PTMs of proteins via mass spec?

PTMs of proteins via mass spec?

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I understand that mass spec is widely used to study PTMs like glycosylation of proteins, but how can mass spec determine correct PTM structure of say glycosylation if two glycan structures have the same mass but different arrangement (i.e. isobaric)? Also, how can mass spec determine the difference between structural isomers?

I am not a MassSpec expert but this is what I understand from my spectroscopy basics:

In tandem MS (MS-MS), the ionized analyte generally undergoes or made to undergo fragmentation (breaking up into smaller ions). Fragmentation allows you to study finer details. It is also essential for de-novo peptide sequencing. Different fragmentation techniques, such as Collision Induced Dissociation (CID) or Photofragmentation, yield different kinds of secondary ions.

You can intuitively understand that Fragmentation of structural isomers will yield different kinds of secondary ions (using the same fragmentation technique). The choice of fragmentation technique depends on the molecular nature of the analyte. In the article that I mentioned previously, they use laser induced photofragmentation to dissociate glycan-ions.

Even small molecules, which had been classically studied by MassSpec, undergo isomerization after ionization in order to stabilize the ion. Reactions such as reverse-Diels-Adler can also happen. The pattern of reactions would differ between structural isomers. For basics you can refer to a book on Spectroscopic techniques by Silverstein

Comprehensive Analysis of the Lysine Succinylome and Protein Co-modifications in Developing Rice Seeds

Lysine succinylation has been recognized as a post-translational modification (PTM) in recent years. It is plausible that succinylation may have a vaster functional impact than acetylation because of bulkier structural changes and more significant charge differences on the modified lysine residue. Currently, however, the quantity and identity of succinylated proteins and their corresponding functions in cereal plants remain largely unknown. In this study, we estimated the native succinylation occupancy on lysine was between 2% to 10% in developing rice seeds. Eight hundred fifty-four lysine succinylation sites on 347 proteins have been identified by a thorough investigation in developing rice seeds. Six motifs were revealed as preferred amino acid sequence arrangements for succinylation sites, and a noteworthy motif preference was identified in proteins associated with different biological processes, molecular functions, pathways, and domains. Remarkably, heavy succinylation was detected on major seed storage proteins, in conjunction with critical enzymes involved in central carbon metabolism and starch biosynthetic pathways for rice seed development. Meanwhile, our results showed that the modification pattern of in vitro nonenzymatically succinylated proteins was different from those of the proteins isolated from cells in Western blots, suggesting that succinylation is not generated via nonenzymatic reaction in the cells, at least not completely. Using the acylation data obtained from the same rice tissue, we mapped many sites harboring lysine succinylation, acetylation, malonylation, crotonylation, and 2-hydroxisobutyrylation in rice seed proteins. A striking number of proteins with multiple modifications were shown to be involved in critical metabolic events. Given that these modification moieties are intermediate products of multiple cellular metabolic pathways, these targeted lysine residues may mediate the crosstalk between different metabolic pathways via modifications by different moieties. Our study exhibits a platform for extensive investigation of molecular networks administrating cereal seed development and metabolism via PTMs.

Keywords: acetylation lysine succinylation mass spectrometry plant biology post-translational modifications protein modification rice seeds storage nutrient succinylome.


Peptide identification from a protein or translated genomic database is probability based. This is done by comparing the observed MS/MS spectrum with the theoretical spectra for the predicted proteolytic peptides (see Figure 6.1) of all proteins in the database. If more than a few modifications at a time are considered, the search time increases exponentially, and the probability score is decreased if more modifications are used in order to achieve a “match.”


Schematic of MS spectrum of a tryptic-digested protein.

Another factor critical to database searching is that the modification must be specific to only certain amino acids. Non-specific modifications, for example, modifications that can react with any amino acid, cannot be searched with some common database search engines, such as Mascot™ (3).

Applications of department mass spectrometry include:

Neurological proteomics: assessing protein populations and PTMs in the brain

One goal of proteomics is move beyond the reductive analysis of individually acting protein isoforms to examine how the composition of protein populations changes in response to natural stimuli and stressors, disease states, and drug therapies. Thus the aim is to conduct broadly encompassing (global) measurement of the relative quantities of the hundreds of different proteins that interact to generate biological outcomes, moving beyond the activities of individual neurotransmitter and receptor species

The synapses of the brain and central nervous system (CNS)—dynamic junctions where neurons exchange chemical signals that are the molecular underpinnings of phenomena such as learning and memory—are the subject of such quantitative and qualitative (PTM) proteomic analysis by department researchers and their neuroscientist collaborators. For example:

Tracking coordinated changes in synaptic proteome during CNS stimulation

MS combined with isotopic tagging of peptides was used to analyze variations in the relative abundance of nearly 900 proteins in mouse post-synaptic densities (PSD) at distinct time intervals in response to broad CNS drug stimulation. The density is a protein-rich structure on the receiving end (dendrite terminal) of inter-neuronal chemical communications across the synaptic cleft. PSDs contain concentrations of receptors that bind neurotransmitters as well as numerous associated synaptic signaling and regulatory proteins, assembled by a variety of scaffold proteins.

By analyzing the changes in relative quantities of the post-synaptic proteins, the study found evidence of the co-regulated activation of certain groups. Thus they were able to identify core functional complexes in which proteins displayed coordinated activity even when not known to physically interact.

Quantifying synaptic phosphorylation and protein expression by brain region

Global MS quantification and isotopic labeling was applied to examine differences in protein expression and phosphorylation at post-synaptic densities by brain region. The analysis of more than 2,100 proteins and 1,500 phosphorylation sites in a mouse model suggested roles for previously unannotated proteins whose greater expression clustered with known functional complexes.

This study also revealed relatively more phosphorylation sites on NMDA than AMPA receptors, which are both involved in synaptic plasticity—changes in the efficacy of synaptic transmissions that underlie learning, memory, and other neurological phenomena. Indeed, the study found higher average levels of enzymes associated with reversible phosphorylation (kinases, phosphatases) in the hippocampus, a region associated with memory formation and spatial orientation is among the first to suffer damage in Alzheimer’s disease.

Graph of relative abundance of a specific protein, chapsyn-110 (a membrane-associated kinase) in post-synaptic densities by brain region, showing its greatest prevalence in the cortex, mid-brain, cerebellum, and hippocampus. (Individual peptides are represented by dots, protein averages by horizontal bars.) This research was originally published in Molecular & Cellular Proteomics. Trinidad, et al., Quantitative Analysis of Synaptic Phosphorylation and Protein Expression. Molecular and Protein Expression. 2008 Vol 7:684-696

Graph shows higher relative expression of enzymes associated with reversible phosphorylation in the hippocampus. This research was originally published in Molecular & Cellular Proteomics. Trinidad et al., Quantitative Analysis of Synaptic Phosphorylation and Protein Expression. Molecular and Protein Expression. 2008 Vol 7:684-696

First large-scale mapping of challenging carbohydrate modifications in synaptic proteome

Department scientists applied liquid chromatography and electron-transfer dissociation MS to conduct the first large-scale study characterizing the specific sites and modifier structures of more than 2,500 unique N- and O-linked glycopeptides from 453 proteins in mouse synaptosomes—membrane-bound sacks of vesicles from neuronal axon terminals.

