Post-translational control of proteins through covalent attachment of ubiquitin plays important roles in all eukaryotic cell functions. The ubiquitin system in humans consists of 2 E1, 35 E2 and >600 E3 ubiquitin ligases as well as hundreds of deubiquitylases, which reverse ubiquitin attachment. Moreover, there are hundreds of proteins with ubiquitin-binding domains that bind one of the eight possible polyubiquitin chains. Dysfunction of the ubiquitin system is associated with many diseases such as cancer, autoimmunity and neurodegeneration, demonstrating the importance of ubiquitylation. Therefore, enzymes of the ubiquitin system are considered highly attractive drug targets. In recent years, mass spectrometry (MS)-based techniques have become increasingly important in the deciphering of the ubiquitin system. This short review addresses the state-of-the-art MS techniques for the identification of ubiquitylated proteins and their ubiquitylation sites. We also discuss the identification and quantitation of ubiquitin chain topologies and highlight how the activity of enzymes in the ubiquitin pathway can be measured. Finally, we present current MS tools that can be used for drug discovery in the ubiquitin space.

Introduction

Ubiquitylation is a post-translational modification involving the covalent binding of the small, highly conserved protein ubiquitin, consisting of 76 amino acids, to a target substrate to regulate protein functions. The ligation of the ubiquitin C-terminus to a target protein is facilitated by a cascade of ubiquitin-activating and ligating enzymes (E1, E2 and E3) that work in tandem to conjugate ubiquitin predominantly to lysine residues in substrate proteins. This post-translational modification can be reversed by deubiquitylating enzymes (DUBs) for which six families have been identified [1,2]. Ubiquitin possesses seven lysine residues and an N-terminus (M1, K6, K11, K29, K33, K48 and K63), any one of which can be ubiquitylated itself leading to the assembly of polyubiquitin chains [3]. These chains have substantially different structures (Figure 1) and lead to different biological outcomes; for example, while K48 chains lead to proteasomal degradation, K63 chains are linked to cell signalling trafficking and lysosomal degradation [4]. As ubiquitin chains can be formed of a single type, mixed or even branched [5,6], there is potential to have a huge range of different ubiquitin chain topologies thus resulting in highly complex and varied signalling [3,4]. Ubiquitin signalling is integral to almost all cellular processes in eukaryotes, and thus, disorders or mutations in ubiquitin pathways often result in diseases including cancer, a wide range of autoimmune diseases and neurodegenerative diseases such as Parkinson's, Alzheimer's and Huntington's disease [710].

The structures of the ubiquitin linkages.

Figure 1.
The structures of the ubiquitin linkages.

Surface representations of crystallized diubiquitin isomers show substantially different structures leading to different biological outcomes [19,7580]. M1 chains are important for innate immune signalling, K48 chains lead to proteasomal degradation and K63 chains have various roles in signalling, endolysosomal trafficking and degradation. The roles of the atypical chains K6, K11, K27, K29 and K33 are less well understood (for review, see [4]).

Figure 1.
The structures of the ubiquitin linkages.

Surface representations of crystallized diubiquitin isomers show substantially different structures leading to different biological outcomes [19,7580]. M1 chains are important for innate immune signalling, K48 chains lead to proteasomal degradation and K63 chains have various roles in signalling, endolysosomal trafficking and degradation. The roles of the atypical chains K6, K11, K27, K29 and K33 are less well understood (for review, see [4]).

Mass spectrometry (MS)-based proteomics has become a powerful method for the analysis of ubiquitylation in the last 15 years. Great technological progress in MS and liquid chromatography instrumentation as well as software and sample preparation techniques have substantially improved speed and sensitivity, allowing now the characterisation of low-abundant post-translational modifications. In a standard peptide-based proteomic ‘bottom-up’ approach (Figure 2), proteins are extracted from whole organisms, tissues, cells or organelles and digested into peptides using highly specific proteases such as trypsin. These peptides are then separated on a (nano)-High-Performance Liquid Chromatography (HPLC) according to their hydrophobicity and eluted into a spray needle from which they are ionised by electrospray ionisation (ESI). Peptide ions enter the mass spectrometer and are then analysed in two ways: first, in a so-called survey scan, the mass (actually, mass-to-charge ratio; m/z) and intensities of the peptide ions are determined. This provides quantitative information about the abundance of the peptide. Secondly, these peptide ions are then isolated and fragmented by tandem mass spectrometry (MS/MS), providing amino acid sequence information about the peptide. In a next step, protein databases of the studied organism can be searched by specialist software utilising both accurate mass of the peptide and the sequence information, leading to the identification of the peptide. If post-translational modifications are targeted, the modification of interest has to be included in the peptide search, thereby increasing the search space.

