTRAQIsobaric amine-specific tagging of peptide products**Relative and absolute quantitation of protease activity by MS•Comparison of up to 8 samples SILACExpressed proteases/peptide products labeled by amino acids††Identification and quantitation of peptide products by MS•Comparison of an unlimited No. of samples MRMMultiple reaction monitoring of known peptide fragments by targeted MS‡‡Absolute quantitation of known peptides••Comparison of an unlimited No. of samplesTechniques specifically designed for protease research CLiPSCombinatorial approach to measure substrate hydrolysis§§Identification of substrates by quantitative screening of whole-cell fluorescence•• Positional scanning synthetic librariesScreening of tetra-peptide libraries by proteolysis-dependent signal intensities∥∥Screen for P1–P4 substrate specificities• Colloidal barcoding bead-based protease profilingScreening of combinatorial libraries using polyelectrolyte-coated fluorescent silica reporter particles***Identification of consensus proteolytic cleavage sites•••• NIR fluorogenic reportersNIR fluorescence signal upon cleavage of protease-sensitive peptide linkers†††In vivo imaging and quantitation of protease activities•••• C- and N-term enrichment of cleavage productsNegative or positive enrichment of C- or N-terminal peptide cleavage products‡‡‡Identification of proteolytic peptides and cleavage sites•Comparison of 2 samples ABPsChemical probes with affinity and fluorescent tags§§§Report on the structure and reactivity of enzyme active sites in cells and tissues••• PALeO18O-labeling of proteolytic peptides during hydrolysis∥∥∥Quantitation and identification of protease activity, peptide substrates and cleavage products••••Comparison of an unlimited No. of samplesORF indicates open reading frames; PCR, polymerase chain reaction; SCX, strong cation exchange; RP-HPLC, reversed phase high-performance liquid chromatography; DIGE, differential gel electrophoresis; MS, mass spectrometry; MudPIT, multidimensional protein identification technique; ICAT, isotope-coded affinity tags; iTRAQ, isobaric tags for relative and absolute quantitation; SILAC, stable isotope labeling by amino acids; MRM, multiple reaction monitoring; CLiPS, cellular libraries of peptide substrates; NIR, near infrared; ABP, activity-based probes; PALeO, protease activity labeling employing 18O-enriched water.*Data from Parrish JR, Gulyas KD, Finley RL Jr. Yeast two-hybrid contributions to interactome mapping. Curr Opin Biotechnol. 2006;17:387–393.†Data from Balestrieri ML, Napoli C. Novel challenges in exploring peptide ligands and corresponding tissue-specific endothelial receptors. Eur J Cancer. 2007;43:1242–1250.‡Data from Unlu M, Morgan ME, Minden JS. Difference gel electrophoresis: a single gel method for detecting changes in protein extracts. Electrophoresis. 1997;18:2071–2077.§Data from Link AJ, Eng J, Schieltz DM, Carmack E, Mize GJ, Morris DR, Garvik BM, Yates JR III. Direct analysis of protein complexes using mass spectrometry. Nat Biotechnol. 1999;17:676–682.∥Data from Gygi SP, Rist B, Gerber SA, Turecek F, Gelb MH, Aebersold R. Quantitative analysis of complex protein mixtures using isotope-coded affinity tags. Nat Biotechnol. 1999;17:994–999.**Data from Ross PL, Huang YN, Marchese JN, Williamson B, Parker K, Hattan S, Khainovski N, Pillai S, Dey S, Daniels S, Purkayastha S, Juhasz P, Martin S, Bartlet-Jones M, He F, Jacobson A, Pappin DJ. Multiplexed protein quantitation in Saccharomyces cerevisiae using amine-reactive isobaric tagging reagents. Mol Cell Proteomics. 2004;3:1154–1169.††Data from Ong SE, Blagoev B, Kratchmarova I, Kristensen DB, Steen H, Pandey A, Mann M. Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol Cell Proteomics. 2002;1:376–386.‡‡Data from Picotti P, Rinner O, Stallmach R, Dautel F, Farrah T, Domon B, Wenschuh H, Aebersold R. High-throughput generation of selected reaction-monitoring assays for proteins and proteomes. Nat Methods. 2010;7:43–46.§§Data from Boulware KT, Daugherty PS. Protease specificity determination by using cellular libraries of peptide substrates (CLiPS). Proc Natl Acad Sci U S A. 2006;103:7583–7588.∥∥Data from Barrios AM, Craik CS. Scanning the prime-site substrate specificity of proteolytic enzymes: a novel assay based on ligand-enhanced lanthanide ion fluorescence. Bioorg Med Chem Lett. 2002;12:3619–3623.***Data from Marcon L, Battersby BJ, Rühmann A, Ford K, Daley M, Lawrie GA, Trau M. ‘On-the-fly' optical encoding of combinatorial peptide libraries for profiling of protease specificity. Mol Biosyst. 2010;6:225–233.†††Data from Baruch A, Jeffery DA, Bogyo M. Enzyme activity–it's all about image. Trends Cell Biol. 2004;14:29–35.‡‡‡Data from Gevaert K, Goethals M, Martens L, van Damme J, Staes A, Thomas GR, Vandekerckhove J. Exploring proteomes and analyzing protein processing by mass spectrometric identification of sorted N-terminal peptides. Nat Biotech. 2003;21:566–569.§§§Data from Cravatt BF, Wright A, Kozarich J. Activity-based protein profiling: from enzyme chemistry to proteomic chemistry. Annu Rev Biochem. 2008;77:383–414.∥∥∥Data from Robinson S, Niles RK, Witkowska HE, Rittenbach KJ, Nichols RJ, Sargent JA, Dixon SE, Prakobphol A, Hall SC, Fisher SJ, Hardt M. A mass spectrometry-based strategy for detecting and characterizing endogenous proteinase activities in complex biological samples. Proteomics. 