Extra-cellular glycosylation is a common, complex suite of structurally related PTMs in which one or more carbohydrates (glycans) are attached to a membrane or secreted protein. The N- or O- designation refers to the nitrogen or oxygen in amino acid (residue) side chains where the glycan is attached. These PTMs aid and stabilize protein folding among other functions.

Such analyses are challenging because glycosidic bonds are more readily broken in collision-induced dissociative processes than the peptide bonds of other covalent PTMs. Also, there is limited software for automating identification of glycan-modification spectra.

Researchers here used two forms of affinity chromatography (lectin, TiO2) to sort and enrich the concentration of glycopeptides for identification. Then, having previously identified thousands of synaptosome proteins, they focused on annotated transmembrane/secreted proteins and used the resource’s Protein Prospector to search those proteins, allowing for modifications with specific mass values corresponding to potential glycan components. They thus identified the most common carbohydrate modifications and matched their masses to sugar (oligosaccharide) structures.

Such MS analyses also identified the number of unique glycans for each N-linked glycosylation site, including a single site with 19 glycan modifications.

Pioneering the identification of sites and roles of a common, important sugar PTM: O-GlcNAcylation

O-GlcNAcylation is a common, vital, and reversible post-translational modification that occurs in all animals and plants(metazoans), in both cellular nuclei and cytosol. It entails the modification of serine and threonine residues of nuclearc and cytosolic proteins with a single oxygen-linked N-acetylglucosamine (GlcNAc). It is implicated in the modulation of:

  • gene regulation (including via modification of transcription factors)
  • responses to cellular stress and nutrient levels, including the effects of the latter on cellular circadian clocks
  • the targeting of proteins for destruction by the proteasome
  • intra-cellular signaling pathways regulated by phosphorylation (the most-studied PTM and commonly an on-off switch of proteins)—via cross-talk on co-modified residues or competition for modification sites
  • diseases such as diabetes and Alzheimer’s (enzymes that add/remove O-GlcNAc are highly expressed in the brain)

While this PTM was discovered three decades ago, detailed determination of its biological functions had been limited by the difficulty of identifying precisely which serine or threonine residues on a protein were modified. The use of collision-induced dissociation to fragment proteins into peptides for analysis typically broke the weaker sugar-oxygen glycosidic bonds before the modified protein’s peptide backbone, thus losing modification site data.

At a time when less than 80 exact residues of O-GlcNAcylation were known on all proteins, mostly via chemical dissection (Edman degradation), scientists here pioneered new methodology, combining lectin (wheat germ agglutin) affinity chromatography with electron-transfer dissociation (ETD) tandem MS. This was applied to the proteomes of mouse brain cells, where high rates of O-GlcNAcylation had been noted. Specifically, researchers analyzed post-synaptic densities—protein-rich structures at the receiving end (dendrite terminal) of inter-neuronal chemical communications across the synaptic cleft.

Upon initially applying the new method, department researchers determined 58 modification sites from a single experiment, including 28 on Bassoon—a protein in the synaptic active zone—matching the number of then-known phosphorylation sites on that protein and suggesting that O-GlcNAcylation might be play an equivalent role in regulating its function. Indeed, the study also found further evidence that the sugar modification might interact (crosstalk) with phosphorylation in the synapse, as several O-GlcNAc-modified sites on Bassoon were previously reported as phosphate modified.

Post-translational modifications on the synaptic protein Bassoon, which is comprised of 3940 amino acids from its N to C terminus. Positions of the 28 O-GlcNAC modifications (top) are indicated relative to phosphorylation sites (bottom). O-GlcNAc sites also reported as phosphorylation sites are indicated in red. From Proceedings of the National Academy of Sciences, Identification of protein O-GlcNAcylation sites using electron transfer dissociation mass spectrometry on native peptides, Chalkley et al., Vol 106 no. 22, 2009, 8894–8899

Combined global characterization of two major modifications

The potential for modulating effects, reciprocal regulation, and other interplay (crosstalk) between O-GlcNAcylation and phosphorylation is suggested by factors that include:

  • Both reversibly modify serine and threonine residue side chains
  • Both are common (most cellular proteins may be phosphorylated, while thousands of O-GlcNAcylation sites are now known)
  • In vitro modulation of global phosphorylation levels changes the O-GlcNAcylation levels of many proteins and vice versa.

To examine the interplay of these two common PTMs, researchers here developed an approach allowing both the combined detection and site determinations of O-GlcNAcylatoin and phosphorylation, in the same biological sample. This applies affinity chromatography (a weak interaction between O-GlcNAc and lectin what germ agglutinin as well as between phosphates and titanium dioxide) for the enrichment of O-GlcNAc- and phosphate-modified peptides with electron capture/transfer dissociation MS. Examples of this dual PTM characterization include:

Global identification, location of O-GlcNAcylation and phosphorylation in same synapse proteome samples

Department scientists and their collaborators applied this approach to detect both modifications and their locations on proteins in the same biological samples from mouse synaptosomes—membrane-bound sacks of vesicles from the delivering side (axon terminals) of inter-neuronal chemical communications across the synaptic cleft

The study identified more than 6000 proteins with either or both modifications and yielded estimates of 19% and 63% of the synaptosome proteins being O-GlcNAcylated or phosphorylated, respectively. It also found that proteins extensively modified by O-GlcNAc were almost always phosphorylated to a similar or greater extent, indicating that O-GlcNAc-transferase (OGT, which catalyzes the PTM), is targeting certain phosphorylated proteins. In addition, kinases (which phosphorylate proteins) were the class of proteins most extensively modified by O-GlcNAc, suggesting a form of crosstalk in which O-GlcNAcylation regulates the enzymes’ activity.

O-GlcNAcylatoin and phosphorylation interplay in circadian clock regulation

The resource provided MS analysis of the interplay between these modifications in the regulation of 24-hour cellular circadian clocks, which coordinate biological processes in organisms from bacteria to humans. The clocks’ rhythms can be affected by nutrient levels as well as by light.

Prior studies had found that O-GlcNAcylation of circadian-related transcription factors and proteins functions as a nutrient sensor and affects the clocks (altering period length in mice and fruit flies). MS analysis of proteins from mouse brains and livers by department scientists showed OGT, the enzyme that catalyzes O-GlcNAcylation, is phosphorylated and thus upregulated by a kinase (glycogen synthase Kinase 3β) but can also be O-GlcNAc modified at the same locations such that the two enzymes compete with and “fine tune” one another’s effects.

A] MS/MS spectrum of a doubly O-GlcNAc-modified region of the brain protein PER2, which regulates circadian clock speed—or time of sleep onset. B] MS/MS identified sites of O-GlcNAc modification of a kinase(CK1) binding domain (amino acids 557 to 771). The identified serines (numbers 662, 668, and 671) are also CK1 phosphorylation sites suggesting competing modifications. From Cell Metabolism, Glucose Sensor O-GlcNAcylation Coordinates with Phosphorylation to Regulate Circadian Clock, Kaasik et. al, February 2013, pp. 291-302.