Bottom-up proteomics workflow.

Figure 2.
Bottom-up proteomics workflow.

The generic workflow of a standard peptide-based (‘bottom-up’) proteomic strategy consists of four steps. Proteins are extracted from whole organisms, tissues, cells or subcellular compartments, digested by a proteolytic enzyme and separated according to their hydrophobicity by HPLC. Eluted peptides are ionised by ESI. Mass-to-charge ratio (m/z) and intensity of peptides are detected and acquired in a mass spectrometer in a ‘survey scan’. Peptides are then selected for fragmentation by MS/MS resulting in information about the amino acid sequence. The resulting spectral data are searched against the protein coding information of the genome of the analysed organism to provide sequence identification.

Figure 2.
Bottom-up proteomics workflow.

The generic workflow of a standard peptide-based (‘bottom-up’) proteomic strategy consists of four steps. Proteins are extracted from whole organisms, tissues, cells or subcellular compartments, digested by a proteolytic enzyme and separated according to their hydrophobicity by HPLC. Eluted peptides are ionised by ESI. Mass-to-charge ratio (m/z) and intensity of peptides are detected and acquired in a mass spectrometer in a ‘survey scan’. Peptides are then selected for fragmentation by MS/MS resulting in information about the amino acid sequence. The resulting spectral data are searched against the protein coding information of the genome of the analysed organism to provide sequence identification.

In recent years, this type of proteomics has been instrumental in understanding ubiquitin biology. In this short review, we present a guide to proteomic techniques that are used to discover substrates of ubiquitylation, identify and quantify ubiquitin chain topologies, and measure the activity of enzymes in the ubiquitin pathway. Finally, we discuss the MS tools available for drug discovery in the ubiquitin field.

Enriching and identifying sites of ubiquitylation

Although the ubiquitin system itself is evolutionarily conserved, the sites of modification are much less so and are often difficult to predict by sequence analysis alone [11]. MS-based proteomics has been particularly important in identifying sites of ubiquitylation. Like almost all post-translational modifications, ubiquitylated peptides must be enriched prior to analysis by MS as the natural abundance of ubiquitylated proteins is usually very low in comparison with unmodified proteins [12]. The most commonly used strategy to enrich modified proteins is by overexpression of a His- or FLAG-tagged version of ubiquitin. However, it needs to be noted that overexpression of ubiquitin can lead to cellular stress and activation of many stress-induced pathways that could lead to induction of novel, unrelated ubiquitylation sites resulting in artefactual results [1316]. A more elegant enrichment of ubiquitylated proteins is through the use of tandem ubiquitin-binding entities (TUBEs) (Figure 3) [17]. TUBEs are artificial proteins with repetitions of two or four ubiquitin-binding domains. They are often attached to a tag, such as a Halo-tag, which allows for covalent binding to beads, and can then be used to pull-down ubiquitylated proteins from cell lysates. By comparing the pull-downs with control samples obtained with a non-binding TUBE, often generated by introduction of single amino acid changes, enriched ubiquitylated proteins can be identified and quantified by quantitative MS. The most commonly used TUBE is the ubiquitin-binding domain UBA of ubiquilin-1, which appears to bind all ubiquitin chains [17]. However, one can also use TUBEs to enrich for specific chains such as M1/linear (NEMO UBAN) [18], K29 (Trabid NZF) [19], K48 (MINDY-1 tUIM) [20] and K63 (Tab2 NZF) [21], thus providing a powerful tool to distinguish chain types and their substrates by proteomic techniques (Table 1). Recently, K63-specific TUBEs were used to show that K63-ubiquitylated proteins accumulate upon oxidative stress [22].

Workflow for enriching ubiquitylated proteins and di-Gly peptides.

Figure 3.
Workflow for enriching ubiquitylated proteins and di-Gly peptides.

