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Snakebite is a major medical concern in many parts of the world with metalloproteases playing important roles in the pathological effects of Viperidae venoms, including local tissue damage, hemorrhage, and coagulopathy. Hemorrhagic Factor 3 (HF3), a metalloprotease from

With 2129 proteins still classified by the Human Proteome Organisation Human Proteome Project (HPP) as "missing" without compelling evidence of protein existence (PE) in humans, we hypothesized that in-depth proteomic characterization of tissues that are technically challenging to access and extract would yield evidence for tissue-specific missing proteins. Paradoxically, although the skeleton is the most massive tissue system in humans, as one of the poorest characterized by proteomics, bone falls under the HPP umbrella term as a "rare tissue". Therefore, we aimed to optimize mineralized tissue protein extraction methodology and workflows for proteomic and data analyses of small quantities of healthy young adult human alveolar bone. Osteoid was solubilized by GuHCl extraction, with hydroxyapatite-bound proteins then released by ethylenediaminetetraacetic acid demineralization. A subsequent GuHCl solubilization extraction was followed by solid-phase digestion of the remaining insoluble cross-linked protein using trypsin and then 6 M urea dissolution incorporating LysC digestion. Bone extracts were digested in parallel using trypsin, LysargiNase, AspN, or GluC prior to liquid chromatography-mass spectrometry analysis. Terminal Amine Isotopic Labeling of Substrates was used to purify semitryptic peptides, identifying natural and proteolytic-cleaved neo N-termini of bone proteins. Our strategy enabled complete solubilization of the organic bone matrix leading to extensive categorization of bone proteins in different bone matrix extracts, and hence matrix compartments, for the first time. Moreover, this led to the high confidence identification of pannexin-3, a "missing protein", found only in the insoluble collagenous matrix and revealed for the first time by trypsin solid-phase digestion. We also found a singleton proteotypic peptide of another missing protein, meiosis inhibitor protein 1. We also identified 17 proteins classified in neXtprot as PE1 based on evidence other than from MS, termed non-MS PE1 proteins, including ≥9-mer proteotypic peptides of four proteins.

Gluten, a group of proteins found in wheat, barley, and rye, is the trigger of celiac disease, an immune disorder that affects about 1% of people worldwide. The toxicity of partially hydrolyzed gluten (PHG) in fermented products is less well understood due to the significant analytical challenges in PHG characterization. In this project, an N-terminal labeling mass spectrometry method, terminal amine isotopic labeling of substrates (TAILS), was optimized for the in-depth analysis of PHG and validated using a test protease (trypsin) with known cleavage specificity. Gluten N-termini in test and control groups were labeled with heavy and light formaldehyde, respectively. Trypsin-generated neo N-termini were identified by exhibiting an MS1 Log2 H:L peak area ratio with a significant difference (p .01) from zero and native N-termini with no significant difference from zero (p .01). Using this strategy, all abundant, theoretical, test protease-generated peptides in exemplar alpha/beta gliadins and gamma gliadins were identified. SIGNIFICANCE: This study is the first study that modified and evaluated TAILS analysis for the analysis of partially hydrolyzed gluten proteins. The evaluation indicated that the TAILS analysis could be modified and expanded to the identification of multiple protease cleavage sites in gluten proteins and is worth further evaluation as a novel strategy for the analysis of natural hydrolysis of gluten in food processes. This strategy also may be further applied to characterize a broader range of partially hydrolyzed allergens in foods and provide reference for their safety assessment to both industry and regulatory authorities.

The comprehensive profiling of the repertoire of secreted proteins from cancer cells samples provides information on the signaling events in oncogenesis as well as on the cross-talk between normal and tumoral cells. Moreover, the analysis of post-translational modifications in secreted proteins may unravel biological circuits regulated by irreversible modifications such as proteolytic processing. In this context, we used Terminal Amine Isotopic Labeling of Substrates (TAILS) to perform a system-wide investigation on the N-terminome of the secretomes derived from a paired set of mouse cell lines: Melan-a (a normal melanocyte) and Tm1 (its transformed phenotype). Evaluation of the amino acid identities at the scissile bond in internal peptides revealed significant differences, suggesting distinct proteolytic processes acting in the normal and tumoral secretomes. The mapping and annotation of cleavage sites in the tumoral secretome suggested functional roles of active proteases in central biological processes related to oncogenesis, such as the processing of growth factors, cleavage of extracellular matrix proteins and the shedding of ectopic domains from the cell surface, some of which may represent novel processed forms of the corresponding proteins. In the context of the tumor microenvironment, these results suggest important biological roles of proteolytic processing in murine melanoma secreted proteins.

