Exploring skyline for both MSE‐based label‐free proteomics and HRMS quantitation of small molecules

The MS^E (where MS^E is low energy (MS) and elevated energy (E) mode of acquisition) acquisition method commercialized by Waters on its Q‐TOF instruments is regarded as a unique data‐independent fragmentation approach that improves the accuracy and dynamic range of label‐free proteomic quantitation. Due to its special format, MS^E acquisition files cannot be independently analyzed with most widely used open‐source proteomic software specialized for processing data‐dependent acquisition files. In this study, we established a workflow integrating Skyline, a popular and versatile peptide‐centric quantitation program, and a statistical tool DiffProt to fulfill MS^E‐based proteomic quantitation. Comparison with the vendor software package for analyzing targeted phosphopeptides and global proteomic datasets reveals distinct advantages of Skyline in MS^E data mining, including sensitive peak detection, flexible peptide filtering, and transparent step‐by‐step workflow. Moreover, we developed a new procedure such that Skyline MS1 filtering was extended to small molecule quantitation for the first time. This new utility of Skyline was examined in a protein–ligand interaction experiment to identify multiple chemical compounds specifically bound to NDM‐1 (where NDM is New Delhi metallo‐β‐lactamase 1), an antibiotics‐resistance target. Further improvement of the current weaknesses in Skyline MS1 filtering is expected to enhance the reliability of this powerful program in full scan‐based quantitation of both peptides and small molecules.     

Development of novel mass spectrometric methods for identifying HOCl‐induced modifications to proteins

Protein oxidation is thought to contribute to a number of inflammatory diseases, hence the development of sensitive and specific analytical techniques to detect oxidative PTMs (oxPTMs) in biological samples is highly desirable. Precursor ion scanning for fragment ions of oxidized amino acid residues was investigated as a label‐free MS approach to mapping specific oxPTMs in a complex mixture of proteins. Using HOCl‐oxidized lysozyme as a model system, it was found that the immonium ions of oxidized tyrosine and tryptophan formed in MS² analysis could not be used as diagnostic ions, owing to the occurrence of isobaric fragment ions from unmodified peptides. Using a double quadrupole linear ion trap mass spectrometer, precursor ion scanning was combined with detection of MS³ fragment ions from the immonium ions and collisionally‐activated decomposition peptide sequencing to achieve selectivity for the oxPTMs. For chlorotyrosine, the immonium ion at 170.1 m/z fragmented to yield diagnostic ions at 153.1, 134.1, and 125.1 m/z, and the hydroxytyrosine immonium ion at 152.1 m/z gave diagnostic ions at 135.1 and 107.1 m/z. Selective MS³ fragment ions were also identified for 2‐hydroxytryptophan and 5‐hydroxytryptophan. The method was used successfully to map these oxPTMs in a mixture of nine proteins that had been treated with HOCl, thereby demonstrating its potential for application to complex biological samples.     

Electron transfer dissociation of iTRAQ labeled peptide ions

Triply and doubly charged iTRAQ ( isobaric tagging for relative and absolute quantitation) labeled peptide cations from a tryptic peptide mixture of bovine carbonic anhydrase II were subjected to electron transfer ion/ion reactions to investigate the effect of charge bearing modifications associated with iTRAQ on the fragmentation pattern. It was noted that electron transfer dissociation (ETD) of triply charged or activated ETD (ETD and supplemental collisional activation of intact electron transfer species) of doubly charged iTRAQ tagged peptide ions yielded extensive sequence information, in analogy with ETD of unmodified peptide ions. That is, addition of the fixed charge iTRAQ tag showed relatively little deleterious effect on the ETD performance of the modified peptides. ETD of the triply charged iTRAQ labeled peptide ions followed by collision-induced dissociation (CID) of the product ion at m/ z 162 yielded the reporter ion at m/ z 116, which is the reporter ion used for quantitation via CID of the same precursor ions. The reporter ion formed via the two-step activation process is expected to provide quantitative information similar to that directly produced from CID. A 103 Da neutral loss species observed in the ETD spectra of all the triply and doubly charged iTRAQ labeled peptide ions is unique to the 116 Da iTRAQ reagent, which implies that this process also has potential for quantitation of peptides/proteins. Therefore, ETD with or without supplemental collisional activation, depending on the precursor ion charge state, has the potential to directly identify and quantify the peptides/proteins simultaneously using existing iTRAQ reagents.     