To identify ubiquitylation sites, ubiquitylated substrates can be enriched either at the protein or peptide level. (Left) For enrichment at the protein level after lysis of cells or tissue, TUBEs can be employed. Some TUBEs are specific for certain chain types thereby allowing enrichment of polyubiquitin subsets. Enriched ubiquitylated proteins are then digested by a specific protease such as trypsin and the peptides are analysed by MS. Digesting each protein provides a large number of peptides, making its identification easier; at the same time, it makes the identification of specific ubiquitylation site-containing peptides more difficult. (Right) Ubiquitylation site di-Gly peptides can be highly enriched using specific commercial antibodies against the di-Gly motif. This enhances the chances of identifying an ubiquitylation site peptide (yellow stars), but proteins where the corresponding di-Gly peptide is not amenable to MS identification might be missed.

Figure 3.
Workflow for enriching ubiquitylated proteins and di-Gly peptides.

To identify ubiquitylation sites, ubiquitylated substrates can be enriched either at the protein or peptide level. (Left) For enrichment at the protein level after lysis of cells or tissue, TUBEs can be employed. Some TUBEs are specific for certain chain types thereby allowing enrichment of polyubiquitin subsets. Enriched ubiquitylated proteins are then digested by a specific protease such as trypsin and the peptides are analysed by MS. Digesting each protein provides a large number of peptides, making its identification easier; at the same time, it makes the identification of specific ubiquitylation site-containing peptides more difficult. (Right) Ubiquitylation site di-Gly peptides can be highly enriched using specific commercial antibodies against the di-Gly motif. This enhances the chances of identifying an ubiquitylation site peptide (yellow stars), but proteins where the corresponding di-Gly peptide is not amenable to MS identification might be missed.

Table 1
TUBEs and their control mutations
TUBEsSpecificityNon-binding control mutationReference
Ubiquilin UBA All ubiquitin chains M557K/L584K Hjerpe et al. [17
NEMO UBAN M1/linear D311N Rahighi et al. [18
Trabid NZF1 K29, K33 M26A Kristariyanto et al. [19
MINDY tMIU K48 A416D Kristariyanto et al. [20
TAB2 NZF K63 (and K6) T674A/F675A Kulathu et al. [21
TUBEsSpecificityNon-binding control mutationReference
Ubiquilin UBA All ubiquitin chains M557K/L584K Hjerpe et al. [17
NEMO UBAN M1/linear D311N Rahighi et al. [18
Trabid NZF1 K29, K33 M26A Kristariyanto et al. [19
MINDY tMIU K48 A416D Kristariyanto et al. [20
TAB2 NZF K63 (and K6) T674A/F675A Kulathu et al. [21

While this is a very sensitive and powerful approach, it needs to be noted that depending on protein size and peptide ionisation characteristics, the number of identified ubiquitylation site carrying peptides in these pull-downs is often low. Still, it remains that only the identification of these specific peptides provides satisfactory evidence that a substrate protein has been ubiquitylated. To identify ubiquitylation sites on a protein, the protein needs to be digested into peptides using specific proteases such as Trypsin (which cleaves C-terminally of Arg and Lys) or Lys-C (which cleaves C-terminally of Lys). The enzymatic digestion of ubiquitylated proteins leaves a specific amino acid remnant on the substrate peptide, a Gly-Gly (di-Gly) motif, attached to the ε-amine of the modified lysine [23,24]. Importantly, ubiquitylation, like other post-translational modifications, needs to be preserved during cell lysis [25]. Unspecific and very active DUBs readily cleave off ubiquitin from their substrates or ubiquitin chains, thus making the use of DUB inhibitors during cell lysis mandatory in a proteomic workflow. There are a range of DUB inhibitors of varying specificity available [26], but to preserve the majority of native ubiquitylation alkylating agents such as N-ethylmaleimide or iodoacetamide are commonly used. These react with active site cysteines present in E1/E2/E3 ligases and most DUBs (except for the JAMM metalloprotease family), inhibiting them permanently. However, as iodoacetamide can side-react with lysines which leaves a modification of the same mass as a di-Gly remnant, it is no longer recommended for this purpose [27].