Enteroviruses encode proteinases that are essential for processing of the translated viral polyprotein. In addition, viral proteinases also target host proteins to manipulate cellular processes and evade innate antiviral responses to promote replication and infection. Although some host protein substrates of enterovirus proteinases have been identified, the full repertoire of targets remains unknown. We used a novel quantitative in vitro proteomics-based approach, termed t erminal a mine i sotopic l abeling of s ubstrates (TAILS), to identify with high confidence 72 and 34 new host protein targets of poliovirus and coxsackievirus B3 (CVB3) 3C proteinases (3C pro s) in HeLa cell and cardiomyocyte HL-1 cell lysates, respectively. We validated a subset of candidate substrates that are targets of poliovirus 3C pro in vitro including three common protein targets, phosphoribosylformylglycinamidine synthetase (PFAS), hnRNP K, and hnRNP M, of both proteinases. 3C pro -targeted substrates were also cleaved in virus-infected cells but not noncleavable mutant proteins designed from the TAILS-identified cleavage sites. Knockdown of TAILS-identified target proteins modulated infection both negatively and positively, suggesting that cleavage by 3C pro promotes infection. Indeed, expression of a cleavage-resistant mutant form of the endoplasmic reticulum (ER)-Golgi vesicle-tethering protein p115 decreased viral replication and yield. As the first comprehensive study to identify and validate functional enterovirus 3C pro substrates in vivo , we conclude that N-terminomics by TAILS is an effective strategy to identify host targets of viral proteinases in a nonbiased manner. IMPORTANCE Enteroviruses are positive-strand RNA viruses that encode proteases that cleave the viral polyprotein into the individual mature viral proteins. In addition, viral proteases target host proteins in order to modulate cellular pathways and block antiviral responses in order to facilitate virus infection. Although several host protein targets have been identified, the entire list of proteins that are targeted is not known. In this study, we used a novel unbiased proteomics approach to identify ∼100 novel host targets of the enterovirus 3C protease, thus providing further insights into the network of cellular pathways that are modulated to promote virus infection.

GREDIL for quantitative N-glycomics largely improves the stability of N-glycan 18O-labeling and greatly decreases the interference of isotopic cluster overlap.

Proteolysis is an irreversible post-translational modification that affects intra- and intercellular communication by modulating the activity of bioactive mediators. Key to understanding protease function is the system-wide identification of cleavage events and their dynamics in physiological contexts. Despite recent advances in mass spectrometry-based proteomics for high-throughput substrate screening, current approaches suffer from high false positive rates and only capture single states of protease activity. Here, we present a workflow based on multiplexed terminal amine isotopic labeling of substrates for time-resolved substrate degradomics in complex proteomes. This approach significantly enhances confidence in substrate identification and categorizes cleavage events by specificity and structural accessibility of the cleavage site. We demonstrate concomitant quantification of cleavage site spanning peptides and neo-N and/or neo-C termini to estimate relative ratios of noncleaved and cleaved forms of substrate proteins. By applying this strategy to dissect the matrix metalloproteinase 10 (MMP10) substrate degradome in fibroblast secretomes, we identified the extracellular matrix protein ADAMTS-like protein 1 (ADAMTSL1) as a direct MMP10 substrate and revealed MMP10-dependent ectodomain shedding of platelet-derived growth factor receptor alpha (PDGFRα) as well as sequential processing of type I collagen. The data have been deposited to the ProteomeXchange Consortium with identifier PXD000503.

Background: Biological roles for intracellular dipeptidyl peptidases 8 and 9 are unknown. Results: By degradomics, 29 new in vivo substrates were identified (nine validated) for DP8/DP9, including adenylate kinase 2 and calreticulin. Conclusion: These substrates indicate roles for DP8 and DP9 in metabolism and energy homeostasis. Significance: Being the first proteomics screen for DP8/DP9 substrates, unexpected new cellular roles were revealed., Dipeptidyl peptidases (DP) 8 and 9 are homologous, cytoplasmic N-terminal post-proline-cleaving enzymes that are anti-targets for the development of DP4 (DPPIV/CD26) inhibitors for treating type II diabetes. To date, DP8 and DP9 have been implicated in immune responses and cancer biology, but their pathophysiological functions and substrate repertoire remain unknown. This study utilizes terminal amine isotopic labeling of substrates (TAILS), an N-terminal positional proteomic approach, for the discovery of in vivo DP8 and DP9 substrates. In vivo roles for DP8 and DP9 in cellular metabolism and homeostasis were revealed via the identification of more than 29 candidate natural substrates and pathways affected by DP8/DP9 overexpression. Cleavage of 14 substrates was investigated in vitro; 9/14 substrates for both DP8 and DP9 were confirmed by MALDI-TOF MS, including two of high confidence, calreticulin and adenylate kinase 2. Adenylate kinase 2 plays key roles in cellular energy and nucleotide homeostasis. These results demonstrate remarkable in vivo substrate overlap between DP8/DP9, suggesting compensatory roles for these enzymes. This work provides the first global investigation into DP8 and DP9 substrates, providing a number of leads for future investigations into the biological roles and significance of DP8 and DP9 in human health and disease.

Stable isotope labeling is at present one of the most powerful methods in quantitative proteomics. Stable isotope labeling has been performed at both the protein as well as the peptide level using either metabolic or chemical labeling. Here, we present a straightforward and cost-effective triplex quantification method that is based on stable isotope dimethyl labeling at the peptide level. Herein, all proteolytic peptides are chemically labeled at their alpha- and epsilon-amino groups. We use three different isotopomers of formaldehyde to enable the parallel analysis of three different samples. These labels provide a minimum of 4 Da mass difference between peaks in the generated peptide triplets. The method was evaluated based on the quantitative analysis of a cell lysate, using a typical "shotgun" proteomics experiment. While peptide complexity was increased by introducing three labels, still more than 1300 proteins could be identified using 60 microg of starting material, whereby more than 600 proteins could be quantified using at least four peptides per protein. The triplex labeling was further utilized to distinguish specific from aspecific cAMP binding proteins in a chemical proteomics experiment using immobilized cAMP. Thereby, differences in abundance ratio of more than two orders of magnitude could be quantified.