LTQ‐iQuant: A freely available software pipeline for automated and accurate protein quantification of isobaric tagged peptide data from LTQ instruments

Pulsed Q dissociation enables combining LTQ ion trap instruments with isobaric peptide tagging. Unfortunately, this combination lacks a technique which accurately reports protein abundance ratios and is implemented in a freely available, flexible software pipeline. We developed and implemented a technique assigning collective reporter ion intensity‐based weights to each peptide abundance ratio and calculating a protein's weighted average abundance ratio and p‐value. Using an iTRAQ‐labeled standard mixture, we compared our technique's performance to the commercial software MASCOT, finding that it performed better than MASCOT's nonweighted averaging and median peptide ratio techniques, and equal to its weighted averaging technique. We also compared performance of the LTQ‐Orbitrap plus our technique to 4800 MALDI TOF/TOF plus Protein Pilot, by analyzing an iTRAQ‐labeled stem cell lysate. We found highly correlated protein abundance ratios, indicating that the LTQ‐Orbitrap plus our technique yields results comparable to the current standard. We implemented our technique in a freely available, automated software pipeline, called LTQ‐iQuant, which is mzXML‐compatible; supports iTRAQ 4‐plex and 8‐plex LTQ data; and can be modified for and have weights trained to a user's LTQ and other isobaric peptide tagging methods. LTQ‐iQuant should make LTQ instruments and isobaric peptide tagging accessible to more proteomic researchers.     

Enhancing Data Reliability in TOMAHAQ for Large‐Scale Protein Quantification

The analytical scale of most mass‐spectrometry‐based targeted proteomics assays is usually limited by assay performance and instrument utilization. A recently introduced method, called triggered by offset, multiplexed, accurate mass, high resolution, and absolute quantitation (TOMAHAQ), combines both peptide and sample multiplexing to simultaneously improve analytical scale and quantitative performance. In the present work, critical technical requirements and data analysis considerations for successful implementation of the TOMAHAQ technique based on the study of a total of 185 target peptides across over 200 clinical plasma samples are discussed. Importantly, it is observed that significant interference originate from the TMTzero reporter ion used for the synthetic trigger peptides. This interference is not expected because only TMT10plex reporter ions from the target peptides should be observed under typical TOMAHAQ conditions. In order to unlock the great promise of the technique for high throughput quantification, here a post‐acquisition data correction strategy to deconvolute the reporter ion superposition and recover reliable data is proposed.     

Ultra-high intra-spectrum mass accuracy enables unambiguous identification of fragment reporter ions in isobaric multiplexed quantitative proteomics

Isobaric tagging using reagents such as tandem mass tags (TMT) and isobaric tags for relative and absolute quantification (iTRAQ) have become popular tools for mass spectrometry based quantitative proteomics. Because the peptide quantification information is collected in tandem mass spectra, the accuracy and precision of this method largely depend on the resolution with which precursor ions can be selected for the fragmentation and the specificity of the generated reporter ion. The latter can constitute an issue if near isobaric ion signals are present in such spectra because they may distort quantification results. We propose a simple remedy for this problem by identifying reporter ions via the accurate mass differences within a single tandem mass spectrum instead of applying fixed mass error tolerances for all tandem mass spectra. Our results show that this leads to unambiguous reporter ion identification and complete removal of interfering signals. This mode of data processing is easily implemented in software and offers advantages for protein quantification based on few peptides.     

High‐performance hybrid Orbitrap mass spectrometers for quantitative proteome analysis: Observations and implications

We present basic workups and quantitative comparisons for two current generation Orbitrap mass spectrometers, the Q Exactive Plus and Orbitrap Fusion Tribrid, which are widely considered two of the highest performing instruments on the market. We assessed the performance of two quantitative methods on both instruments, namely label‐free quantitation and stable isotope labeling using isobaric tags, for studying the heat shock response in Escherichia coli. We investigated the recently reported MS3 method on the Fusion instrument and the potential of MS3‐based reporter ion isolation Synchronous Precursor Selection (SPS) and its impact on quantitative accuracy. We confirm that the label‐free approach offers a more linear response with a wider dynamic range than MS/MS‐based isobaric tag quantitation and that the MS3/SPS approach alleviates but does not eliminate dynamic range compression. We observed, however, that the choice of quantitative approach had little impact on the ability to statistically evaluate the E. coli heat shock response. We conclude that in the experimental conditions tested, MS/MS‐based reporter ion quantitation provides reliable biological insight despite the issue of compressed dynamic range, an observation that significantly impacts the choice of instrument.     