To enrich for ubiquitylated peptides, a powerful method using a commercially available monoclonal antibody against di-Gly remnants was developed by Kim et al. [28]. Coupled with traditional liquid chromatography (LC)–MS methods, this method drastically increased the number of sites and ubiquitylated proteins identified. In total, ∼19 000 di-Gly modified residues within ∼5000 proteins were identified in HCT116 cells with >94% of these sites corresponding to ubiquitin conjugation [28]. Since first reported, this antibody has been used extensively for ubiquitylation discovery proteomics in many studies [24,2933]. While it is not yet known if these antibodies enrich all di-Gly peptides at equal affinity, this immunoprecipitation approach is a versatile tool to study different types of ubiquitylation as it can be used in conjunction with robust quantitative MS approaches.

Moreover, it is important to note that some ubiquitin-like modifiers (UBLs), namely ISG15 and NEDD8, release the same di-Gly motif after tryptic digestion as ubiquitin itself, which renders the distinction between these modifications virtually impossible. While work performed by Kim et al. [28] suggests that the pool of these UBL modified peptides is relatively small with most of the di-Gly modified peptides being ubiquitylated peptides, this distribution might change depending on the tissues, cells or conditions used.

Many of the large-scale ubiquitin proteomic studies published so far focused on studying K48-ubiquitylated proteins which under normal conditions are rapidly degraded by the proteasome [34]. Therefore, proteasome inhibitors such as MG132 [35] or bortezomib [36] are often used during lysis to enrich for this protein population. Unfortunately, proteasome inhibition leads to accumulation of polyubiquitylated proteins resulting in a severe shortage of monoubiquitin in the cell, which can induce other degradative pathways such as autophagy [37]. It is therefore important to consider the effects of the use of proteasome inhibition on the cellular ubiquitin system, limit the exposure time to these inhibitors where possible and ideally validate results without the use of inhibitors.

Choosing the best method of quantifying enriched ubiquitylated proteins and di-Gly peptides can be challenging, as there are several advantages and disadvantages to each method to be considered. Label-free quantitation is probably the most commonly used technique, as it is straightforward, sensitive and provides good quantitation. However, normalisation of peptides in di-Gly pull-downs can be challenging if changes between conditions are substantial. Here, metabolic labelling such as stable isotope labelling using amino acids in cell culture (SILAC) might be preferable as the lysates can be mixed before digestion, thus reducing problems with normalisation and equal loading [38,39]. Other common methods for absolute and relative quantitation in proteomics include labelling of peptides with reactive amine reagents, such as isotopically labelled tandem mass tags or isobaric tags for relative and absolute quantitation, which translate well to the study of ubiquitylation. However, these particular isotopic labelling strategies must be performed after a di-Gly enrichment as the labelled peptides will no longer be recognised by the di-Gly antibody [30,40]. Altogether, sensitive and selective tools are available to enrich for ubiquitylated proteins and di-Gly peptides for quantitative proteomics.

Quantifying chain types

Different ubiquitin chain types and linkages can play significantly different roles in cell signalling, but differentiating and quantifying these chain types is inherently challenging, often requiring the use of antibodies — although for several chain types no specific antibodies are available. Fortunately, MS analysis can bridge this gap as tryptic digestion of ubiquitin chains leads to unique peptides with a di-Gly motif at the site of ubiquitylation. All of these ubiquitin di-Gly peptides of the eight linkage types differ in sequence and can therefore be readily detected and quantified using MS [41]. It should be noted that as these peptides have distinct sequences, the number of ions generated for each of these peptide species, i.e. their ionisation efficiency, differs significantly and will be reflected by their intensities in the resulting spectra. For example, when comparing equimolar amounts of ubiquitin di-Gly peptides, the K63 and K29 peptides ionise substantially worse than K48 and K6 peptides, making their detection at low abundance more challenging. To determine absolute chain type abundance, an internal standard is essential. One of the most powerful tools currently available is the use of isotopically labelled absolute quantification (AQUA) peptides that enable the direct quantification of the abundance levels of ubiquitylated substrates [42]. In the case of ubiquitin, these synthetic peptides correspond to the di-Gly modified peptides carrying the seven lysines and the N-terminus, and are spiked into a cell lysate at a known concentration. Both isotopically labelled internal standards and unlabelled sample peptides can be detected by proteomics and their intensities compared, thus allowing absolute quantification of the different ubiquitin linkage types.

