Abstract

Post-translational modifications (PTMs) are integral to the regulation of protein function, characterising their role in this process is vital to understanding how cells work in both healthy and diseased states. Mass spectrometry (MS) facilitates the mass determination and sequencing of peptides, and thereby also the detection of site-specific PTMs. However, numerous challenges in this field continue to persist. The diverse chemical properties, low abundance, labile nature and instability of many PTMs, in combination with the more practical issues of compatibility with MS and bioinformatics challenges, contribute to the arduous nature of their analysis. In this review, we present an overview of the established MS-based approaches for analysing PTMs and the common complications associated with their investigation, including examples of specific challenges focusing on phosphorylation, lysine acetylation and redox modifications.

Introduction

Following translation, proteins can be modified, often either simultaneously or independently, by multiple post-translational modifications (PTMs). With over 450 forms of PTMs listed in the UniProt database [1], each influencing the structure, function, localisation or selection of interaction partner in differing ways, dependent on PTM-type and location within the protein, it is easy to see that PTMs contribute to a great deal of the diversity observed within the protein population of cells [2,3] (see Box 1). PTMs play a significant role in both the normal biological function of cells [4–7] and the development of a range of diseases including cardiovascular diseases [8], Alzheimer’s disease [9], diabetes [10], and many others. It is therefore vital that PTMs are characterised, and their impact on proteins understood in order to fully elucidate disease mechanisms and enable the development of new targeted therapies. Here we aim to present state of the art mass spectrometry (MS) approaches for the detection and quantification of PTMs, and highlight the associated challenges which continue to persist. To do this, selected examples of PTMs from both well established, specifically phosphorylation and lysine acetylation, as well as newly evolving fields of study, in this case oxidative modifications, have been selected to highlight a number of the specific challenges.

Box 1
Post-translational modifications

  • PTMs of proteins represent an important level of regulation of protein function and activity, from very rapid and transitional to stable and irreversible processes. PTMs affect structure of proteins, and function as molecular switches that regulate the interaction of proteins with other proteins, DNA, cofactors, and lipids. PTMs increase the complexity of organismal proteomes and play a role in almost all aspects of cell biology, allowing for fine-tuning of protein structure, function, and localisation.

Role of MS in the investigation of PTMs

While many technologies and approaches have been developed to study protein PTMs, MS is a highly sensitive detection method capable of both large-scale and directed detection of PTMs, providing both site-specific and quantitative information [11]. This has made it a common approach for the investigation of PTMs, but one of the major benefits of MS-based analysis is that it allows for an unbiased, large-scale characterisation of PTMs, enabling the identification of previously unanticipated modifications [12]. In more general terms, MS-based analysis is often not directed based on prior knowledge and is capable of analysing large numbers of analytes within a single experiment. While PTMs may alter the chemical properties of amino acids, all PTMs involve either the addition or removal of a chemical group, or alternatively, specific proteolytic cleavage to yield the active form of, or remove a signalling peptide from a protein (as reviewed in [13] and [14]). This can be detected by MS as a shift in the measured mass-to-charge ratio (m/z), or by the presence of a diagnostic analyte, i.e. a characteristic ion generated during MS analysis that is specific to the PTM of interest. A number of MS approaches targeting PTMs exist, these fall broadly into the categories of intact, top-down, and bottom-up [15–17] (Figure 1). Intact analysis, as the name implies, involves the analysis of intact proteins, however due to the multiple charge states and the complex isotopic distributions of proteins as a result of their size, the overlap in signals can prove difficult to interpret, this has been described and addressed in a number of articles [18–21], therefore this method is generally limited to the analysis of a single protein or very simple mixtures [22]. While this method can be useful in providing information on the overall modification state of a protein, it does not provide site-specific information. This can be overcome by a top-down approach which involves the fragmentation (MS/MS (MS2)) of intact proteins within the MS instrument [23]. Despite the top-down based-MS approach being considered a powerful tool in the characterisation of PTMs, this approach is still limited in its ability to analyse complex mixtures. It is therefore largely used for the characterisation of proteoforms for a single protein, or simple mixtures, such as protein complexes. More detailed discussions of the benefits and disadvantages of both top-down and bottom-up MS approaches have been undertaken in numerous reviews over the years, including those by Toby et al. [24], Ahlf et al. [25], and Zhou et al. [26]. Although some techniques for larger scale top-down studies do exist, such as those developed by Schmit et al. [27], by far the most common MS approach in the analysis of PTMs is the bottom-up approach, and as such techniques based around bottom-up approaches will be focussed on within this review. However, this approach also comes with its own limitations, such as limited identification of low-intensity ions, which are discussed in the relevant PTM sections. Bottom-up approaches involve the enzymatic digestion of proteins into peptides coupled with separation followed by fragmentation within the MS instrument, allowing for the peptide sequence, and the mass and localisation of PTMs to be determined (see Box 2). MS can be used for both the identification and quantitation of peptides and the PTMs that occur on them. Generally, the approach used for large-scale PTM identification is untargeted, meaning it is not directed towards particular proteins, but rather the selection of analytes for fragmentation is determined, most commonly, by the intensity of the signal [28]. For example, a ‘top 20’ approach will result in the fragmentation and subsequent sequence identification of the 20 most intense signals within an individual MS scan of analytes collected by the instrument. Alternatively, a semi-targeted or targeted MS approach for identification may be used. This involves the selection of analytes for fragmentation based on the presence of diagnostic signals indicating the presence of a PTM of interest (semi-targeted) or based on the m/z and fragmentation pattern which matches a known target (targeted). A semi-targeted approach requires an understanding of the behaviour of a particular PTM within the MS instrument, but can be used in lieu of enrichment prior to MS analysis should an enrichment method not currently be developed [29], while a targeted approach requires prior knowledge of the specific protein or peptide and its modification state, and therefore, in practice, is only really useful as a validation method rather than an investigative tool [30]. Beyond identification, MS approaches also allow for the quantification of PTMs. This can be achieved in two main ways: label-free and labelled. The label-free approach largely involves quantitation through either counting the number of times an analyte of interest is measured by the MS instrument (spectral counting) [31], or through quantifying the area of the peak generated by the analyte within the MS instrument [32]. Alternatively, a labelled approach can be used in which a mass tag is added to the proteins or peptides. This enables quantification to be performed on the MS level, with tags of known differing masses allowing the direct comparison of intensities of the analyte between samples, for example stable isotope labelling by amino acids in cell culture (SILAC) [33] or dimethyl labelling [34]. Or it can occur on the MS2 level, with tags of isobaric mass added that, upon fragmentation, release reporter masses of differing size, allowing relative quantification between samples through the comparison of the reporter ion intensities, for example isobaric tagging for relative and absolute quantitation (iTRAQ) [35] or tandem mass tagging (TMT) [36]. These methods typically quantify the change in abundance of the peptide as a whole and are not specific to the modification. But methods have also been developed which involve specific modification-based labelling, such as the ‘SNO/SOH TMT strategy’ for the direct quantification of S-nitrosylation and sulphenylation modifications using iodoTMT labelling developed by Wojdyla et al. [37] and other thiol-based labelling approaches [38,39]. Most quantitation methods used within proteomic studies are relative between samples. To enable absolute quantitation, standards can also be used, however the generation of modified peptides for absolute quantitation can be expensive, time-consuming, and the incorporation of a particular modification at a particular site can be difficult depending on the type of modification to be investigated [40–42]. Furthermore, MS can be used to calculate the relative site-occupancy of a modification (see Box 3), that is the proportion of the peptide population which is modified at a particular time within a sample by comparing the relative abundance of the modified peptide with the unmodified counterpart. This can prove difficult, however, due to the altered behaviour of modified peptides compared with their unmodified counterparts. Modified peptides can demonstrate differing binding efficiencies with respect to liquid chromatography (LC) separation prior to MS-analysis, or ionisation efficiency and therefore the difference in signal observed may not be associated with the abundance of the modified peptide, but the change in properties of the peptide. This is further complicated by the presence of multiple modifications [43–45]. Despite the many advantages of MS-based analysis of PTMs, a number of limitations related to this approach exist. These are largely associated with the sensitivity of current instrumentation, i.e. how well the instrument can detect low intensity signals, and the speed of the instrument, i.e. how quickly the instrument is able to sequence the selected analytes and therefore the depth of coverage of the sample, as well as bioinformatics limitations. However, with the advent of ever increasingly rapid and sensitive instrumentation, and improved software capabilities, the deeper into the proteome and subsequently the PTMome, it is possible to investigate.