The same study used tandem MS analysis to find that O-GlcNAcylation and phosphorylation are also competing modifications of a specific region of a brain protein (PER2) critical for regulating clock “speed”—a mutation reducing phosphorylation of the region leads to a syndrome of early evening sleep onset.

Deciphering epigenetics: detecting and characterizing modifications to histones and chromatin proteins (DNA)

There are roughly 20,000 protein-encoding genes in every cell in our bodies. Yet gene expression differs not only by cell type and developmental stage, but also during healthy response to stimuli and stresses and via dysfunction and disease, converting genetic make-up (genotype) to observable traits (phenotype).

One way that gene expression is regularly modified is by the attachment of chemical groups and carbohydrates to the combined complex of DNA and its associated proteins (chromatin) in the cell nucleus, leading to a physical remodeling that alters gene accessibility.

Determining how and where such modifications occur is an example of epigenetics—the study of such non-sequence-related changes in gene expression that can be inherited by subsequent generations. A key subset of epigenetic modifications is made to histones—several protein isoforms that package and stabilize DNA in a spool-like manner (a repeating basic unit called a nucleosome is a DNA segment wound around eight histone cores). Upon histones’ modification by multiple chemical groups (e.g., acetyl, methyl, ubiquitin, phosphate), typically on their unstructured N-terminals (dubbed “tails”), they remodel part of the chromatin, regulating the binding affinities of transcriptional proteins and ultimately altering gene expression.

MS/MS spectrum of a histone (H4) peptide (GKGGKGLGKGGAKR) from maize (aka corn) showing its post-translational modifications by acetyl (ac) groups (acetylation). M/Z indicates mass-to-charge ratios.

Department scientists and their collaborators have pioneered in applying mass spectrometry to histone modifications as well epigenetic changes such as DNA methylation. Such research could help decipher the hypothetical histone and epigenetic codes—which propose that complex combinations of multiple modifications and their interactions (cross-talk) are correlated with changes in gene expression in modulated responses to endogenous and environmental stimuli and, when disrupted, with dysfunctional expression in disease.

Histone modifications represent a challenging model for the application of mass spectrometry to analyzing intact proteins, given their complex variety of modifications. More than 60 residues on the four core nucleosome histones have been found to be subject to one or more modifications. Top-down electron transfer/capture dissociation techniques allow the analysis of larger peptides, but modifications to different sites and in different quantities (e.g., trimethylation) must be precisely detected despite varying in frequency and abundance by orders of magnitude.

Examples of department research and applications of MS methodologies into epigenetic modifications include:

Quantifying effects of carcinogenic arsenic exposure on histone modifications

Scientists here applied quantitative MS analysis of comparative histone modifications to a cell culture model for the development of bladder cancer due to arsenic exposure. While arsenic exposure, especially through contaminated groundwater, is known to be associated with a several types of cancers in humans, the underlying mechanisms have been elusive. Working with collaborators, scientists here analyzed histone modifications in human uroepithelium cells (which line the urinary tract) exposed to doses of arsenic over time. The study detected a reduction in acetylation levels on H3 and H4 histone isoforms at specific lysine residues. Noting related findings of arsenic-altered histone acetylation and malignant transformation, the study authors suggested that such epigenetic disturbances could silence key growth-controlling genes, leading to tumor development.

Assessing gene regulatory demethylation of key nucleosome sites

Using liquid chromatography-tandem MS to monitor the removal of methyl groups (demethylation) from a histone substrate over time, researchers here were able to develop a broadly applicable method to assess demethylation of specific nucleosome sites. By reconstituting the process in vitro, the study looked specifically at reaction rates (kinetics) of a histone demethylase JMJD2A catalytic domain which, among its various histone substrates, removes a third methyl group from a trimethylated H3 histone’s ninth lysine residue (H3 K9). This modification represses transcription by altering chromatin structure to inactivate genes. While trimethylation of that histone site has been seen at genes silenced in cancer, overexpression of the JMJD2 enzyme family is also implicated in disease, thus indicating the importance of precise demethylation regulation.

Mass Spectrometric Analysis of the Most Frequent Post-translational Modifications


Phosphorylation is widespread and often studied protein modification. Serine, threonine, and tyrosine residues are covalently modified by the enzymatic addition of a phosphate group. Phosphorylation is a reversible process, typically used to switch on and off various biological processes. The presence of the charged, hydrophilic group changes the structure of proteins, regulating multiple biochemical processes, for instance, cell cycle, metabolism and regulation of receptors [41, 42].

Phosphorylation of a given site typically occurs only in a (small) fraction of molecules. Phosphorylation analysis, therefore, requires both structure assignment (identification of the phosphorylation site) and determining the proportion of phosphorylated and non-phosphorylated analogs. Phosphoproteins constitute only a small fraction of the proteome, consequently enrichment strategies are essential for successful analysis. Several methods have been developed for this purpose, such as immunoaffinity chromatography, immobilized metal ion affinity chromatography (IMAC), and metal oxide affinity chromatography (MOAC) such as titanium dioxide (TiO2) and zirconium dioxide (ZrO2) chromatography [43,44,45,46]. IMAC and MOAC are based on the affinity of the phosphate groups for transition metal ions and metal oxides. However, these materials also show affinity for carboxyl groups, resulting in the isolation of acidic residues such as glutamate and aspartate-containing peptides alongside phosphorylated ones [47, 48]. Numerous strategies have been developed to overcome these specificity issues. Esterification of carboxyl groups prior to enrichment reduces the retention of acidic peptides. Organic acids having higher affinity for the stationary phase than carboxyl groups but lower than phosphate groups are employed as non-phosphopeptide excluders. Phosphopeptides can also be specifically eluted from the IMAC material using beta elimination, which removes the bond between the peptide and the phosphate group [48]. Although MOAC is shown to be more resistant to interfering compounds (e.g. salts, detergents) and more selective than IMAC in many cases, multiply phosphorylated residues may bind with high affinity to the metal oxide, which makes them dramatically difficult to elute. IMAC, therefore, provides improved coverage of multiphosphorylated peptides, and it is also able to enrich full-length phosphoproteins without any prior proteolytic digestion [48,49,50]. Immunoaffinity chromatography is predominantly employed in the enrichment of proteins phosphorylated on the tyrosine residue [51]. Considering that the data obtained with different enrichment methods only partly overlap each other (around 30%), combination of multiple methods is recommended to improve phosphoproteome coverage [52]. The enriched samples can be subsequently fractionated prior to MS analysis by multiple methods such as hydrophilic interaction chromatography (HILIC), RP-HPLC, and SCX [53,54,55].

There are two important issues that make phosphorylated peptide characterization by MS nontrivial. First, the sensitivity of a phosphopeptide is significantly lower than that of its non-phosphorylated analog [56]. Second, phosphopeptides typically fragment in CID by losing phosphoric acid, and therefore, identification of the phosphorylation site may be compromised [43]. Using ECD, ETD and/or special MS/MS techniques (such as MS 3 ) may alleviate this problem and allow both sequence analysis and locating the phosphorylation site [57]. Phosphopeptide analysis may be performed using derivatization as well. Phosphoserine and phosphothreonine residues are selectively transformed into lysine analog and subsequent proteolytic cleavage (using, e.g. trypsin or Lys-C) results in novel peptides [58]. Identifying these peptides using MS/MS and comparing it to the MS/MS of non-derivatized analog can be used to identify the phosphorylation site [46, 58].