Quantitation of protein abundance is a vital component in the proteomic analysis of biological systems, which can be achieved by differential stable isotopic labeling. To analyze tissue-derived samples, the isotopic labeling can be performed using chemical labeling of the peptides post-digestion. Standard chemical labeling procedures often require many manual sample handling steps, reducing the accuracy of measurements. Here, we describe a fully automated, online (in nanoLC columns), labeling procedure, which allows protein quantitation using differential isotopic dimethyl labeling of peptide N termini and lysine residues. We show that the method allows reliable quantitation over a wide dynamic range and can be used to quantify differential protein abundances in lysates and, more targeted, differences in composition between purified protein complexes. We apply the method to determine the differences in composition between bovine liver and spleen 20 S core proteasome complexes. We find that although all catalytically active immunoproteasome subunits were up-regulated in spleen (compared with liver), only one of the normal catalytic subunits was down-regulated, suggesting that the tissue-specific immunoproteasome assembly is more diverse than previously assumed.

Quantification by series of b, y fragment ion pairs generated from isobaric-labeled peptides in MS2 spectra has recently been considered an accurate strategy in quantitative proteomics. Here we developed a novel MS2 quantification approach named quantitation by isobaric terminal labeling (QITL) by coupling (18)O labeling with dimethylation. Trypsin-digested peptides were labeled with two (16)O or (18)O atoms at their C-termini in H(2)(16)O or H(2)(18)O. After blocking all ε-amino groups of lysines through guanidination, the N-termini of the peptides were accordingly labeled with formaldehyde-d(2) or formaldehyde. These indistinguishable, isobaric-labeled peptides in MS1 spectra produce b, y fragment ion pairs in the whole mass range of MS2 spectra that can be used for quantification. In this study, the feasibility of QITL was first demonstrated using standard proteins. An accurate and reproducible quantification over a wide dynamic range was achieved. Then, complex rat liver samples were used to verify the applicability of QITL for large-scale quantitative analysis. Finally, QITL was applied to profile the quantitative proteome of hepatocellular carcinoma (HCC) and adjacent non-tumor liver tissues. Given its simplicity, low-cost, and accuracy, QITL can be widely applied in biological samples (cell lines, tissues, and body fluids, etc.) for quantitative proteomic research.

In the quantitative proteomic studies, numerous in vitro and in vivo peptide labeling strategies have been successfully applied to measure differentially regulated protein and peptide abundance. These approaches have been proven to be versatile and repeatable in biological discoveries. (15)N metabolic labeling is one of these widely adopted and economical methods. However, due to the differential incorporation rates of (15)N or (14)N, the labeling results produce imperfectly matched isotopic envelopes between the heavy and light nitrogen-labeled peptides. In the present study, we have modified the solid Arabidopsis growth medium to standardize the (15)N supply, which led to a uniform incorporation of (15)N into the whole plant protein complement. The incorporation rate (97.43±0.11%) of (15)N into (15)N-coded peptides was determined by correlating the intensities of peptide ions with the labeling efficiencies according to Gaussian distribution. The resulting actual incorporation rate (97.44%) and natural abundance of (15)N/(14)N-coded peptides are used to re-calculate the intensities of isotopic envelopes of differentially labeled peptides, respectively. A modified (15)N/(14)N stable isotope labeling strategy, SILIA, is assessed and the results demonstrate that this approach is able to differentiate the fold change in protein abundance down to 10%. The machine dynamic range limitation and purification step will make the precursor ion ratio deriving from the actual ratio fold change. It is suggested that the differentially mixed (15)N-coded and (14)N-coded plant protein samples that are used to establish the protein abundance standard curve should be prepared following a similar protein isolation protocol used to isolate the proteins to be quantitated.

Terminal amine isotopic labeling of substrates (TAILS), our recently introduced platform for quantitative N-terminome analysis, enables wide dynamic range identification of original mature protein N-termini and protease cleavage products. Modifying TAILS by use of isobaric tag for relative and absolute quantification (iTRAQ)-like labels for quantification together with a robust statistical classifier derived from experimental protease cleavage data, we report reliable and statistically valid identification of proteolytic events in complex biological systems in MS2 mode. The statistical classifier is supported by a novel parameter evaluating ion intensity-dependent quantification confidences of single peptide quantifications, the quantification confidence factor (QCF). Furthermore, the isoform assignment score (IAS) is introduced, a new scoring system for the evaluation of single peptide-to-protein assignments based on high confidence protein identifications in the same sample prior to negative selection enrichment of N-terminal peptides. By these approaches, we identified and validated, in addition to known substrates, low abundance novel bioactive MMP-2 targets including the plasminogen receptor S100A10 (p11) and the proinflammatory cytokine proEMAP/p43 that were previously undescribed.

