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Protein N-termini and their modifications not only represent different protein isoforms but also relate to the functional annotation and proteolytic activities. Currently, negative selection methods, such as terminal amine isotopic labeling of substrates (TAILS), are the most popular strategy to analyze the protein N-terminome, in which dimethylation or acetylation modification is commonly used to block the free amines of proteome samples. However, after tryptic digestion, the generated long peptides, caused by the missing cleavage of blocked lysine, could hardly be identified by MS, which hindered the deep-coverage analysis of N-terminome. Herein, to solve this problem, we developed an approach, named terminal amine guanidination of substrates (TAGS). 1H-Pyrazole-1-carboxamidine was used to effectively guanidinate lysine e-amines and N-terminal α-amines, followed by tryptic digestion to generate N-terminal peptides without free amines and internal peptides with free amines. Then, the internal peptides with free amines were removed by hyperbranched polyglycerol-aldehyde polymers (HPG-ALDs) to achieve the negative enrichment of N-terminome. By TAGS, not only the cleavage rate of blocked lysine could be improved, but also the ionization efficiency of tryptic peptides was increased. In comparison, 1814 and 1620 protein N-termini were, respectively, identified by TAGS and TAILS in Saccharomyces cerevisiae (S. cerevisiae). Among them, 1012 N-termini were uniquely identified in TAGS. Furthermore, by the combination of TAGS and the stable isotope labeling with amino acids in cell culture (SILAC)/label-free quantitative method, we not only identified the known N-terminal cleavage fragment of gasdermin D but also identified some new cleavage sites during Val-boroPro-induced pyroptosis. All these results demonstrated that our developed approach, TAGS, might be of great promise for the comprehensive analysis of N-terminome and beneficial for promoting the identification of protein isoforms and studying in-depth the proteolytic activity of proteins.

Mass spectrometry-based stable isotope labeling has become a key technology for protein and small-molecule analyses. We developed a multiplexed quantification method for simultaneous proteomics and amine metabolomics analyses via nano reversed-phase liquid chromatography–tandem mass spectrometry (nanoRPLC–MS/MS), called mass defect-based N,N-dimethyl leucine (mdDiLeu) labeling. The duplex mdDiLeu reagents were custom-synthesized with a mass difference of 20.5 mDa, arising from the subtle variation in nuclear binding energy between the two DiLeu isotopologues. Optimal MS resolving powers were determined to be 240K for labeled peptides and 120K for labeled metabolites on the Orbitrap Fusion Lumos instrument. The mdDiLeu labeling does not suffer from precursor interference and dynamic range compression, providing excellent accuracy for MS1-centric quantification. Quantitative information is only revealed at high MS resolution without increasing spectrum complexity and overlapping isotope distribution. Chromatographic performance of polar metabolites was dramatically improved by mdDiLeu labeling with modified hydrophobicity, enhanced ionization efficiency, and picomole levels of detection limits. Paralleled proteomics and amine metabolomics analyses using mdDiLeu were systematically evaluated and then applied to pancreatic cancer cells.

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.

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.

A new method for proteolytic stable isotope labeling is introduced to provide quantitative and concurrent comparisons between individual proteins from two entire proteome pools or their subfractions. Two 18O atoms are incorporated universally into the carboxyl termini of all tryptic peptides during the proteolytic cleavage of all proteins in the first pool. Proteins in the second pool are cleaved analogously with the carboxyl termini of the resulting peptides containing two 16O atoms (i.e., no labeling). The two peptide mixtures are pooled for fractionation and separation, and the masses and isotope ratios of each peptide pair (differing by 4 Da) are measured by high-resolution mass spectrometry. Short sequences and/or accurate mass measurements combined with proteomics software tools allow the peptides to be related to the precursor proteins from which they are derived. Relative signal intensities of paired peptides quantify the expression levels of their precursor proteins from proteome pools to be compared, using an equation described in the paper. Observation of individual (unpaired) peptides is mainly interpreted as differential modification or sequence variation for the protein from the respective proteome pool. The method is evaluated here in a comparison of virion proteins for two serotypes (Ad5 and Ad2) of adenovirus, taking advantage of information already available about protein sequences and concentrations. In general, proteolytic 18O labeling enables a shotgun approach for proteomic studies with quantitation capability and is proposed as a useful tool for comparative proteomic studies of very complex protein mixtures.

While peptide-level labeling using isobaric tag reagents has been widely applied for quantitative proteomics experiments, there are comparatively few reports of protein-level labeling. Intact protein labeling could be broadly applied to quantification experiments utilizing protein-level separations or enrichment schemes. Here, protein-level isobaric labeling was explored as an alternative strategy to peptide-level labeling for serum glycoprotein quantification. Labeling and digestion conditions were optimized by comparing different organic solvents and enzymes. Digestions with Asp-N and trypsin were found highly complementary; combining the results enabled quantification of 30% more proteins than either enzyme alone. Three commercial reagents were compared for protein-level labeling. Protein identification rates were highest with iTRAQ 4-plex when compared to TMT 6-plex and iTRAQ 8-plex using higher-energy collisional dissociation on an Orbitrap Elite mass spectrometer. The compatibility of isobaric protein-level labeling with lectin-based glycoprotein enrichment was also investigated. More than 74% of lectin-bound labeled proteins were known glycoproteins, which was similar to results from unlabeled and peptide-level labeled serum samples. Finally, protein-level and peptide-level labeling strategies were compared for serum glycoprotein quantification. Isobaric protein-level labeling gave comparable identification levels and quantitative precision to peptide-level labeling.

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.

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.