Targeted proteomic experiments, such as selective reaction monitoring (SRM) or parallel reaction monitoring (PRM) (Figure 4) [43,44], can be used to quantify known peptides such as the specific ubiquitin chain link peptides and have been widely applied. These technologies quantify peptides by measuring the intensities of fragments after MS/MS fragmentation [45], providing a much wider dynamic range while still quantifying attomole amounts of each ubiquitin link type [46]. Kirkpatrick et al. used SRM coupled with AQUA to reveal the complex ubiquitin chain topology consisting of a mixture of K11, K48 and K63 linkages and Xu et al. were able to unravel some of the critical functions of unconventional polyubiquitin chains in proteasome degradation [47,48]. Moreover, PRM was also utilised to quantify phosphorylated ubiquitin produced by the Parkinson's kinase PINK1 expressed in HeLa cells [49] and the effect of ubiquitin phosphorylation [50,51] on ubiquitin chain formation by E2/E3 ligases [52].

Quantitation of ubiquitin chain linkage peptides by SRM and PRM.

Figure 4.
Quantitation of ubiquitin chain linkage peptides by SRM and PRM.

To identify the ubiquitin chain linkages in a sample, polyubiquitylated proteins are digested, resulting in eight di-Gly remnant peptides of ubiquitin. Isotopically labelled internal peptide standards for AQUA can optionally be added to the mixture at a known concentration. This mixture is then separated by LC and analysed by SRM or PRM. In both MS modes, the peptide of interest is isolated in the first quadrupole (Q1) and then fragmented in the collision cell. While in SRM a single peptide fragment is quantified in quadrupole Q3, many more fragments are simultaneously quantified in PRM, thereby improving quantitation.

Figure 4.
Quantitation of ubiquitin chain linkage peptides by SRM and PRM.

To identify the ubiquitin chain linkages in a sample, polyubiquitylated proteins are digested, resulting in eight di-Gly remnant peptides of ubiquitin. Isotopically labelled internal peptide standards for AQUA can optionally be added to the mixture at a known concentration. This mixture is then separated by LC and analysed by SRM or PRM. In both MS modes, the peptide of interest is isolated in the first quadrupole (Q1) and then fragmented in the collision cell. While in SRM a single peptide fragment is quantified in quadrupole Q3, many more fragments are simultaneously quantified in PRM, thereby improving quantitation.

One limitation of these bottom-up proteomic approaches using proteases is that the stoichiometry, order and length of the chains is lost, which makes the study of mixed linkage types particularly difficult. An emerging solution is UbiCRest, which utilises linkage-specific DUBs in parallel to identify the different linkage positions and build the structure of complex polyubiquitylated chains [53]. Combination of this technology with quantitative MS will be an exciting prospect as it will allow for the identification of chain linkages from complex biological samples.

Assaying DUB and E3 ligase activity

DUBs have been identified as key drug targets as they are dysregulated in diseases such as cancer, autoimmunity and Parkinson's disease [5456]. In particular, USP7 appears to be an attractive target as its inhibition stabilises the tumour suppressor p53 and highly specific inhibitors have been published recently ([57,58]; Kategaya, L., Di Lello, P., Rougé, L., Pastor, R., Clark, K.R., Drummond, J., accepted unpublished work). As multiple DUBs often process the same substrate, assigning an activity to a particular enzyme is difficult. A novel solution has been the development of activity-based probes (ABPs) that mimic the substrate and covalently attach to the active site in an enzyme-catalysed reaction (Figure 5). As they only bind to active enzymes, they provide an accurate readout for the activity of DUBs. Several ABPs have been successfully applied to probe the ubiquitin system such as Ub-Aldehyde probes (Ubal) [59], Ub-vinyl methyl ester probes (UB-VME) [60], Ub-vinyl methyl sulphone probes (UB-VS) [60,61], as well as branched and ubiquitin isopeptide activity-based probes [62]. More recently, diubiquitin activity probes [63] have been developed to identify DUBs, explore their chain linkage preference and characterise their DUB enzymatic activity. These technologies provide very useful tools to analyse the selectivity and potency of inhibitors in cells. In such an experiment, cells are incubated with the ubiquitin ABP +/− inhibitor. Tags on ABPs allow selective pull-downs, enriching modified DUBs which are then analysed by MS. DUBs whose active sites were blocked by an inhibitor will not bind to any of the ABPs, and thus, the relative difference will give an indication about the activity and specificity of the inhibitors. As this process is dose-dependent, one can use this approach to measure enzyme and inhibitor kinetics such as IC50 [64,65].

Quantifying activity of DUBs and their inhibitors by activity-based probes and proteomics.