2008;8:435–445.MS-based proteomics has emerged as the premier tool to identify and quantify proteins and peptides.16–18 Briefly, in a typical proteomics identification workflow, proteins are separated by 1- or 2-dimensional gel electrophoresis, digested by exogenous proteases (ie, trypsin). Resulting peptide fragments are recovered, fractionated by reverse-phase chromatography, and their molecular masses measured by MS. Selected peptides are fragmented in tandem MS experiments, and resulting fragmentation data is submitted to search engines (ie, Mascot) that match it to fragmentation patterns predicted from protein databases.19 Alternatively, in a peptidomics workflow the digestion step is omitted and peptide products formed by endogenous proteases are directly isolated.20 Peptide sequence assignments particularly are challenging in peptidomics-type experiments; however, recent advances in bioinformatics are starting to approach this problem.21How to Develop a Quantitative View of Proteolytic ProcessingWhile MS readily provides protein and peptide identities, MS measurements are inherently poorly quantitative.22 Accurate quantitation of protein and peptide expression levels is a prerequisite to capture the dynamic nature of proteolysis and to gain functional insights. Quantitation by MS is achieved by either label-free or stable isotope labeling methods. Measurements of chromatographic ion intensity (eg, peak areas) and spectral counting are the most commonly applied label-free approaches.23 Label-free methods provide relative quantitation for an unlimited number of samples. Stable isotope labeling strategies, in contrast, can provide relative and absolute quantitation; however, the specifics of the labeling reactions can limit the number of samples interrogated. Over recent years, a broad variety of stable isotope labeling methods have been developed.23,24 Here, we will focus on the most prominent examples that have been used in the context of protease biology. In general, the presented techniques are capable of simultaneously identifying and quantifying novel and known proteases and their substrates. They also can be used to test the efficacy of protease inhibitors.Isotope-coded affinity tags (ICAT) are one of the best-known techniques to measure protein abundances. ICAT reagents are comprised of 3 functional components: (1) a reactive group specific toward cysteinyl residues, (2) a stable isotope label, and (3) a biotin affinity tag.25 The affinity tag allows for the selective enrichment of cysteine-containing peptides, thus reducing sample complexity. The stable isotope label introduces either a light or heavy tag, which in turn allows for the comparative analysis of protein expression levels across 2 states. After labeling with ICAT reagents, 2 samples are combined and digested, and cysteine-containing peptides are enriched by avidin affinity chromatography. Liquid chromatography (LC)-MS/MS analysis yields both protein quantity and identity. The ICAT approach has been used successfully to identify novel protease substrates in complex cellular environments and quantify protease activity.26 The exclusive reliance on cysteine-containing peptides limits the applicability of the ICAT approach as a general quantitation approach.This shortcoming has been addressed by the next generation of chemical labeling strategies that tag peptide N-termini and lysine side chains using N-hydroxy-succinimide chemistry. The tandem mass tag27 and iTRAQ (isobaric tags for relative and absolute quantitation)28 strategies share as an important design feature an isobaric stable isotope moiety, which renders differentially labeled samples indistinguishable during chromatographic and mass spectrometric analyses. Only on MS/MS fragmentation are low molecular weight reporter ions released, and their relative ion abundances can be used for quantitation. Currently, there are f4 and 8 reporter ions available for iTRAQ28,29 and 2 and 6 for tandem mass tag,27,30 each allowing for multiplexed analysis in single LC-MS/MS experiments. iTRAQ labeling has been applied successfully to quantify peptides generated by endogenous proteases31 and to monitor protease activity in cell cultures.32SILAC (stable isotope labeling by amino acids) has been developed as a metabolic labeling method alternative to chemical labeling approaches. SILAC has been applied successfully in cell culture systems33 and more recently in entire organisms.34 SILAC relies on the in vitro incorporation of essential amino acids with substituted stable isotope nuclei (eg, Arg and Lys labeled with 13C, 15N). Heavy and light versions of the amino acids are incorporated into every protein in the system, which, in turn, allows for comparative expression analyses. SILAC has been used to determine which substrates are trapped by a catalytically inactivated protease,35 and more recently to quantify Granzyme B catalyzed proteolysis.36 Compared with chemical derivatization strategies, SILAC offers more efficient and reproducible labeling.High levels of sample complexity and large cleavage products are a challenge for proteomic analyses. To