Proteolysis is a major protein posttranslational modification that, by altering protein structure, affects protein function and, by truncating the protein sequence, alters peptide signatures of proteins analyzed by proteomics. To identify such modified and shortened protease-generated neo-N-termini on a proteome-wide basis, we developed a whole protein isobaric tag for relative and absolute quantitation (iTRAQ) labeling method that simultaneously labels and blocks all primary amines including protein N- termini and lysine side chains. Blocking lysines limits trypsin cleavage to arginine, which effectively elongates the proteolytically truncated peptides for improved MS/MS analysis and peptide identification. Incorporating iTRAQ whole protein labeling with terminal amine isotopic labeling of substrates (iTRAQ-TAILS) to enrich the N-terminome by negative selection of the blocked mature original N-termini and neo-N-termini has many advantages. It enables simultaneous characterization of the natural N-termini of proteins, their N-terminal modifications, and proteolysis product and cleavage site identification. Furthermore, iTRAQ-TAILS also enables multiplex N-terminomics analysis of up to eight samples and allows for quantification in MS2 mode, thus preventing an increase in spectral complexity and extending proteome coverage by signal amplification of low abundance proteins. We compared the substrate degradomes of two closely related matrix metalloproteinases, MMP-2 (gelatinase A) and MMP-9 (gelatinase B), in fibroblast secreted proteins. Among 3,152 unique N-terminal peptides identified corresponding to 1,054 proteins, we detected 201 cleavage products for MMP-2 and unexpectedly only 19 for the homologous MMP-9 under identical conditions. Novel substrates identified and biochemically validated include insulin-like growth factor binding protein-4, complement C1r component A, galectin-1, dickkopf-related protein-3, and thrombospondin-2. Hence, N-terminomics analyses using iTRAQ-TAILS links gelatinases with new mechanisms of action in angiogenesis and reveals unpredicted restrictions in substrate repertoires for these two very similar proteases.

Effective proteome-wide strategies that distinguish the N-termini of proteins from the N-termini of their protease cleavage products would accelerate identification of the substrates of proteases with broad or unknown specificity. Our approach, named terminal amine isotopic labeling of substrates (TAILS), addresses this challenge by using dendritic polyglycerol aldehyde polymers that remove tryptic and C-terminal peptides. We analyze unbound naturally acetylated, cyclized or labeled N-termini from proteins and their protease cleavage products by tandem mass spectrometry, and use peptide isotope quantification to discriminate between the substrates of the protease of interest and the products of background proteolysis. We identify 731 acetylated and 132 cyclized N-termini, and 288 matrix metalloproteinase (MMP)-2 cleavage sites in mouse fibroblast secretomes. We further demonstrate the potential of our strategy to link proteases with defined biological pathways in complex samples by analyzing mouse inflammatory bronchoalveolar fluid and showing that expression of the poorly defined breast cancer protease MMP-11 in MCF-7 human breast cancer cells cleaves both endoplasmin and the immunomodulator and apoptosis inducer galectin-1.

Matrix metalloproteinases (MMPs) are important players in skin homeostasis, wound repair, and in the pathogenesis of skin cancer. It is now well established that most of their functions are related to processing of bioactive proteins rather than components of the extracellular matrix (ECM). MMP10 is highly expressed in keratinocytes at the wound edge and at the invasive front of tumors, but hardly any non-ECM substrates have been identified and its function in tissue repair and carcinogenesis is unclear. To better understand the role of MMP10 in the epidermis, we employed multiplexed iTRAQ-based Terminal Amine Isotopic Labeling of Substrates (TAILS) and monitored MMP10-dependent proteolysis over time in secretomes from keratinocytes. Time-resolved abundance clustering of neo-N termini classified MMP10-dependent cleavage events by efficiency and refined the MMP10 cleavage site specificity by revealing a so far unknown preference for glutamate in the P1 position. Moreover, we identified and validated the integrin alpha 6 subunit, cysteine-rich angiogenic inducer 61 and dermokine as novel direct MMP10 substrates and provide evidence for MMP10-dependent but indirect processing of phosphatidylethanolamine-binding protein 1. Finally, we sampled the epidermal proteome and degradome in unprecedented depth and confirmed MMP10-dependent processing of dermokine in vivo by TAILS analysis of epidermis from transgenic mice that overexpress a constitutively active mutant of MMP10 in basal keratinocytes. The newly identified substrates are involved in cell adhesion, migration, proliferation, and/or differentiation, indicating a contribution of MMP10 to local modulation of these processes during wound healing and cancer development. Data are available via ProteomeXchange with identifier PXD002474.