Amine-reactive Neutron-encoded Labels for Highly Plexed Proteomic Quantitation

We describe a novel amine-reactive chemical label that exploits differential neutron-binding energy between ¹³C and ¹⁵N isotopes. These neutron-encoded (NeuCode) chemical labels enable up to 12-plex MS1-based protein quantification. Each structurally identical, but isotopically unique, tag is encoded with a 12.6-mDa mass difference—relative to its nearest neighbor—so that peptides bearing these NeuCode signatures do not increase spectral complexity and are detected only upon analysis with very high mass-resolving powers. We demonstrate that the method provides quantitative performance that is comparable to both metabolic labeling and isobaric tagging while combining the benefits of both strategies. Finally, we employ the tags to characterize the proteome of Saccharomyces cerevisiae during the diauxic shift, a metabolic transition from fermentation to aerobic respiration.Proteome quantification is an increasingly essential component of modern biology and translational medicine (1, 2). Whether targeted or global, stable isotope incorporation with mass spectrometry (MS) analysis is a core technique for protein abundance measurements. There are numerous approaches that can be used to introduce stable isotopes into peptides, the most frequently used being stable isotope labeling with amino acids in cell culture (SILAC)¹ and isobaric tagging (tandem mass tags or isobaric tags for relative and absolute quantitation) (3–7). Both of these methods incorporate heavy isotopes to increase mass by at least 1 Da. SILAC is the quantification gold standard for global proteomic analysis. However, the SILAC approach is not easily adapted for tissue sample analysis; SILAC mouse labeling, for example, requires feeding mice a specialized diet for multiple generations (8). Tissue samples can be analyzed if they are mixed with SILAC cell-culture-based labeled standards, but this strategy does not permit multiplexing (9). Isobaric labels, in contrast, are conjugated to the primary amines of peptides following proteolytic digestion and thus have the advantage of being completely compatible with samples from virtually any source (10, 11). That said, isobaric tagging suffers from dynamic range suppression caused by co-isolation of precursor peptides (12). Multiple studies have revealed that this problem greatly erodes quantitative accuracy—for example, 10-fold changes often are detected as much smaller ∼4-fold changes (13, 14). Here we propose a new approach to protein quantification, one that achieves tissue-compatible 4-plexed MS1-based quantification without increasing spectral complexity.Recently, we described the use of mass defects to expand SILAC quantification from 3-plex to 12-plex and beyond (i.e., NeuCode SILAC) (15). NeuCode SILAC exploits the subtle mass differences that exist in atoms as a result of the varying energies of nuclear binding in common stable isotopes (e.g., carbon, nitrogen, hydrogen, and oxygen) by using the extremely high resolving power of modern Fourier transform mass spectrometer systems (16, 17). For example, the multiplexing capability of tandem mass tagging was increased from 6 to 8 by incorporating a difference in mass of 6.3 mDa in specific reporter ions by swapping ¹⁴N for a ¹⁵N atom while concomitantly switching a ¹³C with a ¹²C atom (18, 19). This method requires only 30,000 resolving power to resolve the reporter ions, but it still suffers from the interference problem described above. Repetition of this process, within the context of an analyte molecule, can generate several chemically identical isotopologues that, when analyzed under normal MS analysis conditions (resolving power R) are indistinguishable (i.e., produce one m/z peak). Analysis of these NeuCode m/z peaks with high resolving power (480,000), however, often reveals distinct m/z peaks whose abundances can be extracted and used to determine analyte quantity across the sundry conditions. This strategy permits very high levels of MS1-based multiplexing (>10), which has several advantages. First, MS1 scans across the entire analyte elution profile can be averaged to increase quantitative accuracy and precision. Second, a tandem mass spectrum is not necessary for quantification. Mann and colleagues showed that once a peptide is identified via tandem MS analysis, it can be confidently identified in other runs with only the exact mass and elution profile matching (20). Third, MS1-based quantification does not suffer from the pervasive problem of precursor interference that cripples the quantitative accuracy of the isobaric tagging strategies (13).The NeuCode SILAC approach, however, relies on the use of amino acid isotopologues and still requires metabolic incorporation. Chemical labeling strategies for proteome quantification can be convenient and, for certain systems, are requisite. We reasoned that our NeuCode strategy could be extended to create novel chemical reagents for proteome quantification. We describe here the design, synthesis, and use of 12-plex NeuCode amine reactive labels for global proteome quantification.     