Over the last few years, study of protein acetylation has become a major issue in PTM analysis. Proteins can be acetylated at the N terminus and at the ε-amino group of lysine residues, the latter having the greater biological relevance. Acetylation has been most extensively characterized for the regulation of histones, concluding that it plays an important role in cell cycle processes, such as gene expression and DNA repair [59, 60].

Since acetylation usually occurs at a very low stoichiometry, enrichment of the modified proteins and peptides is of critical relevance. Acetyllysine-specific antibodies are highly valuable for this purpose, and used as the initial purification step of very complex samples [61,62,63]. However, the method works properly only for peptides, as the accessibility of acetylated amino-acid residues is limited in the case of intact proteins. Moreover, there is no available antibody for the enrichment of N-terminal acetylation [64]. A number of methods are accessible for the prefractionation of histones including size exclusion chromatography (SEC), SDS-PAGE, and CE [65,66,67]. Acetylation increases the hydrophobic character of proteins and removes the positive charge from the N-terminal amino group. Acetylated residues, therefore, can be separated from their non-acetylated counterparts based on hydrophobicity (RP-HPLC), electrostatic interactions (SCX), or both (zwitterionic hydrophilic interaction liquid chromatography, ZIC-HILIC) [64, 68]. HILIC also has the added value of being able to separate methylated and acetylated histones, significantly decreasing sample complexity [69]. Another possibility for reducing sample complexity is derivatization by propionic anhydride, which converts lysine residues and the N-terminal amino group into propionyl amides providing a + 56 Da mass shift and protection from enzymatic digestion [37, 70].

Lysine acetylation is considered to be a stable PTM under mass spectrometry analysis, i.e. the functional group is retained on the protein using high-energy CID [23]. Acetylation can be identified through a 42.01 Da mass shift, in contrast with the unmodified variant. Although another modification called trimethylated lysine has a very similar weight (42.04 Da) to lysine acetylation, high-resolution mass spectrometers are capable to differentiate such modifications [71]. Since acetylation neutralizes the positive charge of lysine residue and blocks the tryptic cleavage, the absence of this characteristic cleavage can be another sign of acetylation [72].

O/N Glycosylation

Approximately, more than the half of human proteins bears simple or complex glycan side-chains, influencing protein folding and stability [73, 74]. There are two major types of glycan side-chains: (1) O-linked glycans attached to the hydroxy group of serine, threonine, tyrosine, while, (2) N-linked glycans are attached to the nitrogen of asparagine side-chains. In contrast to the synthesis of nucleic acids and proteins, glycosylation processes are not template directed, providing great heterogeneity for glycoconjugates. Consequently, a glycoprotein is a mixture of protein isoforms (glycoforms), instead of being a well-defined, exact chemical entity [75].

Complete characterization of glycoproteins and glycopeptides requires multiple analytical methods. In one approach, glycan side-chains are released from the protein by the application of an enzyme (e.g. peptide N-glycosidase F, PNGase F) or by reductive elimination. The former can be used in the case of N-glycans, and the latter for O-glycans [76,77,78,79]. When the sugars are released, they are typically separated from the protein mixture and derivatized. Replacing all reactive hydrogens with methyl groups (permethylation) is the most widespread method for this purpose, which enables detailed structural determination of complex glycans (e.g. branching sites, interglycosidic linkages, configurational isomers) [82]. As the procedure stabilizes sialic acid residues, permethylated derivatives show more effective ionization and more predictable fragmentation properties compared to their native counterparts [80,81,82,83,84]. Permethylated glycans are most often analyzed by MALDI-MS. Anthranilic acid (2-aminobenzoic acid, 2-AA) labeling of reducing carbohydrates is carried out in mild conditions, therefore, undesirable desialylation can be avoided [85]. 2-AA tag improves both chromatographic and mass spectrometric (in negative ion mode) detectabilities of glycans by acting as a chromophore, fluorophore and adding a negative charge to all the molecules. It also provides more informative fragments facilitating sequential analysis of saccharides [81, 85,86,87,88]. The derivatives can be subsequently separated by a number of methods, such as HILIC and CE prior to mass spectrometric analysis. CE is capable of separating positional isomers as well as differently linked glycans with high resolution and short analysis time, while HILIC tends to yield better retention time repeatability [89, 90]. It is also demonstrated that the techniques show notable complementarity in the glycoform profiling of therapeutic proteins [91].

An alternative to release glycan analysis is digestion of the protein (mixture) with proteolytic enzymes (e.g. trypsin). This yields a mixture of peptides and glycopeptides, and can be analyzed by CID- or ECD-based tandem mass spectrometry. Enzymatic digestion is often followed by enrichment techniques prior to the MS analysis. Glycopeptides may be enriched by boronate affinity chromatography, lectin affinity chromatography, HILIC, and electrostatic repulsion-hydrophilic interaction chromatography (ERLIC) [75, 92,93,94,95,96]. Boronate affinity chromatography is able to capture glycan moieties through formation of borate diesters with vicinal diols of the sugar residues. This ensures selective enrichment of glycopeptides, even in complex mixtures. In contrast to boronate affinity chromatography, lectin affinity chromatography allows the enrichment of unique glycoproteins/peptides having specific glycan structure, which may be exceptionally useful in the exploration of disease-related, abnormal glycosylation pattern [97]. As glycosylation increases hydrophilicity of proteins and peptides, glycopeptides can be easily separated from non-glycosylated ones by HILIC. ERLIC, in which positively charged functional groups are attached to the stationary phase, can be considered as a combination of hydrophilic interaction and ion-exchange chromatography. Sialylated moieties with negative charge are, therefore, highly retained, while positively charged peptides are easily eluted [98]. It is also reported that certain gradient HILIC methods can be converted into isocratic separations by ERLIC [99].

There are various mass spectrometric approaches for glycopeptide analysis, identifying the glycan structure, the glycosylation site and also the abundance ratio of various glycoforms. CID fragmentation often cleaves off the sugar residue, while ECD and ETD can yield information on the peptide sequence [75]. Ion mobility–mass spectrometry (IM-MS) is also a promising analytical tool in this field with the added value of being able to separate analytes based on their size. This technique has been successfully applied for the separation and characterization of isobaric polysaccharides and glycopeptides [100, 101].


Approximately, half of the endocrine and neural peptides bear C-terminal amidation [102]. The presence of such modification is indispensable for signal transduction and receptor recognition [102, 103]. Furthermore, derivatization of amidated peptides with glycosylphosphatidylinositol is responsible for anchoring peptides and proteins to the cell membrane [104].

As in most cases in the field of PTMs, proteins that are presumably amidated at the C terminus can be separated or enriched by specifically engineered approaches based on the affinity of such proteins. For a long time, only radioimmunoassay (RIA) and immunoprecipitation (IP) have been applied for identification of amidated proteins [105, 106]. Separation of amidated peptides from their precursors can be carried out with RP-HPLC [105]. Weak cation-exchange chromatography (WCX) has been reported to be able to separate the α-amidated heavy chain of immunoglobulin G1 from other isoforms [107].