Detection of PTMs by top-down and bottom-up MS approaches

Figure 1
Detection of PTMs by top-down and bottom-up MS approaches

Top-down MS approaches involve analysis of the intact protein, with fragmentation providing sequence, PTM information and PTM localisation, but the complexity of the spectra can make it difficult to interpret. Therefore, in general, only isolated proteins or complexes are being efficiently analysed. High resolution MS and MS2 spectra using electron-transfer dissociation (ETD) fragmentation are the preferred analysis techniques. Middle-down involves analysis of large peptides, which allows for identification of coexisting PTMs. Separation of larger peptides is efficiently carried out only by specialised approaches e.g. weak cation exchange hydrophilic interaction LC (WCX-HILIC). Bottom-up MS includes analysis of proteolytically digested peptides typically ranging from 8 to 20 aa in length. Peptides carrying PTMs are very often separated from unmodified ones by application of various enrichment strategies (examples described in the text). The most commonly utilised peptide separation method is reverse phase chromatography (RPC). Mass spectrometry analysis can be performed using low resolution instruments, though some quantification methods require high resolution in MS or MS2 modes. Collision-induced dissociation (CID) or higher energy collisional dissociation (HCD) fragmentation methods are used to provide sequence and PTM localisation information.

Figure 1
Detection of PTMs by top-down and bottom-up MS approaches

Top-down MS approaches involve analysis of the intact protein, with fragmentation providing sequence, PTM information and PTM localisation, but the complexity of the spectra can make it difficult to interpret. Therefore, in general, only isolated proteins or complexes are being efficiently analysed. High resolution MS and MS2 spectra using electron-transfer dissociation (ETD) fragmentation are the preferred analysis techniques. Middle-down involves analysis of large peptides, which allows for identification of coexisting PTMs. Separation of larger peptides is efficiently carried out only by specialised approaches e.g. weak cation exchange hydrophilic interaction LC (WCX-HILIC). Bottom-up MS includes analysis of proteolytically digested peptides typically ranging from 8 to 20 aa in length. Peptides carrying PTMs are very often separated from unmodified ones by application of various enrichment strategies (examples described in the text). The most commonly utilised peptide separation method is reverse phase chromatography (RPC). Mass spectrometry analysis can be performed using low resolution instruments, though some quantification methods require high resolution in MS or MS2 modes. Collision-induced dissociation (CID) or higher energy collisional dissociation (HCD) fragmentation methods are used to provide sequence and PTM localisation information.