Amine-reactive isobaric tagging reagents: requirements for absolute quantification of proteins and peptides

Amine-reactive isobaric tagging reagents such as iTRAQ (isobaric tags for relative and absolute quantitation) have recently become increasing popular for relative protein quantification, cell expression profiling, and biomarker discovery. This is due mainly to the possibility of simultaneously identifying and quantifying multiple samples. The principles of iTRAQ may also be applied to absolute protein quantification with the use of synthetic peptides as standards. The prerequisites that must be fulfilled to perform absolute quantification of proteins by iTRAQ have been investigated and are described here. Three samples of somatropin were quantified using iTRAQ and synthetic peptides as standards, corresponding to a portion of the protein sequence. The results were compared with those obtained by quantification of the same protein solutions using double exact matching isotope dilution mass spectrometry (IDMS). To obtain reliable results, the appropriate standard peptides needed to be selected carefully and enzymatic digestion needed to be optimized to ensure complete release of the peptides from the protein. The kinetics and efficiency of the iTRAQ derivatization reaction of the standard peptides and digested proteins with isobaric tagging reagents were studied using a mixture of seven synthetic peptides and their corresponding labeled peptides. The implications of incomplete derivatization are also presented.     

Protein labeling by iTRAQ: a new tool for quantitative mass spectrometry in proteome research

A novel, MS-based approach for the relative quantification of proteins, relying on the derivatization of primary amino groups in intact proteins using isobaric tag for relative and absolute quantitation (iTRAQ) is presented. Due to the isobaric mass design of the iTRAQ reagents, differentially labeled proteins do not differ in mass; accordingly, their corresponding proteolytic peptides appear as single peaks in MS scans. Because quantitative information is provided by isotope-encoded reporter ions that can only be observed in MS/MS spectra, we analyzed the fragmentation behavior of ESI and MALDI ions of peptides generated from iTRAQ-labeled proteins using a TOF/TOF and/or a QTOF instrument. We observed efficient liberation of reporter ions for singly protonated peptides at low-energy collision conditions. In contrast, increased collision energies were required to liberate the iTRAQ label from lysine side chains of doubly charged peptides and, thus, to observe reporter ions suitable for relative quantification of proteins with high accuracy. We then developed a quantitative strategy that comprises labeling of intact proteins by iTRAQ followed by gel electrophoresis and peptide MS/MS analyses. As proof of principle, mixtures of five different proteins in various concentration ratios were quantified, demonstrating the general applicability of the approach presented here to quantitative MS-based proteomics.     

Comparative Analyses of Data Independent Acquisition Mass Spectrometric Approaches: DIA,WiSIM‐DIA,and Untargeted DIA

Data‐independent acquisition (DIA) is an emerging technology for quantitative proteomics. Current DIA focusses on the identification and quantitation of fragment ions that are generated from multiple peptides contained in the same selection window of several to tens of m/z. An alternative approach is WiSIM‐DIA, which combines conventional DIA with wide‐SIM (wide selected‐ion monitoring) windows to partition the precursor m/z space to produce high‐quality precursor ion chromatograms. However, WiSIM‐DIA has been underexplored; it remains unclear if it is a viable alternative to DIA. We demonstrate that WiSIM‐DIA quantified more than 24 000 unique peptides over five orders of magnitude in a single 2 h analysis of a neuronal synapse‐enriched fraction, compared to 31 000 in DIA. There is a strong correlation between abundance values of peptides quantified in both the DIA and WiSIM‐DIA datasets. Interestingly, the S/N ratio of these peptides is not correlated. We further show that peptide identification directly from DIA spectra identified >2000 proteins, which included unique peptides not found in spectral libraries generated by DDA.     