Amidation of a carboxyl group manifests in a 1 Da mass shift, which is very difficult to identify. The situation can be further complicated when the C-terminal amino acid is glutamine or asparagine, which cannot be distinguished from the amidated form of glutamate or aspartate, except when an auxiliary chemical or enzymatic procedure is performed. An approach has been developed for detecting C-terminal amidation by a combination of chemical derivatization and MALDI-TOF/TOF MS [86]. First step is the derivatization of the free carboxyl group at the C terminal, resulting in a methylamide structure [108, 109]. This results in a 13 Da mass increment observable in the spectrum, enabling the distinction between the amidated and the unmodified peptides. This method was shown to be able to distinguish isobaric residues at the C terminal, such as Gln-OH and Glu-NH2 or Asn-OH and Asp-NH2, suggesting a wider application to investigate modified and unmodified C terminals of proteins [104].


Although hydroxylation can take place on several amino acids including lysine, histidine, asparagine, aspartate, and aromatic residues, the most commonly hydroxylated amino acid is proline, which makes up almost one-third of collagen protein [110,111,112]. Hydroxylation generally takes place at the γ-C atom, yielding 4-hydroxyproline (4-Hyp), which has the ability to stabilize the secondary structure of the protein by the powerful electronegative effect of the hydroxy group [112]. Hyp is also associated with the decreased availability of oxygen in the cellular environment, as the alpha subunits of hypoxia-induced factor (HIF) contain hydroxylated proline residues [113].

Mass spectrometric investigation of collagenous proteins is intricate due to the large number of proline residues and the high abundance of hydroxylation. These structural features result in a vast number of isobaric but differently modified peptides in a typical proteomic workflow, which usually co-elute during chromatographic separation and provide chimeric spectra troublesome to elucidate. Moreover, the nominal mass of Hyp is 113 Da, which is the same as leucine and isoleucine. To overcome such difficulties, a mass spectrometer with high mass accuracy and abundant product ions from the tandem MS experiment are required [114, 115]. Unfortunately, MS n sequence analysis is ruined by the abnormal fragmentation of Pro- and Hyp-rich proteins and peptides. These residues promote characteristic and dominant cleavages instead of those nonselective ones, which may provide sequence information. This phenomenon is the so-called “proline-effect”, by which the identification of collagens and other proline-rich proteins are greatly hampered [114, 116, 117]. An approach has been suggested that is based on the application of five different proteolytic enzymes to increase sequence coverage of collagenous proteins. Thereafter, the analytes were separated by nano-LC and introduced into a linear ion trap–orbitrap instrument. Differently modified peptides with the same sequence were eluted and ionized together producing chimeric mass spectra. As ETD-induced fragmentation did not yield sufficient amount of data, additional dissociation techniques (CID and HCD) were also applied. In general, multistage activation made the identification of PTM-carrying residues possible, however, in some cases, the exact localization of the modification has remained unknown [114].

Table 2 summarizes the separation/enrichment techniques, fragmentation methods, and commonly used derivatization methods of modified peptides and proteins.


Csizmok, V. & Forman-Kay, J. D. Complex regulatory mechanisms mediated by the interplay of multiple post-translational modifications. Current opinion in structural biology 48, 58–67 (2018).

Santos, A. L. & Lindner, A. B. Protein posttranslational modifications: Roles in aging and age-related disease. Oxidative Medicine and Cellular Longevity 2017 (2017).

Ubersax, J. A. & Ferrell, J. E. Mechanisms of specificity in protein phosphorylation. Nature reviews. Molecular cell biology 8, 530–541 (2007).

Tsiatsiani, L. & Heck, A. J. R. Proteomics beyond trypsin. The FEBS journal 282, 2612–2626 (2015).

Swaney, D. L., Wenger, C. D. & Coon, J. J. Value of using multiple proteases for large-scale mass spectrometry-based proteomics. Journal of proteome research 9, 1323–1329 (2010).

Giansanti, P., Tsiatsiani, L., Low, T. Y. & Heck, A. J. R. Six alternative proteases for mass spectrometry-based proteomics beyond trypsin. Nature protocols 11, 993–1006 (2016).

Casanovas, A., Gallardo, O., Carrascal, M. & Abian, J. Tcellxtalk facilitates the detection of co-modified peptides for the study of protein post-translational modification cross-talk in t cells. Bioinformatics (Oxford, England) (2018).

Liu, Y., Wang, M., Xi, J., Luo, F. & Li, A. Ptm-ssmp: A web server for predicting different types of post-translational modification sites using novel site-specific modification profile. International journal of biological sciences 14, 946–956 (2018).

Li, F. et al. Quokka: a comprehensive tool for rapid and accurate prediction of kinase family-specific phosphorylation sites in the human proteome. Bioinformatics (Oxford, England) 34, 4223–4231 (2018).

Li, G. X. H., Vogel, C. & Choi, H. Ptmscape: an open source tool to predict generic post-translational modifications and map modification crosstalk in protein domains and biological processes. Molecular omics 14, 197–209 (2018).

Patrick, R., Lê Cao, K.-A., Kobe, B. & Bodén, M. Phosphopick: modelling cellular context to map kinase-substrate phosphorylation events. Bioinformatics (Oxford, England) 31, 382–389 (2015).

He, W., Wei, L. & Zou, Q. Research progress in protein posttranslational modification site prediction. Briefings in functional genomics (2018).

Chen, Z. et al. Large-scale comparative assessment of computational predictors for lysine post-translational modification sites. Briefings in bioinformatics (2018).

Xu, Y., Yang, Y., Wang, Z., Li, C. & Shao, Y. A systematic review on posttranslational modification in proteins: Feature construction, algorithm and webserver. Protein and peptide letters 25, 807–814 (2018).

Wang, D., Liang, Y. & Xu, D. Capsule network for protein post-translational modification site prediction. Bioinformatics ( Oxford, England ) (2018).

Eisenhaber, B. & Eisenhaber, F. Prediction of posttranslational modification of proteins from their amino acid sequence. Methods in molecular biology (Clifton, N.J.) 609, 365–384 (2010).

Hui, E. et al. T cell costimulatory receptor cd28 is a primary target for pd-1-mediated inhibition. Science (New York, N.Y.) 355, 1428–1433 (2017).

Venne, A. S., Kollipara, L. & Zahedi, R. P. The next level of complexity: crosstalk of posttranslational modifications. Proteomics 14, 513–524 (2014).

Wiese, H. et al. Comparison of alternative ms/ms and bioinformatics approaches for confident phosphorylation site localization. Journal of proteome research 13, 1128–1137 (2014).

Collins, M. O., Wright, J. C., Jones, M., Rayner, J. C. & Choudhary, J. S. Confident and sensitive phosphoproteomics using combinations of collision induced dissociation and electron transfer dissociation. Journal of proteomics 103, 1–14 (2014).

Imanishi, S. Y. et al. Reference-facilitated phosphoproteomics: fast and reliable phosphopeptide validation by microlc-esi-q-tof ms/ms. Molecular & cellular proteomics: MCP 6, 1380–1391 (2007).