Box 2
Bottom-up MS analysis
  • The bottom-up analyses of PTMs follow a similar strategy. Proteins are digested into peptides, followed by an enrichment step in which modified peptides are separated from unmodified ones. Peptide mixtures are then separated by LC and analysed by MS. The obtained mass spectra of peptide masses (MS) and peptide fragment ions (MS2) are used to determine peptide sequences, PTM type and localisation, and protein identity. Thousands of modified peptides may be reliably identified in these studies.

  • Various peptide fragmentation methods are used to obtain peptide sequence information and PTM localisation site:

  • Collision-induced dissociation (CID), peptide ions are accelerated by an electrical potential and allowed to collide with neutral molecules (e.g. nitrogen). The collision results in peptide bond breakage and fragmentation into b- and y-ions. CID is effective for small, low-charged peptides, and is not suitable for fragmentation of intact proteins, and peptides with labile PTMs.

  • Higher energy collisional dissociation (HCD) is a CID technique specific to the orbitrap mass spectrometer. HCD also generates b- and y-type fragment ions. In contrast with CID, HCD does not suffer from the low mass cutoff. HCD is useful for de novo peptide sequencing and isobaric tag-based quantification. In PTM studies, low mass ions specific for HCD facilitate PTM identification.

  • Electron-transfer dissociation (ETD) induces fragmentation of multiply charged cations by transferring electrons to them. This causes peptide backbone cleavage into c- and z-ions while leaving labile PTMs intact. ETD is suitable for fragmentation of large peptides and intact proteins.

Box 3
Relative site occupancy of PTMs
  • Relative site occupancy is a measure of the percentage of a given site in a protein that is modified in comparison with the unmodified form (i.e. acetylation occupancy). It requires determination of the levels of the modified and unmodified forms of the peptide. Knowing site occupancy is important when investigating changes in PTM levels and the contribution of the various competing PTMs at a given residue following treatment or stimulation. A site with low occupancy (<1%) may increase five-fold to <5% occupancy, while the same increase to a site with higher occupancy (20%) would result in 100% occupancy. A site occupied at 50% cannot increase by more than two-fold.

Common problems associated with PTM investigation

While each PTM often comes with its own unique challenges, a number of hurdles are common amongst the PTMs. These can be broadly defined as a challenge of abundance, chemical properties, and bioinformatics (Table 1).

Table 1
Summary of common challenges associated with PTM analysis
ChallengeCurrent approach
Abundance  
Broad dynamic range of protein abundance
Substoichiometric occupancy of PTMs 
Modification-specific enrichment and LC-based separation of peptides 
Chemical properties  
Diversity Select appropriate sample preparation and analysis strategy 
Instability
High reactivity 
Take appropriate precautions during lysis and sample handling to both stabilise PTM and prevent PTM loss 
Select appropriate MS analysis techniques that minimise loss of labile modifications  
Low ionisation efficiency Modification-specific enrichment, chemically altering modifications and LC-based separation 
Impaired digestion Use alternate proteolytic enzymes 
Enrichment strategies  
Specificity Selection of appropriate, existing enrichment approach 
Availability Alternate MS approaches may be used, e.g. semi-targeted or targeted MS 
Loss of information Alternate MS approach which retains relevant information, e.g. top-down or middle-down MS 
Bioinformatics approaches  
Impaired digestion Increasing the number of expected missed cleavages when searching data 
Limited number of PTMs can be simultaneously searched Alternative analysis approaches e.g. de novo sequencing or open database searches 
ChallengeCurrent approach
Abundance  
Broad dynamic range of protein abundance
Substoichiometric occupancy of PTMs 
Modification-specific enrichment and LC-based separation of peptides 
Chemical properties  
Diversity Select appropriate sample preparation and analysis strategy 
Instability
High reactivity 
Take appropriate precautions during lysis and sample handling to both stabilise PTM and prevent PTM loss 
Select appropriate MS analysis techniques that minimise loss of labile modifications  
Low ionisation efficiency Modification-specific enrichment, chemically altering modifications and LC-based separation 
Impaired digestion Use alternate proteolytic enzymes 
Enrichment strategies  
Specificity Selection of appropriate, existing enrichment approach 
Availability Alternate MS approaches may be used, e.g. semi-targeted or targeted MS 
Loss of information Alternate MS approach which retains relevant information, e.g. top-down or middle-down MS 
Bioinformatics approaches  
Impaired digestion Increasing the number of expected missed cleavages when searching data 
Limited number of PTMs can be simultaneously searched Alternative analysis approaches e.g. de novo sequencing or open database searches 

Abundance

In a complex mixture resulting from, for example whole cell lysate, proteins exist within a broad dynamic range of abundance which can be a challenge due to the speed and sensitivity limitations of current instrumentation. In addition, PTMs are generally present at substoichiometric levels. This is illustrated through the observation that while it has been shown that approximately 75% of the proteome can be phosphorylated [46], the occupancy of the identified sites at a given time has been estimated to largely range between 0 and 20% [47]. As a result, PTMs can be difficult to analyse by MS as the signal resulting from the modified species often is dwarfed by the signal from the unmodified species, preventing its identification. This is because, as described in the previous example of a typical ‘top 20’ MS2 experiment, only the most intense ions are selected for fragmentation. These difficulties can be overcome by enrichment of the modified species, reducing the interference from the unmodified signal, and by separation through the commonly used approach of LC, simplifying the mixture of peptides and increasing the probability of the less abundant signals being observed. However, with more sample handling comes the increased risk of sample loss, therefore it is often necessary to use relatively large starting amounts of material to overcome this [48].