reduce complexity and determine cleavage sites within large proteins, investigators developed strategies to enrich for N-terminal cleavage products that are of interest.37 In positive enrichment methods, α-amino groups that are formed newly by proteolysis are captured by chemical or enzymatic processes.38,39 In depletion methods, internal peptides are removed by either altered chromatographic properties40 or by chemical means.41 Recently, a positive enrichment strategy termed C terminomics has been introduced for the until then elusive C-terminal cleavage products.42 Combining N- and C-terminal peptide data provides complementary information and increased coverage of cleavage sites. However, such enrichment steps considerably extend sample processing and analysis time.Another strategy to address sample complexity is targeted MS. These types of experiments typically are performed on triple quadrupole instruments operating in multiple reaction monitoring (MRM) mode. MRM assays determine the abundances of peptide analytes by measuring the intensities of their characteristic MS/MS fragment ions and can be used in relative and absolute quantitation mode. Compared with other MS approaches, MRM provides higher sensitivity and specificity.17 MRM assays have been used to study the pharmacokinetics of enalapril, an ACE inhibitor used in the treatment of hypertension and congestive heart failure.43 The MRM approach also was used to simultaneously quantify 7 angiotensin II receptor antagonists in plasma of patients undergoing cardiovascular treatment.44 Despite their broad usage in pharmacological studies, MRM assays so far have not been employed in protease research, which can be attributed partially to the laborious assay development that requires optimized LC and MS conditions for each analyte.45How to Determine the Substrate Specificity and Activity State of ProteasesProteases are typically low abundant molecules, which makes their functional characterization by MS-based workflows challenging. Therefore, activity-based probes (ABPs) have been devised as chemical warheads directed at the active sites of enzymes. ABP-based profiling has been used to detect and capture the subset of catalytically active proteases in complex biological systems and to give insights into the catalytic mechanism and substrate specificity of enzymes.46 ABPs report on the functional state of proteases by selectively labeling enzymes that share active site features (affinity, reactivity). Currently, ABPs have been developed for serine-, cysteine-, aspartyl-, and metallo-proteases.47 One of the most important design features of ABPs is their ability to distinguish active proteases from inactive precursors (zymogens)48 or proteases whose activity is modulated by other mechanisms.49 ABP profiling thereby provides functional information that is beyond standard protein expression data. ABPs with various combinations of affinity and fluorescent tags have been developed to study diseases in vivo.50,51 In conjunction with downstream analytic technologies like MS, ABP profiling offers the unique advantage to enrich for proteome fractions that share functional properties. ABP profiling generally can be classified between (1) gel-based methods and (2) LC-MS strategies.52,53 The LC-MS based platform uses biotin-tagged probes that capture intact enzymes on streptavidin beads. On-bead digestion and subsequent LC-MS analysis provide protein identification.54 In the bottom-up variation of the strategy, the enrichment step occurs on the peptide level after all proteins have been digested.55 Shortcomings of ABP profiling include the limited specificity and availability of functional probes. Also, some of the probes are toxic and cannot be used in vivo. Finally, ABPs inactivate captured proteases, and therefore enzyme function cannot be further studied.How to Gain Insights Into the Dynamics of Proteolytic ProcessingIn contrast to the methods described above that primarily aim at the identification of protease network components, few experimental approaches exist that capture the dynamics of proteolytic processing.56 Classically, chromo- or fluorogenic protease cleavage assays are used to measure the kinetics of individual proteolytic reactions. Fluorescence-based assays rely on the activation or dequenching of fluorophores on proteolytic cleavage. The design of such assays requires precise knowledge of the targeted cleavage sites, so that appropriate constructs can be synthesized that specifically report substrate-to-product conversions.57 Introduction of fluorescent reporter functionalities may alter substrate-enzyme interaction kinetics. Also, cleavage assays are likely to fall short when complex proteolytic reactions are being investigated, that is, when a protease recognizes multiple, alternate cleavage sites on the same substrate molecule. MS can fill this critical technological gap by its ability to detect and identify unknown peptide metabolites on a large-scale in a virtually unbiased way. MS-based monitoring of enzymatic reactions has been described for electrospray and matrix-assisted laser desorption ionization (MALDI)-ionization techniques.58 For example, in the MALDI-MES (MS-assisted enzyme screening) strategy,59 