Isobaric Labeling and Data Normalization without Requiring Protein Quantitation

Isobaric multiplexed quantitative proteomics can complement high-resolution sample isolation techniques. Here, we report a simple workflow exponentially modified protein abundance index (emPAI)-MW deconvolution (EMMOL) for normalizing isobaric reporter ratios within and between experiments, where small or unknown amounts of protein are used. EMMOL deconvolutes the isobaric tags for relative and absolute quantification (iTRAQ) data to yield the quantity of each protein of each sample in the pool, a new approach that enables the comparison of many samples without including a channel of reference standard. Moreover, EMMOL allows using a sufficient quantity of control sample to facilitate the peptide fractionation (isoelectric-focusing was used in this report), and mass spectrometry MS/MS sequencing yet relies on the broad dynamic range of iTRAQ quantitation to compare relative protein abundance. We demonstrated EMMOL by comparing four pooled samples with 20-fold range differences in protein abundance and performed data normalization without using prior knowledge of the amounts of proteins in each sample, simulating an iTRAQ experiment without protein quantitation prior to labeling. We used emPAI,¹ the target protein MW, and the iTRAQ reporter ratios to calculate the amount of each protein in each of the four channels. Importantly, the EMMOL-delineated proteomes from separate iTRAQ experiments can be assorted for comparison without using a reference sample. We observed no compression of expression in iTRAQ ratios over a 20-fold range for all protein abundances. To complement this ability to analyze minute samples, we report an optimized iTRAQ labeling protocol for using 5 μg protein as the starting material.     

Multiplexed protein quantitation in Saccharomyces cerevisiae using amine-reactive isobaric tagging reagents

We describe here a multiplexed protein quantitation strategy that provides relative and absolute measurements of proteins in complex mixtures. At the core of this methodology is a multiplexed set of isobaric reagents that yield amine-derivatized peptides. The derivatized peptides are indistinguishable in MS, but exhibit intense low-mass MS/MS signature ions that support quantitation. In this study, we have examined the global protein expression of a wild-type yeast strain and the isogenic upf1Delta and xrn1Delta mutant strains that are defective in the nonsense-mediated mRNA decay and the general 5' to 3' decay pathways, respectively. We also demonstrate the use of 4-fold multiplexing to enable relative protein measurements simultaneously with determination of absolute levels of a target protein using synthetic isobaric peptide standards. We find that inactivation of Upf1p and Xrn1p causes common as well as unique effects on protein expression.     

Candidate verification of iron-regulated Neisseria meningitidis proteins using isotopic versions of tandem mass tags (TMT) and single reaction monitoring

Tandem Mass Tags (TMT) are suited to both global and targeted quantitation approaches of proteins and peptides. Different versions of these tags allow for the generation of both isobaric and isotopic sets of reagents sharing the same common structure. This feature allows for a straightforward transfer of data obtained during discovery studies into targeted investigations. In prior discovery studies, an isobaric set of these reagents was used to identify Neisseria meningitidis proteins expressed under iron-limitation. Here, we apply isotopic versions of those reagents in combination with single reaction monitoring to verify selected candidates found to be differentially regulated in these discovery studies, representing both well-known and novel iron-regulated proteins, such as the MtrCDE drug efflux pump. In this targeted approach (TMT–SRM), the selectivity of SRM is maintained while allowing the incorporation of an internal reference standard into the experiment. By monitoring 184 transitions, TMT–SRM resulted in the quantitation of 33 peptides representing 12 proteins. The acquired data corroborated the results obtained during the discovery phase. Furthermore, these data obtained by MS-based quantitation of peptides were independently confirmed by western blotting results, an orthogonal approach based on quantitation at the protein level.     