Chemical properties

A more complicated challenge is the chemical properties of the PTMs. While enrichment can be used to overcome the problem of low abundance, the method by which enrichment occurs can be difficult to develop. Numerous enrichment strategies have emerged over the years, in general these are commonly based on chemical affinity (such as the affinity between phosphorylation moieties and metal ions [49]), antibody-based affinity [50], and chemical alteration of the modification with the use of chemical tags or probes [51]. In order to be effective, enrichment methods must be specific for the target modification to minimise non-specific binding or reactions [52,53]. Additionally, information can be lost through the use of enrichment. For example, as modified and unmodified peptides are often analysed separately, it is no longer possible to determine the site-occupancy of a particular modification. This means that although a proportional change in modification between conditions can be observed, it is no longer possible to determine the proportion of a protein population modified at a particular site within a single condition [54]. The enrichment method utilised must be carefully considered as many come with limitations. For example, antibody-based methods can often be limited due to the specificity of the antibody. This approach is common in the enrichment of lysine acetylation, however, many of the antibodies used are found to be biased towards specific motifs, rather than the modification in general. As a result, a cocktail of antibodies may be used to achieve a more global enrichment [55]. Alternatively, enrichments based on chemical modification or chemical tagging rely on efficient and specific chemical reactions, such as the selective reduction in oxidative modifications of cysteine. But inefficient chemical reactions and unintended side reactions can hamper these forms of enrichment [38]. It is therefore important to consider not only the benefits of the chosen enrichment technique, but also the limitations. Further to the difficulties associated with the enrichment of PTMs is the issue of stability. Many PTMs, such as phosphorylation or oxidative modifications are labile, reversible, and capable of being further modified [56,57]. As a result, it is important to take steps to minimise the loss of modifications during sample preparation. This can be achieved through, for example the inclusion of inhibitors if modifications are enzymatically driven, or stabilising the modification through the composition of the lysis buffers preventing further reactions, examples of this are described in the relevant PTM sections [58,59]. To be compatible with MS, peptides need to be ionisable. Unfortunately, many PTMs will often, but not always, reduce the ionisation efficiency of peptides, therefore the modified peptide will often give a much weaker signal than the unmodified peptide [60]. As above, by enriching the PTMs of interest and reducing complexity through LC, the probability of the modified peptide being selected for analysis by the instrument is increased [61]. For bottom-up MS approaches, efficient digestion is vital to ensure accurate database matching and maximum coverage of the proteome, but many PTMs impair efficient enzymatic digestion [62]. One example of this is lysine acetylation, the most commonly used enzyme in bottom-up studies is trypsin, which cleaves on the C-terminal side of lysine and arginine residues except when the following residue is proline. However, with the addition of acetylation, the enzyme may no longer be able to cleave following a modified lysine residue [63]. This can be overcome through the use of alternative proteolytic enzymes, such as GluC, which typically cleaves on the C-terminal side of glutamic acid, or AspN, which cleaves on the N-terminal side of aspartic acid [64,65]. The selection of alternate proteases comes with its own caveats, such as variable digestion efficiency and reduced MS ionisation and fragmentation efficiency [65–68]. Alternatively, this cleavage inefficiency can be overcome through changing the way data are searched against known databases.

Bioinformatics

Typically, data are matched against a known database (see Box 4) with specific parameters, in the case of digestion, most commonly through restricting the search to allow for either one or two missed cleavage sites. However, when studying lysine acetylation, this may be increased to three or four missed cleavage sites. While this may increase the possibility of false identifications, it also increases the likelihood of identifying the modified peptides. This increase in false-positives due to increased ‘search space’, or number of possible variables, is one of the limitations of the traditional method of sequence matching commonly used, as this limits the number of modifications that can also be searched [69]. In general, the challenges and limitations of bioinformatics processing of MS data can be grouped into a number of categories, the first of which is the identification of the modification. The most common approach for bottom-up MS PTM experiments is to search the experimental spectra against a database of theoretical spectra generated in silico. This requires prior knowledge regarding the modifications expected within the dataset and the condition of the data, for example the expected number of missed cleavages resulting from enzymatic digestion. This approach, as mentioned above, is limited in that as the number of variables searched increases, such as increased numbers of modifications, the effective search space and therefore probability of false identifications is increased [70,71]. More recently, search tools that work by matching experimental spectra with a database of known spectra, as opposed to theoretical spectra predicted from the sequence, such as pMatch [12], QuickMod [72], and ANN-SoLo [73], or that predict a sequence’s potential modifications based on de novo sequencing, such as MSFragger [74], have been developed. These tools allow for an agnostic approach to searching PTM datasets, enabling the identification of unexpected modifications within the data, overcoming the limitation of traditional search tools with regard to the number of modifications that can be searched, and thus improving the depth of coverage of the PTMome. However, due to the computing time often required by these approaches and the need to individually validate the peptide matches due to the increased possibility of false identifications, the searching of large-scale datasets can prove to be impractical, and as such these approaches are generally limited to small-scale datasets. When selecting a search engine, it is vital to understand the concepts behind how the software matches data, detailed explanations of the common approaches can be found in [75–77]. In addition to the identification of the modification, the localisation of the modification can also be challenging. This is largely due to the high level of sequence coverage surrounding potential modification sites required (see Box 5 for more detail). The most common causes of poor sequence coverage, and therefore impaired site localisation, are low-signal intensity leading to poor signal-to-noise ratios, and poor fragmentation. This is usually the result of low abundance or poor ionisation of either the modified peptide or the specific fragment ion containing the modification, or as a result of modifications that can impair fragmentation [78,79]. Further to the bioinformatics challenges associated with identification is the prevalence of non-unique peptides. Often, particularly with closely related proteins or isoforms, there exists a high level of sequence similarity between proteins, with many peptides possessing identical sequences. This can limit the ability of software to accurately identify which protein is modified as it is often not possible to distinguish which protein the peptides originated from [80]. Another challenge of bioinformatics analysis is that of quantitation, in order to quantify a PTM, it is not enough to calculate the peak area of an analyte (label-free) or a reporter ion (labelled) for the modified peptide of interest. It is also essential to quantify the abundance of the total protein in order to normalise the change observed in the modification against any change in abundance of the overall protein. Traditional PTM-focussed bottom-up MS approaches, however, will often separate the analysis of modified and unmodified peptides. As a result, it is only possible to calculate the relative change in abundance of a modification across samples and prevents the calculation of site-occupancy, as discussed above.