endogenous proteases are captured from crude samples and immobilized on functionalized beads. The beads subsequently are incubated with substrate solutions, whose compositions are monitored over time by MALDI-MS analyses. Enzyme immobilization and the use of separate substrate solutions minimize interferences from the biological matrix (eg, salt, proteins) during MS-analysis. Villanueva et al60 showed that even against complex biological backgrounds (ie, serum), ex vivo incubation of endogenous proteases and substrates can yield characteristic protease activity signatures that can be used as cancer biomarkers. In the PALeO approach (protease activity labeling employing 18O-enriched water),61 ex vivo incubations occur in the presence of 18O-enriched water. Hydrolysis of peptide bonds results in the concomitant incorporation of solvent 18O-atoms into the C-termini of nascent cleavage products, which can be detected readily by MS based on their characteristic isotope patterns.62 Interestingly, some substrate-protease interactions extend beyond the initial cleavage reaction; cleavage products formed by serine proteases (eg, trypsin) have been shown to rebind to the protease and reform acyl-enzyme intermediates. In the presence of H218O, hydrolysis of the acyl-enzyme intermediate facilitates the incorporation of a second 18O-atom into the C-terminus.6318O-based strategies such as PALeO, therefore, can provide additional information regarding aspects of protease-substrate interaction and enzymatic mechanism. The 18O-label allows positively selecting cleavage products and disregarding signals derived from the biological matrix background. Strategies such as PALeO are important steps toward mapping proteolytic networks in more global and dynamic ways. Studying complex proteolytic networks, however, remains an analytic challenge, and accurate quantitative information is needed. Such efforts can be aided by providing nondegradable quantitation standards (eg, all d-amino acid peptides) and defined protease substrates.64 Stable isotope labeling of these peptides (eg, by acid-catalyzed 18O-exchange) allows one to track their degradation in complex mixtures.65The emerging field of dynamic proteomics promises to greatly facilitate the elucidation of complex proteolytic networks. Already there is ample evidence that such approaches can shed new light into even well studied subject areas such as the renin-angiotensin system. Using MALDI-MES, Schlüter et al66 determined that renal Cathepsin G can generate angiotensin II. Using the PALeO assay, we demonstrated that endothelin-converting enzyme-1 (ECE-1), a member of the neprilysin protease family, also can generate angiotensin II (Figure 3; M. Hardt, unpublished data, 2011). ECE-1-mediated peptide processing included additional cleavage reactions that could only be resolved by temporal analysis.67 Further, the data reconfirmed that pH conditions influence the substrate specificity and activity profile of ECE-1.68 The existence of alternative angiotensin II biosynthesis pathways illustrates that the renin-angiotensin system is far more complex than previously thought.Download figureDownload PowerPointFigure 3. Protease activity labeling employing 18O-enriched water (PALeO), a dynamic MS-based protease assay, shows that endothelin-converting enzyme-1 (ECE-1) converts angiotensin I to multiple biologically active metabolites, including angiotensin II. The waterfall plot shows the subsequent generation of angiotensin (Ang) [1–9], angiotensin II, and Ang [1–7] at extracellular pH (7.4). Red letters in the insert specify sites of proteolytic 18O-incorporation. Blue arrows indicate novel cleavage sites detected by PALeO, while the black arrow denotes the previously known cut site.63Future PerspectivesProteomics and peptidomics technologies continue to positively impact protease research by aiding in the identification and quantitation of proteases, their substrates, and inhibitors. Advances in MS instrument technology that improve mass accuracy and sensitivity will further increase identification rates.69 Advances in MS-quantitation methods will help shift the analytic focus from static to dynamic measurements and assist in moving the field from identification- to activity-based workflows. In addition, improved quantitation techniques will enhance comparative studies that, for example, investigate the effect of perturbations (eg, pathological conditions) on proteolytic networks.Due to their key roles in diseases, proteases have tremendous potential as therapeutic targets and biomarkers. Taking advantage of the catalytic properties could lead to more specific and sensitive diagnostic tools compared with what is achievable with noncatalytic biomarkers. Proteases such as ACE are prime examples for how proteolytic enzymes have been targeted in drug discovery.3 MS provides valuable tools to these endeavors. However, it is important to keep in mind that a single analytic platform is unlikely to explain complex and dynamic proteolytic systems by themselves. In vivo validation, such as classical overexpression and knock out experiments, are needed to confirm the biochemical roles of proteases. Novel live imaging probes are changing the validation process. Near infrared