Twoplex 12/13C6 aniline stable isotope and linkage‐specific sialic acid labeling 2D‐LC‐MS workflow for quantitative N‐glycomics

Quantitative glycomics represents an actively expanding research field ranging from the discovery of disease‐associated glycan alterations to the quantitative characterization of N‐glycans on therapeutic proteins. Commonly used analytical platforms for comparative relative quantitation of complex glycan samples include MALDI‐TOF‐MS or chromatographic glycan profiling with subsequent data alignment and statistical evaluation. Limitations of such approaches include run‐to‐run technical variation and the potential introduction of subjectivity during data processing. Here, we introduce an offline 2D LC‐MS^E workflow for the fractionation and relative quantitation of twoplex isotopically labeled N‐linked oligosaccharides using neutral ¹²C₆ and ¹³C₆ aniline (Δmass = 6 Da). Additional linkage‐specific derivatization of sialic acids using 4‐(4,6‐dimethoxy‐1,3,5‐trizain‐2‐yl)‐4‐methylmorpholinium chloride offered simultaneous and advanced in‐depth structural characterization. The potential of the method was demonstrated for the differential analysis of structurally defined N‐glycans released from serum proteins of patients diagnosed with various stages of colorectal cancer. The described twoplex ¹²C₆/¹³C₆ aniline 2D LC‐MS platform is ideally suited for differential glycomic analysis of structurally complex N‐glycan pools due to combination and analysis of samples in a single LC‐MS injection and the associated minimization in technical variation.     

Whole cell,label free protein quantitation with data independent acquisition: Quantitation at the MS2 level

Label free quantitation by measurement of peptide fragment signal intensity (MS2 quantitation) is a technique that has seen limited use due to the stochastic nature of data dependent acquisition (DDA). However, data independent acquisition has the potential to make large scale MS2 quantitation a more viable technique. In this study we used an implementation of data independent acquisition—SWATH—to perform label free protein quantitation in a model bacterium Clostridium stercorarium. Four tryptic digests analyzed by SWATH were probed by an ion library containing information on peptide mass and retention time obtained from DDA experiments. Application of this ion library to SWATH data quantified 1030 proteins with at least two peptides quantified (～40% of predicted proteins in the C. stercorarium genome) in each replicate. Quantitative results obtained were very consistent between biological replicates (R² ～ 0.960). Protein quantitation by summation of peptide fragment signal intensities was also highly consistent between biological replicates (R² ～ 0.930), indicating that this approach may have increased viability compared to recent applications in label free protein quantitation. SWATH based quantitation was able to consistently detect differences in relative protein quantity and it provided coverage for a number of proteins that were missed in some samples by DDA analysis.     

Sequential Window Acquisition of all Theoretical Mass Spectra (SWATH) Analysis for Characterization and Quantification of Histone Post-translational Modifications