Box 4
Methods of data searching

  • MS data analysis, Peptide fragmentation spectra (MS2) are used for assigning peptide sequence and identification of the originating protein. This can be achieved using different approaches.

  • Database search algorithms are based on interrogation of databases containing all amino acid sequences assumed to be present in the analysed sample are used.

  • De novo sequencing is the process in which differences between masses of fragment ions from a mass spectrum are used to determine amino acids sequence. This analysis is carried out without prior knowledge of protein sequence data.

  • Spectral matching compares new spectra with libraries of identified spectra. In contrast with database searches this approach can make use of the intensities of fragment peaks in library spectra to assess the quality of a match.

Box 5
PTM localisation by MS
  • Localisation of modification site based on the MS2 spectrum of a peptide is possible only if the presence of one or more ions can be unambiguously assigned to fragmentation between two amino acids in the peptide that can bear the modification. If such an ion is not observed, the modification can not be confidently assigned to either site.

Phosphorylation

One of the most abundant and commonly studied PTMs, phosphorylation is associated with many cellular processes [81]. The addition and removal of the phosphate moiety is enzymatically catalysed by kinases and phosphatases respectively and occurs most often, but not exclusively, on serine, threonine, and tyrosine residues [81,82]. It has a well-established central role in signal transduction cascades [83], but despite being one of the most heavily studied PTMs, much of the phosphoproteome is yet to be fully characterised, as emphasised in a recent review [84]. Numerous methods have been developed to enrich phosphopeptides, the most common of these are based on the affinity between the phosphate moiety and metal oxides or metal ions, for example titanium dioxide (TiOx), or Fe3+-based immobilised metal ion affinity chromatography (IMAC) (as reviewed in [85]). Despite these widely used, well-established approaches, more specific enrichment strategies continue to be developed. One such strategy is the use of molecularly imprinted polymers (MIPs) to create synthetic recognition sites, similar in effect to antibodies [86]. Due to the labile nature of the phosphorylation modification, the identification, and particularly localisation, of the modification was traditionally difficult using MS [87]. This was because MS2 fragmentation was performed using collision-induced dissociation (CID), which is the result of multiple low-energy collisions with gas molecules [88]. However, this method of fragmentation is slow, allowing the modification to ‘jump’ to previously unmodified amino acids, and low energy, allowing the modification to be lost from the peptide without fragmentation of the backbone. The development of new fragmentation methods, such as electron-transfer dissociation (ETD), and particularly the advent of beam-type CID, such as higher energy collisional dissociation (HCD) has largely improved the fragmentation of phosphorylated peptides [89,90]. HCD in particular has become a common-place method for analysing phosphorylation on a large scale as the acceleration of ions into the neutral gas allows for faster, higher energy collisions preventing the modification from jumping, and allowing for fragmentation of the peptide backbone [90].

Lysine acetylation

Similar to phosphorylation, lysine acetylation is thought to be involved in almost all biological processes within the cell, largely through regulation of transcription [91,92]. But it is particularly involved in metabolic control through the suppression of enzymatic activity [93], and mitochondrial function (as reviewed in [94]). Typically, the addition and removal of the acetyl group is enzymatically driven through the action of histone/lysine acetyltransferases and histone/lysine deacetylases respectively [95], however it has also been shown to occur nonenzymatically within the mitochondrial matrix [96]. The most prevalent complications associated with the study of lysine acetylation are abundance and impaired digestion efficiency. Common approaches for the enrichment of lysine acetylation include antibody-based enrichments, with most approaches utilising a combination of antibodies, rather than a single antibody in order to prevent a biased enrichment based on the recognition motif [55]. As described above, lysine acetylation impairs digestion efficiency which is often overcome through sample preparation, using alternative proteolytic enzymes, or bioinformatically by increasing the number of predicted missed cleavages [63]. Similarly, two of the most common methods of labelling for quantification, iTRAQ and TMT, work by reacting with free amine groups and therefore label the N-terminus and lysine residues, as a result the labelling efficiency may be reduced [35,36]. To overcome this, alternative labelling strategies such as SILAC may be used. However, depending on the sample type, this may not be possible as SILAC requires the incorporation of differing isotopes of amino acids, which while possible in cell culture and some animal models, is not possible in, for example clinical samples [97]. Alternatively, as with the challenge of digestion efficiency, this may be accounted for by matching the data against a known database with the label considered to be a dynamic modification rather than a fixed modification. Again, this increases the possibility of false positives by expanding the search space but overcomes the problem of the label not being incorporated by the modified lysines. In addition, for highly modified proteins such as histones, the presence of a high number of lysine residues and therefore tryptic cleavage sites can lead to the generation of peptides that are too short to analyse with a typical bottom-up approach [98,99]. To overcome this, proteolytic enzymes with fewer cleavage sites, generating longer peptides and taking a middle-down approach, as discussed below, can be used.