Stable-isotope labeling coupled with liquid-phase separation and MS analysis is a powerful technique for comparative proteomics. We developed a dimethyl labeling strategy (Anal. Chem. 2003, 75, 6843-6852 and J. Proteome Res. 2005, 4, 101-108) to label peptide N-terminus and epsilon-amino groups of Lys with water-soluble formaldehyde via reductive methylation, and an isotopic pair of formaldehyde is used for binary labeling on two sets of samples. In this study, this approach is extended to a four sample labeling by combining the binary isotopic reagents of formaldehyde (d0, d2) and the binary isotopic reducing reagents, sodium cyanoborohydride (d0, d3). To ensure sufficient mass difference, this multiplexed labeling is coupled with endoproteinase Lys-C instead of trypsin for digestion, resulting in at least two labeling sites with a mass difference of 4 Da for each pair of peptide digest. Moreover, multiplex dimethyl labeling was proved to have no significant isotopic effect during RP LC elution. This method was further applied for monitoring Lys-C digestion using hemoglobin as a model. Dimethyl labeled digests derived from seven time points (1-30 h) were grouped into two sets of sample mixtures, separated by nano-LC to reduce the complexity, and then analyzed by ESI-MS/MS. The temporal study reveals that Lys-C digestion was completed in 10-15 h for all detected peptides. The multiplex dimethyl method has not only provided a simultaneous detection mean for four sample sets but has also conserved all the advantages associated with the original binary method.

A powerful approach to relative quantification by mass spectrometry is to employ labeling reagents that target specific functional groups in molecules of interest. A quantitative comparison of two or more samples may be readily accomplished by using a chemically identical but isotopically distinct labeling reagent for each sample. The samples may then be combined, subjected to purification steps, and mass analyzed. Comparison of the signal intensities obtained from the isotopically labeled variants of the target analyte(s) provides quantitative information on their relative concentrations in the sample. In this report, we describe the synthesis and use of heavy and light isotopic forms of methyl acetimidate for the relative quantification of amine-containing species. The principal advantages of methyl acetimidate as a labeling reagent are that the reaction product is positively charged and hydrophobicity is increased, both of which enhance electrospray ionization efficiency and increase detection sensitivity. The quantitative nature of the analysis was demonstrated in model metabolomics experiments in which heavy and light labeled Arabidopsis extracts were combined in different ratios. Finally, the labeling strategy was employed to determine differences in the amounts of amine-containing metabolites for Arabidopsis seeds germinated under two different conditions.

Secreted proteases act on interstitial tissue secretomes released from multiple cell types. Thus, substrate proteins might be part of higher molecular complexes constituted by many proteins with diverse and potentially unknown cellular origin. In cell culture, these may be reconstituted by mixing native secretomes from different cell types prior to incubation with a test protease. Although current degradomics techniques could identify novel substrate proteins in these complexes, all information on the cellular origin is lost. To address this limitation, we combined iTRAQ-based terminal amine isotopic labeling of substrates (iTRAQ-TAILS) with SILAC to assign proteins to a specific cell type by MS1- and their cleavage by MS2-based quantification in the same experiment. We demonstrate the power of our newly established workflow by monitoring matrix metalloproteinase (MMP) 10 dependent cleavages in mixtures from light-labeled keratinocyte and heavy-labeled fibroblast secretomes. This analysis correctly assigned extracellular matrix components, such as laminins and collagens, to their respective cellular origins and revealed their processing in an MMP10-dependent manner. Hence, our newly devised degradomics workflow facilitates deeper insight into protease activity in complex intercellular compartments such as the epidermal-dermal interface by integrating multiple modes of quantification with positional proteomics. All MS data have been deposited in the ProteomeXchange with identifier PXD001643 (http://proteomecentral.proteomexchange.org/dataset/PXD001643).

While phosphorylation and O-GlcNAc (cytoplasmic and nuclear glycosylation) are linked to normal and pathological changes in cell states, these post-translational modifications have been difficult to analyze in proteomic studies. We describe advances in beta-elimination / Michael addition-based approaches which allow for mass spectrometry-based identification and comparative quantification of O-phosphate or O-GlcNAc-modified peptides, as well as cysteine-containing peptides for expression analysis. The method (BEMAD) involves differential isotopic labeling through Michael addition with normal dithiothreitol (DTT) (d0) or deuterated DTT (d6), and enrichment of these peptides by thiol chromatography. BEMAD was comparable to isotope-coded affinity tags (ICAT; a commercially available differential isotopic quantification technique) in protein expression analysis, but also provided the identity and relative amounts of both O-phosphorylation and O-GlcNAc modification sites. Specificity of O-phosphate vs. O-GlcNAc mapping is achieved through coupling enzymatic dephosphorylation or O-GlcNAc hydrolysis with differential isotopic labeling. Blocking of cysteine labeling by prior oxidation of a cytosolic lysate from mouse brain allowed specific targeting of serine / threonine post-translational modifications as demonstrated through identification of 21 phosphorylation sites (5 previously reported) in a single mass spectrometry analysis. These results demonstate BEMAD is suitable for large-scale quantitative analysis of both protein expression and serine / threonine post-translational modifications.