Histone post-translational modifications (PTMs) have a fundamental function in chromatin biology, as they model chromatin structure and recruit enzymes involved in gene regulation, DNA repair, and chromosome condensation. High throughput characterization of histone PTMs is mostly performed by using nano-liquid chromatography coupled to mass spectrometry. However, limitations in speed and stochastic sampling of data dependent acquisition methods in MS lead to incomplete discrimination of isobaric peptides and loss of low abundant species. In this work, we analyzed histone PTMs with a data-independent acquisition method, namely SWATH™ analysis. This approach allows for MS/MS-based quantification of all analytes without upfront assay development and no issues of biased and incomplete sampling. We purified histone proteins from human embryonic stem cells and mouse trophoblast stem cells before and after differentiation, and prepared them for MS analysis using the propionic anhydride protocol. Results on histone H3 peptides verified that sequential window acquisition of all theoretical mass spectra could accurately quantify peptides (<9% average coefficient of variation, CV) over four orders of magnitude, and we could discriminate isobaric and co-eluting peptides (e.g. H3K18ac and H3K23ac) using MS/MS-based quantification. This method provided high sensitivity and precision, supported by the fact that we could find significant differences for remarkably low abundance PTMs such as H3K9me2S10ph (relative abundance <0.02%). We performed relative quantification for few sample peptides using different fragment ions and observed high consistency (CV <15%) between the fragments. This indicated that different fragment ions can be used independently to achieve the same peptide relative quantification. Taken together, sequential window acquisition of all theoretical mass spectra proved to be an easy-to-use MS acquisition method to perform high quality MS/MS-based quantification of histone-modified peptides.Chromatin is a highly organized and dynamic entity in cell nuclei, mostly composed of DNA and histone proteins. Its structure directly influences gene expression, DNA repair, and cell duplication events such as mitosis and meiosis (1). Histones are assembled in octamers named nucleosomes, wrapped by DNA every ∼200 base pairs. Histones are heavily modified by dynamic post-translational modifications (PTMs)¹, which affect chromatin structure because of their chemical properties and their ability to recruit chromatin modifier enzymes and binding proteins (2). Moreover, histone PTMs can be inherited through cell division and thus are crucial components of epigenetic memory (3). The function of histone PTMs has been extensively studied in the last 15–20 years, and several links have been found between aberrations of histone PTM levels and development of diseases (4, 5). Such discoveries revealed the importance of histone PTMs in fine-tuning cell phenotype. Because of this, technology has been rapidly evolving to investigate histone PTM relative abundance with higher accuracy and throughput.Mass spectrometry (MS)-based strategies have continuously evolved toward higher throughput and flexibility, allowing not only identification and quantification of single histone PTMs, but also their combinatorial patterns and even characterization of the intact proteins (reviewed in (6, 7)). For histone analysis, a widely adopted workflow for nano-liquid chromatography–tandem mass spectrometry (nLC-MS/MS) includes derivatization of lysine residue side chains with propionic anhydride, proteolytic digestion with trypsin, and subsequent derivatization of peptide N termini (8, 9). Such protocol leads to generation of ArgC-like peptides (only cleaved after arginine residues) after digestion. Moreover, propionylation of N termini increases peptide hydrophobicity, thereby improving LC retention of shorter ones, and thus the MS signal. Because of the high mass accuracy, sensitivity, and the possibility to perform label-free quantification MS has become the technique of choice, outperforming antibody-based strategies, to study both known and novel global histone PTMs.Several acquisition methods have been developed for MS analysis to accomplish different needs of identification and quantification (10). The most widely adopted in shotgun or discovery proteomics is the data-dependent acquisition (DDA) mode. Such acquisition method does not require any previous knowledge about the analyte, as it automatically selects precursor ions detectable at the full scan level in a given order (commonly from the most intense) to perform MS/MS fragmentation (11). Label-free quantification is performed at the full MS scan level by integrating the area of the LC peak from an extracted ion chromatogram of the precursor mass corresponding to the given peptide. On the other hand, the selected reaction monitoring (SRM) mode is the most widely used acquisition method in targeted proteomics. Such method performs cyclic precursor ion selection, MS/MS fragmentation, and product ion selection of a list of masses input by the user. Even though the method preparation is intuitively more complex than DDA, SRM is highly popular because of the high selectivity and sensitivity, which leads to more accurate label-free quantification (12). However, both methods have inevitable drawbacks; a DDA approach cannot perform accurate quantification of isobaric and co-eluting peptides, for example, KacQLATKAAR and KQLATKacAAR (histone H3 aa 9–17), as the fragment ions should be monitored through the entire peptide peak elution to define the ratio between the two similar analytes. On the contrary, an SRM experiment prevents future data mining of unpredicted peptides, and thus such method cannot be used for any classical PTM discovery. Therefore, LC-MS/MS analysis of histone peptides is commonly performed by integrating shotgun and targeted acquisition within the same MS method (13). This method requires previous knowledge about retention time and mass of co-eluting isobaric species, and tedious manual peak integration or dedicated software to deconvolute such complex raw data. Although this mixed MS mode is a powerful approach, the targeted sequences in the method reduce the duty cycle and number of DDA MS/MS spectra that can be acquired, making it far from ideal.Data independent acquisition (DIA) modes are a third option that recently gained popularity in proteomics (14, 15). Sequential window acquisition of all theoretical mass spectra (SWATH™)-MS is a data independent workflow that uses a first quadrupole isolation window to step across a mass range, collecting high resolution full scan composite MS/MS at each step and generating an ion map of fragments from all detectable precursor masses (15, 16). From such data set, a virtual SRM, or pseudo-SRM, can be performed by extracting the product ion chromatogram of a given peptide (17) with bioinformatics tools such as Peakview®, Skyline (18), or OpenSWATH™ (19). In order to define which fragment masses should be used to quantify a given peptide, a spectral library of identified peptides can be manually programmed, downloaded (if available), or generated by previous DDA experiments. In terms of quantification power, SWATH™ combines the advantages of both DDA and SRM, as it allows for MS/MS-based label-free quantification, discrimination of isobaric peptides, and subsequent data mining of unpredicted species.Histone proteins are an excellent target sample to test SWATH™, as the peptides are heavily modified by PTMs and often have isobaric proteoforms present. We analyzed with both DDA and SWATH™ two model systems: (1) extracted histones from untreated (pluripotent) and retinoic acid (RA) treated (differentiated) human embryonic stem cells (hESCs, strain H9), and (2) extracted histones from undifferentiated and differentiated mouse trophoblast stem cells (mTSCs). The results from the DDA experiment were used to evaluate the reproducibility of peptide retention time and the variety of species identified. For the SWATH™ analysis we focused on histone H3, as it is the histone with the highest variety of modified peptides (6). Results highlighted that such acquisition method provides sensitive and precise MS/MS-based quantification of both isobaric and nonisobaric peptides. Our data demonstrate that quantification at the MS/MS level is highly reproducible, and identification of the peptide elution profile is assisted by the high mass accuracy and the large number of overlapping elution profiles of the fragment ions. Moreover, we show that by using different fragment ions for MS/MS quantification we achieved similar quantification results. Thus, we used all unique fragment ions for a given species to provide a robust quantification method, where by unique is intended fragment ions that belong to only one of the possible isobaric peptide proteoforms. Taken together, we prove that SWATH™-MS is a reliable and simple-to-use acquisition method to perform epigenetic histone PTM analysis.     