Oxidative modifications

Although challenges still exist in the analysis of phosphorylation and lysine acetylation, these modifications have been investigated, and technologies and methodologies for their analysis improved for a number of decades [91,100]. However, there are many other, less well-established, fields of PTM-focussed research which are coming to the forefront of disease research and pose a significant challenge in their analysis due in part to less well-established methods, one such group of PTMs are the oxidative modifications. Oxidative modifications are a chemically diverse group of modifications that were once traditionally thought to be associated with damage, however they are now understood to often play an important signalling role within the normal function of the cell [101]. In particular, cysteines can act as redox molecular switches [102], an example of this is the control of the transcription factor Nuclear factor erythroid 2-related factor 2 (Nrf2). The translocation of Nrf2 is regulated by the oxidative state of its interaction partners, such as Kelch-like ECH-associating protein 1 (Keap1) which, when oxidised, dissociates from Nrf2 allowing its translocation to the nucleus [103]. These modifications can occur both enzymatically and through a chemical reaction with oxidising species [104–106]. The chemistries that dictate the formation of these modifications are as diverse and varied as the modifications themselves, with extensive work being undertaken to characterise these processes [107]. Oxidative modifications can occur on a number of amino acids, the most reactive of which are cysteine and methionine residues, but also extends to histidine, proline, lysine, arginine, tyrosine, and tryptophan [22]. Oxidative modifications are a highly challenging group of modifications to investigate. This is in part due to many of them being reversible, and able to be further oxidised into other forms of modification. In addition, they cover a broad range of chemistries, and are difficult to completely inhibit during sample preparation in the same way as other modifications, although performing sample preparation under anaerobic conditions or with the inclusion of antioxidants can mitigate the formation of artefactual modifications. This is because many are not enzymatically driven, this increases the possibility of the introduction of oxidation artefacts [108]. Despite its challenging nature, the investigation of oxidative modifications has become increasingly prevalent, particularly with regard to their role in diseases associated with increased oxidative stress, such as diabetes and cardiovascular disease [109,110].

Cysteine thiol oxidation

As one of the more reactive amino acids, oxidative modifications of cysteines have become an area of significant focus. Due to the wide range of oxidative states the sulphur atom of the thiol group can exist within (−II to +IV), cysteines can be modified in a variety of ways, the vast majority of which are considered to be biologically reversible, such as disulphide bonds, S-nitrosylation and S-glutathionylation amongst others. In addition, cysteines can be modified in a way which is generally considered to be biologically irreversible through sulphinylation and sulphonylation. These modifications in particular are thought to be damaging, while the reversible modifications are more generally thought to be associated with signalling [38,111]. Due to the low abundance, labile nature, and chemical diversity of these modifications, combined with the possibility of artefactual oxidation and the ability of the modifications to interchange, the analysis of these modifications presents a significant technical challenge. One of the major challenges associated with the analysis of cysteine oxidation is its instability and the possibility of artefactual modifications during sample preparation. As a result, it is vital to stabilise the free thiols as rapidly as possible and minimise the opportunity for continued oxidative reactions. This can be achieved by combining quenching reagents, such as trichloroacetic acid, and alkylating agents, such as N-ethylmaleimide or iodoacetamide, to block biologically free thiols during initial lysis and denaturation [112–114]. Furthermore, metal ions can often catalyse oxidative reactions, therefore the addition of chelating agents such as diethylenetriaminepentaacetic acid, neocuprione, or ethylenediaminotetraacetic acid can also be added during initial lysis to further protect the integrity of the sample and prevent artefactual oxidation [37,59,115].

Beyond the handling of the samples prior to MS analysis, the detection of these modifications can prove problematic due to their highly labile nature. Although direct detection of a number of the modification species is possible, and has indeed been reported previously [116,117], it is more common for the unique chemistry of the modifications to be utilised for chemical derivatisation methods, or direct chemical labelling approaches. These methods also enable enrichment of the modifications, thus overcoming the challenge of low abundance common to PTMs.

Methods for the enrichment and detection of oxidatively modified cysteines can be grouped in a number of ways. They can be either general, large-scale approaches aimed at detecting all modified cysteine residues irrespective of the type of modification, or they can be specific, targeted approaches aimed at detecting a particular type of modification. While generally, large-scale approaches, such as those developed by Paulech et al. [118] and Huang et al. [119] are useful in gaining a global picture of the oxidative state of the system under investigation and to identify proteins which are potentially regulated by oxidative modifications, the removal of the identity of the modification makes it difficult to fully understand the implications of the changes observed without targeted validation. Alternatively, targeted methods such as that developed by Wojdyla et al. [37], designed to enrich S-nitrosylation and sulphenylation, provide information about the type of modification impacting the protein. However, these methods also ‘limit the field of view’ as only a restricted snapshot of the specific modifications can be observed, as opposed to a global view of the impact of oxidation on the system. Currently, no method has been developed which enables both simultaneously. Additionally, methods for the investigation of oxidative modifications of cysteine can be grouped as either direct or indirect chemical strategies (Figure 2). Direct methods involve a reaction between the specific modification and a chemical group which is often either tagged or bound to a resin to allow enrichment, for example the enrichment of S-nitrosylation through its reaction with organomercury bound to a resin for enrichment [120]. While indirect methods generally involve the selective (e.g. reduction of S-nitrosylation by ascorbate) or global (e.g. by dithiothreitol) reduction of modified cysteine residues into free thiols, and enrichment via either chemical tagging or affinity-based enrichment. However, it is important to note that many of these methods are multi-stage enrichments and involve a significant amount of sample handling. It is therefore important to ensure complete blocking of the biologically free thiols and to reduce the oxidative environment of the sample as much as possible to prevent artefactual oxidation and conversion or loss of oxidative modifications. It is also important to be aware of the limitations of the enrichment method used, for instance while a number of reducing agents are considered to be specific to a particular modification, many have been found to be less specific than originally thought, for example ascorbate has also been shown to reduce disulphide bonds [53].