In this paper, we report a novel, stable-isotope labeling strategy for quantitative proteomics that uses a simple reagent, formaldehyde, to globally label the N-terminus and epsilon-amino group of Lys through reductive amination. This labeling strategy produces peaks differing by 28 mass units for each derivatized site relative to its nonderivatized counterpart and 4 mass units for each derivatized isotopic pair. This labeling reaction is fast (less than 5 min) and complete without any detectable byproducts based on the analysis of MALDI and LC/ESI-MS/MS spectra of both derivatized and nonderivatized peptide standards and tryptic peptides of hemoglobin molecules. The intensity of the a(1) and y(n-1) ions produced, which were not detectable from most of the nonderivatized fragments, was substantially enhanced upon labeling. We further tested the method based on the analysis of an isotopic pair of peptide standards and a pair of defined protein mixtures with known H/D ratios. Using LC/MS for quantification and LC/MS/MS for peptide sequencing, the results show a negligible isotopic effect, a good mass resolution between the isotopic pair, and a good correlation between the experimental and theoretical data (errors 0-4%). The relative standard deviation of H/D values calculated from peptides deduced from the same protein are less than 13%. The applicability of the method for quantitative protein profiling was also explored by analyzing changes in nuclear protein abundance in an immortalized E7 cell with and without arsenic treatment.

Cysteine-selective proteomics approaches simplify complex protein mixtures and improve the chance of detecting low abundant proteins. It is possible that cysteinyl-peptide/protein enrichment methods could be coupled to isotopic labeling and isobaric tagging methods for quantitative proteomics analyses in as few as two or up to 10 samples, respectively. Here we present two novel cysteine-selective proteomics approaches: cysteine-selective dimethyl labeling (cysDML) and cysteine-selective combined precursor isotopic labeling and isobaric tagging (cPILOT). CysDML is a duplex precursor quantification technique that couples cysteinyl-peptide enrichment with on-resin stable-isotope dimethyl labeling. Cysteine-selective cPILOT is a novel 12-plex workflow based on cysteinyl-peptide enrichment, on-resin stable-isotope dimethyl labeling, and iodoTMT tagging on cysteine residues. To demonstrate the broad applicability of the approaches, we applied cysDML and cPILOT methods to liver tissues from an Alzheimer’s disease (AD) mouse model and wild-type (WT) controls. From the cysDML experiments, an average of 850 proteins were identified and 594 were quantified, whereas from the cPILOT experiment, 330 and 151 proteins were identified and quantified, respectively. Overall, 2259 unique total proteins were detected from both cysDML and cPILOT experiments. There is tremendous overlap in the proteins identified and quantified between both experiments, and many proteins have AD/WT fold-change values that are within ~20% error. A total of 65 statistically significant proteins are differentially expressed in the liver proteome of AD mice relative to WT. The performance of cysDML and cPILOT are demonstrated and advantages and limitations of using multiple duplex experiments versus a single 12-plex experiment are highlighted.

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a method to accurately quantify protein expression differences between samples using mass spectrometry. ReDi labeling is performed using either regular (light) or deuterated (heavy) forms of formaldehyde and sodium cyanoborohydride to add two methyl groups to each free amine. Here we demonstrate a robust protocol for ReDi labeling and quantitative comparison of complex protein mixtures. Protein samples for comparison are digested into peptides, labeled to carry either light or heavy methyl tags, mixed, and co-analyzed by LC-MS/MS. Relative protein abundances are quantified by comparing the ion chromatogram peak areas of heavy and light labeled versions of the constituent peptide extracted from the full MS spectra. The method described here includes sample preparation by reversed-phase solid phase extraction, on-column ReDi labeling of peptides, peptide fractionation by basic pH reversed-phase (BPRP) chromatography, and StageTip peptide purification. We discuss advantages and limitations of ReDi labeling with respect to other methods for stable isotope incorporation. We highlight novel applications using ReDi labeling as a fast, inexpensive, and accurate method to compare protein abundances in nearly any type of sample.

We describe an approach for the accurate quantification and concurrent sequence identification of the individual proteins within complex mixtures. The method is based on a class of new chemical reagents termed isotope-coded affinity tags (ICATs) and tandem mass spectrometry. Using this strategy, we com- pared protein expression in the yeast Saccharomyces cerevisiae, using either ethanol or galactose as a carbon source. The measured differences in protein expression correlated with known yeast metabolic function under glucose-repressed conditions. The method is redundant if multiple cysteinyl residues are present, and the relative quantification is highly accurate because it is based on stable isotope dilution techniques. The ICAT approach should provide a widely applicable means to compare quantitatively glob- al protein expression in cells and tissues.

Data analysis in proteomics is complex and with the extra challenges involved in the interpretation of data from N-terminomics experiments, this can be daunting. Therefore, we have devised a rational pipeline of steps to approach N-terminomics data analysis in a statistically-based and valid manner. We have automated these steps in CLIPPER, an add-on to the Trans-Proteomic Pipeline (TPP). Applying CLIPPER to the analysis of N-terminomics data generated by terminal amine isotopic labeling of substrates (TAILS) enables high confidence peptide to protein assignment, protein N-terminal characterization and annotation, and for protease analysis readily allows protease substrate discovery with high confidence.