Consortium, T. U. Uniprot: the universal protein knowledgebase. Nucleic acids research 45, D158–D169 (2017).

Hornbeck, P. V. et al. Phosphositeplus, 2014: mutations, ptms and recalibrations. Nucleic acids research 43, D512–D520 (2015).

Bezanson, J., Edelman, A., Karpinski, S. & Shah, V. B. Julia: A fresh approach to numerical computing. SIAM Review 59, 65–98, (2017).

Pont, F. & Fournié, J. J. Sorting protein lists with nwcompare: A simple and fast algorithm for n-way comparison of proteomic data files. Proteomics 10, 1091–1094 (2010).

A High Resolution Mass Spectrometry Study Reveals the Potential of Disulfide Formation in Human Mitochondrial Voltage-Dependent Anion Selective Channel Isoforms (hVDACs)

The voltage-dependent anion-selective channels (VDACs), which are also known as eukaryotic porins, are pore-forming proteins, which allow for the passage of ions and small molecules across the outer mitochondrial membrane (OMM). They are involved in complex interactions that regulate organelle and cellular metabolism. We have recently reported the post-translational modifications (PTMs) of the three VDAC isoforms purified from rat liver mitochondria (rVDACs), showing, for the first time, the over-oxidation of the cysteine residues as an exclusive feature of VDACs. Noteworthy, this peculiar PTM is not detectable in other integral membrane mitochondrial proteins, as defined by their elution at low salt concentration by a hydroxyapatite column. In this study, the association of tryptic and chymotryptic proteolysis with UHPLC/High Resolution nESI-MS/MS, allowed for us to extend the investigation to the human VDACs. The over-oxidation of the cysteine residues, essentially irreversible in cell conditions, was as also certained in VDAC isoforms from human cells. In human VDAC2 and 3 isoforms the permanently reduced state of a cluster of close cysteines indicates the possibility that disulfide bridges are formed in the proteins. Importantly, the detailed oxidative PTMs that are found in human VDACs confirm and sustain our previous findings in rat tissues, claiming for a predictable characterization that has to be conveyed in the functional role of VDAC proteins within the cell. Data are available via ProteomeXchange with identifier PXD017482.

Keywords: Cysteine over-oxidation Orbitrap Fusion Tribrid hydroxyapatite mitochondria mitochondrial intermembrane space outer mitochondrial membrane post-translational modification.

Conflict of interest statement

The authors declare no conflict of interest.


Sequence coverage map of hVDAC1…

Sequence coverage map of hVDAC1 obtained by tryptic and chymotryptic digestion. The gray…

MS/MS spectrum of the doubly…

MS/MS spectrum of the doubly charged molecular ion at m/z 1083.9756 (calculated 1083.9754)…

MS/MS spectrum of the triply…

MS/MS spectrum of the triply charged molecular ion at m/z 812.0634 (calculated 812.0635)…

Sequence coverage map of hVDAC2…

Sequence coverage map of hVDAC2 obtained by tryptic and chymotryptic digestion. Gray solid…

Sequence coverage map of hVDAC3…

Sequence coverage map of hVDAC3 obtained by tryptic and chymotryptic digestion. The gray…

Scheme of the cysteine localization…

Scheme of the cysteine localization in the aligned sequences of human VDAC isoforms.…

The basics of mass spectrometry

Since their inception in 1912, mass spectrometers have undergone continuous development, and these sophisticated bioanalytical instruments have now reached unrivalled detection limits, speed and diversity in applications. They detect the presence and abundance of peptides (or other biomolecules such as metabolites, lipids and proteins) using fundamental properties of molecules, such as mass, and net charge. When peptides obtain a net charge (usually through gain of protons), they are referred to as peptide ions.

All mass spectrometers have three fundamental components: an ion source, mass analyser and detector (Figure 1A). As mass spectrometers can only analyse gaseous ions, methods such as electrospray ionization (ESI) are needed to convert peptides from the liquid phase to gaseous ions. The liquid containing the peptides is pumped through a micrometre-sized orifice held at a high voltage (2–4 kV). Upon reaching this emitter, the steady stream of liquid disintegrates into extremely small, highly charged and rapidly evaporating charged droplets, leaving peptide ions in the gas phase. Even 20 years after John Fenn received the Nobel Prize for this discovery, the exact mechanisms are not completely understood. We know that the abundance of gaseous peptide ions is proportional to their original concentration, making it beneficial to use the lowest flow rates possible, thereby maximizing sensitivity. It is common in proteomics to separate peptide mixtures using high-performance liquid chromatography (HPLC) systems with flow rates of only a few hundred nanolitres per minute rather than millilitres in conventional HPLC.

Overview of sample preparation and instrumentation used in MS-based proteomics. (A) Proteins are digested into peptides using sequence-specific proteases. Optionally, post-translational modification (PTM)-containing peptides can be enriched using beads with specific surface chemistry or coupled antibodies. High-performance liquid chromatography (HPLC) separates peptides based on hydrophobicity, and they are subsequently analysed by a TOF mass spectrometer. (B) Alternatively, peptides can be analysed by an Orbitrap mass spectrometer, which is a mainstream instrument in proteomics.

The principal role of a mass analyser is to separate ions by their mass-to-charge ratios (m/z). Fundamentally, all ions are separated by modulating their trajectories in electrical fields. Mass analysers differ in the principle they use for separating ions, and this defines their preferred application areas. Quadrupoles, usually combined with time-of-flight (TOF) or Orbitrap analysers, are the most common in proteomics. Quadrupole mass analysers separate ions using an oscillating electrical field between four cylindrical rods in a parallel arrangement, where each pair of rods produces a radio frequency electrical field with a phase offset. The resulting electrical fields define a pseudo-potential surface that is configured to allow the transmission of all ions, or to selectively transmit ions of a specific m/z window.

TOF mass analysers separate ions based on the differences in velocities after acceleration to about 20 kV and subsequent different arrival times at the detector. A TOF can measure mass differences of one part per million (ppm) by detecting time differences of sub-microseconds. In contrast, the Orbitrap mass analyser distinguishes ions based on their oscillation frequencies. Ions are tangentially injected and then trapped in the Orbitrap, and they move along the length axis of a central metal spindle (Figure 1B). Although an Orbitrap is only a few centimetres long, the ions can rapidly travel up to several kilometres, enabling very high resolution (typically tens of thousands) and low ppm mass accuracy.

In proteomics, the quadrupole element is normally followed by a ‘collision cell’, which is a quadrupole where the ions can be fragmented. Either intact peptide ions or fragment ions enter the final stage that also contains the detector – the resulting spectra are called MS 1 or precursor ion spectra in the former case and MS 2 or product or MS/MS spectra in the latter. TOF instruments have microchannel plate (MCP) detectors, where each individual ion ejects electrons from a surface that are then amplified. Individual ions can be readily measured with MCPs, but this exquisite sensitivity comes with the caveat that the detector can easily saturate in case of high signals. In Orbitrap analysers, the ‘image current’ induced by the rapidly oscillating ions is measured, and it represents a quantitative readout of the strength of the individual ion packages. The current is recorded in the time domain and is converted into the frequency domain using Fourier transformation. Advances in signal processing algorithms have repeatedly doubled the achievable resolution with a given transient time of the signal, but these are still orders of magnitude slower than those of TOF analysers (tens to hundreds of milliseconds vs typically 100 microseconds for a single TOF pulse).