Summary of the common components of cysteine oxidative modification enrichment strategies

Figure 2
Summary of the common components of cysteine oxidative modification enrichment strategies

Direct methods (left side) take advantage of the reactivity of the individual modifications often linking reactants with a tag such as biotin to enable the targeted analysis of individual modifications, for example S-nitrosylation reacts with organomercury. Indirect methods (right side) involve the removal of individual modifications in either a global or specific manner using reducing agents, enrichment is then reliant on thiol-based affinity or reactivity. Both direct and indirect methods generally require prior alkylation of biologically free thiols to minimise artefactual oxidation.

Figure 2
Summary of the common components of cysteine oxidative modification enrichment strategies

Direct methods (left side) take advantage of the reactivity of the individual modifications often linking reactants with a tag such as biotin to enable the targeted analysis of individual modifications, for example S-nitrosylation reacts with organomercury. Indirect methods (right side) involve the removal of individual modifications in either a global or specific manner using reducing agents, enrichment is then reliant on thiol-based affinity or reactivity. Both direct and indirect methods generally require prior alkylation of biologically free thiols to minimise artefactual oxidation.

Nitration and chlorination

Tyrosine is another amino acid which is known to undergo a variety of oxidative modifications, many of which can undergo further oxidation into secondary modifications. One example of this is the oxidation of tyrosine leading to the formation of 3,4-dihydroxyphenylalanine, which can be further oxidised into dopaquinone [121]. Although this amino acid is less reactive when compared with cysteine, it is of particular interest due to the potential interference its oxidation may have with tyrosine phosphorylation and subsequent signalling events [122]. Two forms of oxidative modification that are known to occur on tyrosine are nitration, the addition of an NO2 group to the phenolic ring structure of tyrosine as a result of the action of reactive nitrogen species such as peroxynitrite, and chlorination, the addition of a Cl to the phenolic ring structure of tyrosine as a result of the action of hypochlorous acid [123,124]. Both these modifications are largely stable and markers of oxidative stress [125,126]. Of the two modifications, nitration is the most extensively studied, and although it is not enzymatically driven, it has been shown to be a highly selective modification with only a small group of proteins known to be modified in this way [127]. Although nitration is a relatively new field of study when compared with a well-established field such as phosphorylation, a number of methods have been developed to investigate this modification. A common approach utilises separation by two-dimensional gel electrophoresis, followed by antibody-based detection and protein identification by MS [128–130]. While this approach is useful, it is limited by a number of factors. These include sensitivity; only highly abundant proteins will likely be observed when complex samples are being analysed, and compatibility; this approach is biased towards highly soluble proteins while membrane-bound proteins are likely to not be compatible with gel-based separation [131]. Attempts are being made to develop antibody-based enrichment methods similar to those routinely used for the analysis of lysine acetylation. However, these are currently limited due to the specificity of the antibodies available as many are targeted towards specific consensus sequences rather than the modification in general, or show a bias towards a particular localisation of the modification, for example the N-terminal [132]. Alternatively, chemical derivatisation methods have been developed, many of which revolve around blocking the existing primary amines followed by conversion of the nitrated tyrosine residue into an aminotyrosine and enrichment using, for example, biotin tagging. These methods often involve multi-step preparations that rely on the efficiency of chemical reactions, and therefore can be problematic due to sample loss and inefficient chemical reactions [133,134].

Unlike the modifications mentioned previously, the less extensively studied tyrosine chlorination does not currently have any enrichment methods published. This modification has largely been studied in either single protein or highly simplified systems with few large-scale studies undertaken to date [29,135]. A number of biomarker studies do exist, however, these merely identify the presence and/or level of chlorinated tyrosine within biological fluids such as plasma or urine, but they do not provide information regarding the protein of origin, or site-localisation within the protein [124]. In the absence of an enrichment method, a semi-targeted MS approach can be utilised to study chlorination in more complex protein samples. In the case of chlorination this involves scanning for signals that, upon fragmentation, generate a diagnostic ion with an m/z of 170.1, however the presence of isobaric signals can lead to false-positive identifications. As a result, an additional fragmentation of a chosen ion (MS/MS/MS (MS3)) can be used to increase the specificity of the identifications [22,29,136]. Despite this semi-targeted approach, due to the low abundance of chlorinated tyrosines, this method can still be limited when used to analyse more complex samples.