In this paper, we describe the development of a novel stable isotope N-phosphorylation labeling (SIPL) strategy for peptide de novo sequencing and protein quantification based on organic phosphorus chemistry. The labeling reaction could be performed easily and completed within 40 min in a one-pot reaction without additional cleanup procedures. It was found that N-phosphorylation labeling reagents were activated in situ to form labeling intermediates with high reactivity targeting on N-terminus and ε-amino groups of lysine under mild reaction conditions. The introduction of N-terminal-labeled phosphoryl group not only improved the ionization efficiency of peptides and increased the protein sequence coverage for peptide mass fingerprints but also greatly enhanced the intensities of b ions, suppressed the internal fragments, and reduced the complexity of the tandem mass spectrometry (MS/MS) fragmentation patterns of peptides. By using nano liquid chromatography chip/time-of-flight mass spectrometry (nano LC-chip/TOF MS) for the protein quantification, the obtained results showed excellent correlation of the measured ratios to theoretical ratios with relative errors ranging from 0.5% to 6.7% and relative standard deviation of less than 10.6%, indicating that the developed method was reproducible and precise. The isotope effect was negligible because of the deuterium atoms were placed adjacent to the neutral phosphoryl group with high electrophilicity and moderately small size. Moreover, the SIPL approach used inexpensive reagents and was amenable to samples from various sources, including cell culture, biological fluids, and tissues. The method development based on organic phosphorus chemistry offered a new approach for quantitative proteomics by using novel stable isotope labeling reagents.

Proteolysis represents one of the most tightly controlled physiological processes, as proteases create events that will typically commit pathways in an irreversible manner. Despite their implication in nearly all biological systems, our understanding of the role of proteases in disease pathology is often limited. Several approaches to studying proteolytic activity as it relates to biology, pathophysiology, and drug therapy have been published, including the recently described terminal amine isotopic labeling of substrates (TAILS) strategy by Kleifeld and colleagues. Here, we investigate TAILS as a methodology based on targeted enrichment and mass spectrometric detection of endogenous N-terminal peptides from clinically relevant biological samples and its potential to provide quantitative information on proteolysis and elucidation of the protease cleavage sites. While optimizing the most current protocol, by switching to a streamlined one-tube format and simplifying the reagents' removal steps, we demonstrate the advantages over previously published methods and provide solutions to some of the technical challenges presented in the Kleifeld publication. We also identify some of the current and unresolved limitations. We use human plasma as a model system to provide data, which illustrates some of the key analytical parameters of the modified TAILS procedure, including specificity, sensitivity, quantitative precision, and accuracy.

Analysis of the sequence and nature of protein N termini has many applications. Defining the termini of proteins for proteome annotation in the Human Proteome Project is of increasing importance. Terminomics analysis of protease cleavage sites in degradomics for substrate discovery is a key new application. Here we describe the step-by-step procedures for performing terminal amine isotopic labeling of substrates (TAILS), a 2- to 3-d (depending on method of labeling) high-throughput method to identify and distinguish protease-generated neo–N termini from mature protein N termini with all natural modifications with high confidence. TAILS uses negative selection to enrich for all N-terminal peptides and uses primary amine labeling-based quantification as the discriminating factor. Labeling is versatile and suited to many applications, including biochemical and cell culture analyses in vitro; in vivo analyses using tissue samples from animal and human sources can also be readily performed. At the protein level, N-terminal and lysine amines are blocked by dimethylation (formaldehyde/sodium cyanoborohydride) and isotopically labeled by incorporating heavy and light dimethylation reagents or stable isotope labeling with amino acids in cell culture labels. Alternatively, easy multiplex sample analysis can be achieved using amine blocking and labeling with isobaric tags for relative and absolute quantification, also known as iTRAQ. After tryptic digestion, N-terminal peptide separation is achieved using a high-molecular-weight dendritic polyglycerol aldehyde polymer that binds internal tryptic and C-terminal peptides that now have N-terminal alpha amines. The unbound naturally blocked (acetylation, cyclization, methylation and so on) or labeled mature N-terminal and neo-N-terminal peptides are recovered by ultrafiltration and analyzed by tandem mass spectrometry (MS/MS). Hierarchical substrate winnowing discriminates substrates from the background proteolysis products and non-cleaved proteins by peptide isotope quantification and bioinformatics search criteria.

Stable isotope labeling based on relative peptide/protein abundance measurements is commonly applied for quantitative proteomics. Recently, trypsin-catalyzed oxygen-18 labeling has grown in popularity due to its simplicity, cost-effectiveness, and its ability to universally label peptides with high sample recovery. In (18)O labeling, both C-terminal carboxyl group atoms of tryptic peptides can be enzymatically exchanged with (18)O, thus providing the labeled peptide with a 4 Da mass shift from the (16)O-labeled sample. Peptide (18)O labeling is ideally suited for generating a labeled "universal" reference sample used for obtaining accurate and reproducible quantitative measurements across large number of samples in quantitative discovery proteomics.

Accurate quantification of protein expression in biological systems is an increasingly important part of proteomics research. Incorporation of differential stable isotopes in samples for relative protein quantification has been widely used. Stable isotope incorporation at the peptide level using dimethyl labeling is a reliable, cost-effective and undemanding procedure that can be easily automated and applied in high-throughput proteomics experiments. Although alternative multiplex quantitative proteomics approaches introduce isotope labels at the organism level ('stable isotope labeling by amino acids in cell culture' (SILAC)) or enable the simultaneous analysis of eight samples (isobaric tagging for relative and absolute quantification (iTRAQ)), stable isotope dimethyl labeling is advantageous in that it uses inexpensive reagents and is applicable to virtually any sample. We describe in-solution, online and on-column protocols for stable isotope dimethyl labeling of sample amounts ranging from sub-micrograms to milligrams. The labeling steps take approximately 60–90 min, whereas the full protocol including digestion and (two-dimensional) liquid chromatography-mass spectrometry takes approximately 1.5–3 days to complete.