How do the MS instruments sequence or identify peptides? Precursor ions with a specific m/z are first isolated by the quadrupole and fragmented through collision with inert gases such as N2, He or Ar. This causes them to break apart at the lowest energy bonds – typically, some of the amide bonds (peptide bonds) connecting the amino acid residues – and leaves MS/MS spectra with incomplete ladders of peaks differing by amino acid masses. This information is incredibly specific and is used for identification of the peptide sequence. A sequence of just a few amino acids and the flanking masses – a peptide sequence tag – is sufficient for identifying a peptide from the entirety of human proteome. More usually, database identification involves generating all possible fragmentation spectra and then statistically scoring them against the experimental spectra.

The chromatographic retention time is an important level of information when matching a dataset against a previous measurement and is key to ‘targeted proteomics’ technologies. Furthermore, ion mobility analysis, an additional dimension of separation of peptide ions, has recently become mainstream. Ions are either filtered based on their cross-section (FAIMS, field asymmetric ion mobility spectrometry) or actually separated during their analysis (T-Wave or TIMS, trapped ion mobility spectrometry). TIMS is the basis of the parallel accumulation–serial fragmentation (PASEF) technology, which multiplies sequencing speed 10-fold while improving sensitivity.


Protozoan parasitic infections are health, social and economic issues impacting both humans and animals, with significant morbidity and mortality worldwide. Protozoan parasites have complicated life cycles with both intracellular and extracellular forms. As a consequence, protozoan adapt to changing environments in part through a dynamic enzyme-catalyzed process leading to reversible posttranslational modifications (PTMs). The characterization by proteomics approaches reveals the critical role of the PTMs of the proteins involved in host-pathogen interaction. The complexity of PTMs characterization is increased by the high diversity, stoichiometry, dynamic and also co-existence of several PTMs in the same moieties which crosstalk between them. Here, we review how to understand the complexity and the essential role of PTMs crosstalk in order to provide a new hallmark for vaccines developments, immunotherapies and personalized medicine. In addition, the importance of these motifs in the biology and biological cycle of kinetoplastid parasites is highlighted with key examples showing the potential to act as targets against protozoan diseases.

Cohen, P. The regulation of protein function by multisite phosphorylation—a 25 year update. Trends Biochem. Sci. 25, 596–601 (2000).

Tyers, M. & Jorgensen, P. Proteolysis and the cell cycle: with this RING I do thee destroy. Curr. Opin. Genet. Dev. 10, 54–64 (2000).

Carr, S.A. et al. Protein and carbohydrate structural analysis of a recombinant soluble CD4 receptor by mass spectrometry. J. Biol. Chem. 264, 21286–21295 (1989).

Ling, V. et al. Characterization of the tryptic map of recombinant DNA-derived tissue plasminogen activator by high-performance liquid chromatography–electrospray ionization mass spectrometry. Anal. Chem. 63, 2909–2915 (1991).

Roepstorff, P. Mass spectrometry in protein studies from genome to function. Curr. Opin. Biotechnol. 8, 6–13 (1997).

Aebersold, R. & Mann, M. Mass spectrometry-based proteomics. Nature, in press (2003).

Jensen, O.N. Modification-specific proteomics: strategies for systematic studies of post-translationally modified proteins. in Proteomics: A Trends Guide (eds. Blackstock, W. & Mann, M.) 36–42 (Elsevier Science, London 2000).

McLachlin, D.T. & Chait, B.T. Analysis of phosphorylated proteins and peptides by mass spectrometry. Curr. Opin. Chem. Biol. 5, 591–602 (2001).

Sickmann, A. & Meyer, H.E. Phosphoamino acid analysis. Proteomics 1, 200–206 (2001).

Mann, M. et al. Analysis of protein phosphorylation using mass spectrometry: deciphering the phosphoproteome. Trends Biotechnol. 20, 261–268 (2002).

Loughrey Chen, S. et al. Mass spectrometry-based methods for phosphorylation site mapping of hyperphosphorylated proteins applied to Net1, a regulator of exit from mitosis in yeast. Mol. Cell Proteomics 1, 186–196 (2002).

Gorg, A. et al. The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 21, 1037–1053 (2000).

Rabilloud, T. Two-dimensional gel electrophoresis in proteomics: old, old fashioned, but it still climbs up the mountains. Proteomics 2, 3–10 (2002).

Fey, S.J. & Larsen, P.M. 2D or not 2D. Two-dimensional gel electrophoresis. Curr. Opin. Chem. Biol. 5, 26–33 (2001).

Larsen, M.R., Larsen, P.M., Fey, S.J. & Roepstorff, P. Characterization of differently processed forms of enolase 2 from Saccharomyces cerevisiae by two-dimensional gel electrophoresis and mass spectrometry. Electrophoresis 22, 566–575 (2001).

Knebel, A., Morrice, N. & Cohen, P. A novel method to identify protein kinase substrates: eEF2 kinase is phosphorylated and inhibited by SAPK4/p38δ. EMBO J. 20, 4360–4369 (2001).

MacDonald, J.A., Mackey, A.J., Pearson, W.R. & Haystead, T.A. A strategy for the rapid identification of phosphorylation sites in the phosphoproteome. Mol. Cell Proteomics 1, 314–322 (2002).

Covey, T.R., Shushan, B.I., Bonner, R., Schroder, W. & Hucho, F. LC/MS and LC/MS/MS screening for the sites of post-translational modifications in proteins. in Methods in Protein Sequence Analysis (eds Jörnvall, H., Höög, J.O. & Gustavsson, A.M.). 249–256 (Birkhäuser Verlag, Basel, 1991).

Bateman, R.H. et al. A novel precursor ion discovery method on a hybrid quadrupole orthogonal acceleration time-of-flight (Q-TOF) mass spectrometer for studying protein phosphorylation. J. Am. Soc. Mass Spectrom. 13, 792–803 (2002).

Wilm, M., Neubauer, G. & Mann, M. Parent ion scans of unseparated peptide mixtures. Anal. Chem. 68, 527–533 (1996).

Carr, S.A., Huddleston, M.J. & Annan, R.S. Selective detection and sequencing of phosphopeptides at the femtomole level by mass spectrometry. Anal. Biochem. 239, 180–192 (1996).

Steen, H., Kuster, B., Fernandez, M., Pandey, A. & Mann, M. Detection of tyrosine-phosphorylated peptides by precursor ion scanning quadrupole TOF mass spectrometry in positive ion mode. Anal. Chem. 73, 1440–1448 (2001).

Steen, H., Kuster, B., Fernandez, M., Pandey, A. & Mann, M. Tyrosine phosphorylation mapping of the epidermal growth factor receptor signaling pathway. J. Biol. Chem. 277, 1031–1039 (2002).