PTM cross-talk

PTMs do not exist in isolation. Proteins can be modified at multiple sites, and by multiple types of PTMs, as a result it is vital to understand how modifications interact, and how this interaction impacts protein function, termed ‘cross-talk’ [2] (see Box 6). Cross-talk can be either positive or negative in nature and can occur in a number of ways (Figure 3). PTMs can act as a signal for other PTMs to be either added or removed or form a recognition site for proteins in positive cross-talk. While negative cross-talk may take the form of direct competition for the same site, or may act indirectly by obscuring a secondary modification site (as reviewed in [137]). Cross-talk can also be linear in behaviour, that is modifications can have an additive effect leading to a stronger response, or non-linear in behaviour, that is there is a tipping point between a functional impact and no functional impact with the addition or removal of multiple PTMs (as reviewed in [2]) (Figure 4). PTMs often interact through proximity, either in sequence or 3D structure, or by inducing conformational change. However, it is often difficult to determine whether two PTMs are present on the same protein at the same time or not as most studies utilise bottom-up approaches, typically using trypsin as the digestion enzyme of choice. While trypsin is an ideal enzyme for traditional studies, part of what makes it so useful; the generation of relatively short peptides, is also what makes cross-talk studies difficult. This is because the shorter the peptides, the lower the probability of identifying several PTMs on the same peptide. Therefore, utilising alternative enzymes, such as GluC [138,139], which generates longer peptides, and as a result taking what is termed a ‘middle-down’ approach, increases the probability of PTMs being identified together [139]. Cross-talk studies are also limited by the methods available, as only a few PTMs have well-established enrichment procedures associated with them, limiting which PTM interactions can be studied on a large-scale. Currently most PTM cross-talk studies focus on either a small subset of PTMs, usually two to three, or a more complete characterisation of PTMs on a single protein. For example, the PTM interplay on histones is one of the more extensively studied single-protein systems [140–142]. While advances are being made in the analysis of individual PTMs, and some progress has begun on the investigation of cross-talk, the large-scale analysis of large numbers of PTMs and their interactions remains a significant challenge in a complex systems environment.

Interaction mechanisms in PTM cross-talk

Figure 3
Interaction mechanisms in PTM cross-talk

Positive cross-talk can involve one PTM triggering the (A) addition or (B) removal of another PTM through, for example conformational change of the protein, or alteration of the microenvironment. Negative cross-talk typically involves (C) direct competition for the same modification site or (D) obscuring a modification site in close proximity through altered protein confirmation, microenvironment, or steric hindrance.

Figure 3
Interaction mechanisms in PTM cross-talk

Positive cross-talk can involve one PTM triggering the (A) addition or (B) removal of another PTM through, for example conformational change of the protein, or alteration of the microenvironment. Negative cross-talk typically involves (C) direct competition for the same modification site or (D) obscuring a modification site in close proximity through altered protein confirmation, microenvironment, or steric hindrance.

Effect mechanisms of PTM cross-talk

Figure 4
Effect mechanisms of PTM cross-talk

(A) Linear cross-talk refers to PTMs that are cumulative, leading to a gradient-type effect, in which the addition of consecutive PTMs gradually increases the effect. (B) Non-linear cross-talk refers to PTMs that have a binary-type effect, with a specific number of PTMs or PTM combinations required before an effect is observed.

Figure 4
Effect mechanisms of PTM cross-talk

(A) Linear cross-talk refers to PTMs that are cumulative, leading to a gradient-type effect, in which the addition of consecutive PTMs gradually increases the effect. (B) Non-linear cross-talk refers to PTMs that have a binary-type effect, with a specific number of PTMs or PTM combinations required before an effect is observed.

Box 6
PTM cross-talk mechanisms
  • PTM cross-talk, The combination of different PTMs on a protein can create a ‘PTM code’, which can be recognised by specific effectors to initiate/inhibit downstream events, only inducing/retaining a signal once the complementary incoming signals are present at the same time and place.

  • In positive cross-talk, one PTM can serve as a signal for the addition or removal of a second PTM, or for recognition by a binding protein that carries out a second modification.

  • Negative cross-talk occurs through a direct competition for modification of a single residue in a protein, or indirectly by masking the recognition site for a second PTM.

Conclusion

Several technical challenges persist in the field of PTM analysis, ranging from biochemical challenges associated with improving and developing enrichments, to the bioinformatics challenges associated with the interpretation of MS data. However, identifying which proteins are modified, how and where, is only half the battle. While databases such as PhosphoSitePlus [143], RedoxDB [144], and CrossTalkDB [145] are being filled with PTM identifications, a relatively small percentage of these sites have a known biological function. The vast gap in knowledge and biological understanding of PTMs, and their roles in biology combined with the technical challenges described in this review is one of the most complex scientific challenges since the advent of the Human Genome Project.

Summary

  • PTMs are chemical modifications of proteins that are involved in both normal function and disease mechanisms within the cell.

  • Despite a large focus on the characterisation of PTMs, their analysis remains a challenge.

  • MS is a fast and sensitive method for the large-scale and site-specific mapping of PTMs.

  • Future studies need to focus on developing robust large-scale methods for analysis of all biologically relevant PTMs and their interplay.

Competing Interests

The authors declare that there are no competing interests associated with the manuscript.

Funding

This work was supported by the Independent Research Fund Denmark [grant number DFF – 4181-00172 (to L.E.S.)].

Author Contribution

This review was designed by L.E.S. and A.R.W., written by L.E.S. and edited by A.R.W.

Abbreviations

     
  • CID

    collision-induced dissociation

  •  
  • ETD

    electron-transfer dissociation

  •  
  • HCD

    higher energy collisional dissociation

  •  
  • iTRAQ

    isobaric tagging for relative and absolute quantitation

  •  
  • LC

    liquid chromatography

  •  
  • m/z

    mass-to-charge ratio

  •  
  • MS

    mass spectrometry

  •  
  • MS2

    MS/MS

  •  
  • Nrf2

    nuclear factor erythroid 2-related factor 2

  •  
  • PTM

    post-translational modification

  •  
  • SILAC

    stable isotope labelling by amino acids in cell culture

  •  
  • TMT

    tandem mass tagging

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