Higher-energy collision-activated dissociation without a dedicated collision cell

Beam-type collisional activation dissociation (HCD) offers many advantages over resonant excitation collision-activated dissociation, including improved identification of phosphorylated peptides and compatibility with isobaric tag-based quantitation (e.g. tandem mass tag (TMT) and iTRAQ). However, HCD typically requires specially designed and dedicated collision cells. Here we demonstrate that HCD can be performed in the ion injection pathway of a mass spectrometer with a standard atmospheric inlet (iHCD). Testing this method on complex peptide mixtures revealed similar identification rates to collision-activated dissociation (2883 versus 2730 IDs for iHCD/CAD, respectively) and precursor-product-conversion efficiency comparable to that achieved within a dedicated collision cell. Compared with pulsed-q dissociation, a quadrupole ion trap-based method that retains low-mass isobaric tag reporter ions, iHCD yielded isobaric tag for relative and absolute quantification reporter ions 10-fold more intense. This method involves no additional hardware and can theoretically be implemented on any mass spectrometer with an atmospheric inlet.     

Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on Arabidopsis thaliana, a genetically tractable host plant^1,2. The accumulation of reactive oxygen species (ROS) in cotyledons inoculated with DC3000 indicates a role of ROS in modulating necrotic cell death during bacterial speck disease of tomato³. Hydrogen peroxide, a component of ROS, is produced after inoculation of tomato plants with Pseudomonas³. Hydrogen peroxide can be detected using a histochemical stain 3''-3'' diaminobenzidine (DAB)⁴. DAB staining reacts with hydrogen peroxide to produce a brown stain on the leaf tissue⁴. ROS has a regulatory role of the cellular redox environment, which can change the redox status of certain proteins⁵. Cysteine is an important amino acid sensitive to redox changes. Under mild oxidation, reversible oxidation of cysteine sulfhydryl groups serves as redox sensors and signal transducers that regulate a variety of physiological processes^6,7. Tandem mass tag (TMT) reagents enable concurrent identification and multiplexed quantitation of proteins in different samples using tandem mass spectrometry^8,9. The cysteine-reactive TMT (cysTMT) reagents enable selective labeling and relative quantitation of cysteine-containing peptides from up to six biological samples. Each isobaric cysTMT tag has the same nominal parent mass and is composed of a sulfhydryl-reactive group, a MS-neutral spacer arm and an MS/MS reporter¹⁰. After labeling, the samples were subject to protease digestion. The cysteine-labeled peptides were enriched using a resin containing anti-TMT antibody. During MS/MS analysis, a series of reporter ions (i.e., 126-131 Da) emerge in the low mass region, providing information on relative quantitation. The workflow is effective for reducing sample complexity, improving dynamic range and studying cysteine modifications. Here we present redox proteomic analysis of the Pst DC3000 treated tomato (Rio Grande) leaves using cysTMT technology. This high-throughput method has the potential to be applied to studying other redox-regulated physiological processes.