Hinsby, A.M., Olsen, J.V., Bennett, K.L. & Mann, M. Signaling initiated by overexpression of the fibroblast growth factor receptor-1 investigated by mass spectrometry. Mol. Cell. Proteomics, in press (2003).

Mann, M. & Wilm, M.S. Error-tolerant identification of peptides in sequence databases by peptide sequence tags. Anal. Chem. 66, 4390–4399 (1994).

Eng, J.K., McCormack, A.L. & Yates, J.R.I. An approach to correlate MS/MS data to amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 5, 976–989 (1994).

Perkins, D.N., Pappin, D.J., Creasy, D.M. & Cottrell, J.S. Probability-based protein identification by searching sequence databases using mass spectrometry data. Electrophoresis 20, 3551–3567 (1999).

MacCoss, M.J., Wu, C.C. & Yates, J.R. Probability-based validation of protein identifications using a modified SEQUEST algorithm. Anal. Chem. 74, 5593–5599 (2002).

Creasy, D.M. & Cottrell, J.S. Error-tolerant searching of uninterpreted tandem mass spectrometry data. Proteomics 2, 1426–1434 (2002).

Marshall, A.G., Hendrickson, C.L. & Jackson, G.S. Fourier transform ion cyclotron resonance mass spectrometry: a primer. Mass Spectrom. Rev. 17, 1–35 (1998).

Martin, S.E., Shabanowitz, J., Hunt, D.F. & Marto, J.A. Subfemtomole MS and MS/MS peptide sequence analysis using nano-HPLC micro-ESI Fourier transform ion cyclotron resonance mass spectrometry. Anal. Chem. 72, 4266–4274 (2000).

Zubarev, R.A. et al. Electron-capture dissociation for structural characterization of multiply charged protein cations. Anal. Chem. 72, 563–573 (2000).

Stensballe, A., Jensen, O.N., Olsen, J.V., Haselmann, K.F. & Zubarev, R.A. Electron-capture dissociation of singly and multiply phosphorylated peptides. Rapid Commun. Mass Spectrom. 14, 1793–1800 (2000).

Shi, S.D. et al. Phosphopeptide/phosphoprotein mapping by electron-capture dissociation mass spectrometry. Anal. Chem. 73, 19–22 (2001).

Kelleher, N.L. et al. Localization of labile posttranslational modifications by electron-capture dissociation: the case of γ-carboxyglutamic acid. Anal. Chem. 71, 4250–4253 (1999).

Kjeldsen, F., Haselmann, K.F., Budnik, B.A., Soerensen, E.S. & Zubarev, R.A. Complete characterization of post-translational modification sites in the bovine milk protein PP3 by tandem mass spectrometry with electron-capture dissociation as the last stage. Anal. Chem., in press (2003).

Sze, S.K., Ge, Y., Oh, H. & McLafferty, F.W. Top-down mass spectrometry of a 29-kDa protein for characterization of any posttranslational modification to within one residue. Proc. Natl. Acad. Sci. USA 99, 1774–1779 (2002).

Soskic, V., Gorlach, M., Poznanovic, S., Boehmer, F.D. & Godovac-Zimmermann, J. Functional proteomics analysis of signal transduction pathways of the platelet-derived growth factor-β receptor. Biochemistry 38, 1757–1764 (1999).

Yamagata, A. et al. Mapping of phosphorylated proteins on two-dimensional polyacrylamide gels using protein phosphatase. Proteomics 2, 1267–1276 (2002).

Stensballe, A., Andersen, S. & Jensen, O.N. Characterization of phosphoproteins from electrophoretic gels by nanoscale Fe(iii) affinity chromatography with off-line mass spectrometry analysis. Proteomics 1, 207–222 (2001).

Pandey, A. et al. Analysis of receptor signaling pathways by mass spectrometry: identification of Vav-2 as a substrate of the epidermal and platelet-derived growth factor receptors. Proc. Natl. Acad. Sci. USA 97, 179–184 (2000).

Gronborg, M. et al. A mass spectrometry-based proteomic approach for identification of serine/threonine-phosphorylated proteins by enrichment with phospho-specific antibodies: identification of a novel protein, Frigg, as a protein kinase A substrate. Mol. Cell. Proteomics 1, 517–527 (2002).

Peng, J. & Gygi, S.P. Proteomics: the move to mixtures. J. Mass Spectrom. 36, 1083–1091 (2001).

MacCoss, M.J. et al. Shotgun identification of protein modifications from protein complexes and lens tissue. Proc. Natl. Acad. Sci. USA 99, 7900–7905 (2002).

Ho, Y. et al. Systematic identification of protein complexes in Saccharomyces cerevisiae by mass spectrometry. Nature 415, 180–183 (2002).

Gavin, A.C. et al. Functional organization of the yeast proteome by systematic analysis of protein complexes. Nature 415, 141–147 (2002).

Andersson, L. & Porath, J. Isolation of phosphoproteins by immobilized metal (Fe 3+ ) affinity chromatography. Anal. Biochem. 154, 250–254 (1986).

Posewitz, M.C. & Tempst, P. Immobilized gallium(iii) affinity chromatography of phosphopeptides. Anal. Chem. 71, 2883–2892 (1999).

Ficarro, S.B. et al. Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nat. Biotechnol. 20, 301–305 (2002).

Salomon, A.R. et al. Profiling of tyrosine-phosphorylation pathways in human cells using mass spectrometry. Proc. Natl. Acad. Sci. USA 100, 443–448 (2003).

Zhou, H., Watts, J.D. & Aebersold, R. A systematic approach to the analysis of protein phosphorylation. Nat. Biotechnol. 19, 375–378 (2001).

Oda, Y., Nagasu, T. & Chait, B.T. Enrichment analysis of phosphorylated proteins as a tool for probing the phosphoproteome. Nat. Biotechnol. 19, 379–382 (2001).

Steen, H. & Mann, M. A new derivatization strategy for the analysis of phosphopeptides by precursor ion scanning in positive ion mode. J. Am. Soc. Mass Spectrom. 13, 996–1003 (2002).

Medzihradszky, K.F. et al. The characteristics of peptide collision-induced dissociation using a high-performance MALDI-TOF/TOF tandem mass spectrometer. Anal. Chem. 72, 552–558 (2000).

Gygi, S.P. et al. Quantitative analysis of complex protein mixtures using isotope-coded affinity tags. Nat. Biotechnol. 17, 994–999 (1999).

Oda, Y., Huang, K., Cross, F.R., Cowburn, D. & Chait, B.T. Accurate quantitation of protein expression and site-specific phosphorylation. Proc. Natl Acad. Sci. USA 96, 6591–6596 (1999).

Stemmann, O., Zou, H., Gerber, S.A., Gygi, S.P. & Kirschner, M.W. Dual inhibition of sister chromatid separation at metaphase. Cell 107, 715–726 (2001).

Ong, S.E., Kratchmarova, I. & Mann, M. Properties of 13 C-substituted arginine in stable isotope labeling by amino acids in cell culture (SILAC). J. Proteome Res., in press (2003).

Blagoev, B. et al. A proteomics strategy to elucidate functional protein–protein interactions applied to EGF signaling. Nat. Biotechnol. 21, 315–318 (2003).

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