Labeling of proteins and peptides with stable heavy isotopes (deuterium, carbon-13, nitrogen-15, and oxygen-18) is widely used in quantitative proteomics. These are either incorporated metabolically in cells and small organisms, or postmetabolically in proteins and peptides by chemical or enzymatic reactions. Only upon measurement with mass spectrometers holding sufficient resolution, light, and heavy labeled peptide ions or reporter peptide fragment ions segregate and their intensity values are subsequently used for quantification. Targeted use of these labels or mass tags further leads to specific monitoring of diverse aspects of dynamic proteomes. In this review article, commonly used isotope labeling strategies are described, both for quantitative differential protein profiling and for targeted analysis of protein modifications.

Although stable isotopic labeling has found widespread use in the proteomics field, its application to carbohydrate quantification has been limited. Herein we report the design, synthesis, and application of a novel series of compounds that allow for the incorporation of isotopic variation within glycan structures. The novel feature of the compounds is the ability to incorporate the isotopes in a controlled manner, allowing for the generation of four tags that vary only in their isotopic content. This allows for the direct comparisons of three samples or triplicate measurements with an internal standard within one mass spectral analysis. Quantitation of partially depolymerized glycosaminoglycan mixtures, as well as N-linked glycans released from fetuin, is used to demonstrate the utility of the tetraplex tagging strategy.

Labeling with 18O is currently one of the most commonly used methods for incorporating a stable isotopic label into samples for comparative proteomic studies. In this approach, isotopic labeling involves the enzymatic digestion, typically performed with trypsin, of a protein population in 18O-water, which incorporates the stable isotope into the C termini of the newly formed peptides. Although trypsin is often used to facilitate isotopic incorporation after digestion, it is typically overlooked that this same mechanism can lead to isotopic loss even under conditions such as low pH where it is assumed that trypsin is inactive. To examine the role that trypsin plays in isotopic loss, several experiments were performed on the rate of delabeling under conditions relevant to multidimensional proteomic experiments. Results from these studies demonstrate that enzyme-facilitated exchange of 18O in the peptide with 16O in the aqueous solvent was the major process by which the label is removed from the peptides, even under conditions of low pH and temperature where trypsin is thought to be inactive. This study brings the rapid, tryptic-facilitated exchange to the attention of laboratories using this scheme to prevent inaccuracies in quantitative labeling due to loss of the isotopic label.

Phenotype in multicellular organisms is the consequence of dynamic metabolic events that occur in a spatially dependent fashion. This spatial and temporal complexity presents challenges for investigating metabolism; creating a need for improved methods that effectively probe biochemical events such as amino acid biosynthesis. Isotopic labeling can provide a temporal-spatial recording of metabolic events through, for example, the description of enriched amino acids in the protein pool. Proteins are therefore an important readout of metabolism and can be assessed with modern mass spectrometers. We compared the measurement of isotopic labeling in MS2 spectra obtained from tandem mass spectrometry under either higher energy collision dissociation (HCD) or collision induced dissociation (CID) at varied energy levels. Developing soybean embryos cultured with or without 13C-labeled substrates, and Escherichia coli MG1655 enriched by feeding 7% uniformly labeled glucose served as a source of biological material for protein evaluation. CID with low energies resulted in a disproportionate amount of heavier isotopologues remaining in the precursor isotopic distribution. HCD resulted in fewer quantifiable products; however deviation from predicted distributions were small relative to the CID-based comparisons. Fragment ions have the potential to provide information on the labeling of amino acids in peptides, but our results indicate that without further development the use of this readout in quantitative methods such as metabolic flux analysis is limited.

Determining the relative abundances of proteins in biological systems is an important aspect of proteomics. Quantitation provides the possibility to unravel the often subtle molecular differences that regulate biological processes in cells and organisms. A common method to analyze differences in protein expression in complex samples is differential stable isotopic labeling combined with 2D-LC-MS separation. In such experiments, proteins or peptides from different samples are labeled with different stable isotopes and their relative amounts are determined from the peptide ion intensities using mass spectrometry. When human tissue samples are investigated, chemical stable isotope labeling strategies instead of metabolic labeling strategies are required. However, biological variation in protein expression between individuals is a key concern. Here we describe a method that allows for fully automated quantitative proteome analysis; involving desalting, triplex stable isotopic dimethyl labeling and multi-dimensional strong cation exchange/reversed phase separation of peptides prior to mass spectrometric analysis that can be applied to complex samples such as human tissue lysates. We highlight the usability of the method by characterizing the extent of biological variation between the proteomes of primary human leukocytes from three healthy donors. Using our method we were able to quantify 967 proteins with a minimum of 2 peptides, revealing very limited biological variation between the donors. The discovery is noteworthy considering the presence of significant endogenous protease activity, originating primarily from the enzyme neutrophil elastase. This dataset represents the largest quantitative dataset for human leukocytesproteins, which was made possible by the use of an automated labeling strategy.