Figure 5.
Quantifying activity of DUBs and their inhibitors by activity-based probes and proteomics.

(A) To test for inhibitor potency and selectivity, cells are treated with a DUB inhibitor or DMSO (control). (B) A tagged ubiquitin ABP is added that covalently attaches to all active DUBs, but not to DUBs whose active site is blocked by inhibitors. (C) Next, DUBs are enriched by a specific pull-down of the tagged ubiquitin ABP, and then (D) digested and analysed by quantitative proteomics. DUBs binding to the inhibitor are lost and can be identified by comparing intensities of DUBs in control and inhibitor-treated samples.

Figure 5.
Quantifying activity of DUBs and their inhibitors by activity-based probes and proteomics.

(A) To test for inhibitor potency and selectivity, cells are treated with a DUB inhibitor or DMSO (control). (B) A tagged ubiquitin ABP is added that covalently attaches to all active DUBs, but not to DUBs whose active site is blocked by inhibitors. (C) Next, DUBs are enriched by a specific pull-down of the tagged ubiquitin ABP, and then (D) digested and analysed by quantitative proteomics. DUBs binding to the inhibitor are lost and can be identified by comparing intensities of DUBs in control and inhibitor-treated samples.

Similar approaches for E3 ligases were only recently developed and involve attaching a cysteine-reactive chemical group to the C-terminus of ubiquitin, which allowed measurement of the transfer of ubiquitin to the RING-in-between-RING E3 ligase Parkin [66]. The potential to study DUB and E2/E3 ligase activity in cells makes this technique a powerful approach to identify selective inhibitors, making it a valuable tool for drug discovery.

MS tools for drug discovery in the ubiquitin system

The development of DUB inhibitors and screening assays has previously involved the use of non-physiological substrates, including linear fusion of ubiquitin to a reporter protein or fluorogenic reporters [6769], which have been used to describe a range of small-molecule DUB inhibitors. However, these artificial substrates do not contain physiological ubiquitin isopeptide bonds and are not suitable for assessing the linkage specificity of DUBs. Their usage may therefore lead to false positive or false negative compound identification. To overcome the shortcomings of chemical probes, it is possible to screen DUB activity and specificity with physiologically related diubiquitin molecules [70] that are cleaved into monoubiquitin using SDS–PAGE methodology [71]. However, the throughput using this technology is notoriously low and not suitable for screening of large compound libraries. Recently, a novel technique for label-free DUB screening utilising high-throughput matrix-assisted laser desorption ionisation (MALDI) time-of-flight (TOF) MS was developed in our laboratory [26]. This method uses unmodified diubiquitins of all eight chain linkages or monoubiquitylated substrate [72] and 15N-labelled ubiquitin as an internal standard (Figure 6). Compared with SDS–PAGE assays, the MALDI-TOF DUB assay requires significantly less amount of diubiquitin substrate and is dramatically faster and more sensitive. This allowed to profile the activity and specificity of 42 different human DUBs and to characterise the potency and selectivity of many DUB inhibitors against a panel of 32 DUBs. Recent developments of ultra-fast MALDI-TOF mass spectrometers, which can screen 1536 samples in <10 min, have led to applications of this technology for other enzyme classes such as histone deacetylases [73], kinases [74] and E2/E3 ligases (De Cesare et al. in preparation). The great potential of label-free high-throughput MALDI-TOF MS in drug discovery will likely make this technology instrumental for the discovery of inhibitors in the ubiquitin system.

The MALDI-TOF DUB assay for drug discovery.

Figure 6.
The MALDI-TOF DUB assay for drug discovery.

To screen for DUB inhibitors, DUBs are incubated with a diubiquitin isomer with (A) or without (B) chemical compounds, leading to subsequent cleavage of diubiquitin into monoubiquitin. The reaction is stopped by the addition of trifluoroacetic acid (TFA) and 15N-labelled ubiquitin is added as an internal standard. The sample is then spotted onto a MALDI target and analysed by MALDI-TOF MS, and the ratio of light and 15N-labelled ubiquitin can be used to determine differences in DUB activity between control and inhibitor-treated reactions. (C) For drug discovery, all these steps are performed by liquid handling robots in a 1536-well format.

Figure 6.
The MALDI-TOF DUB assay for drug discovery.

To screen for DUB inhibitors, DUBs are incubated with a diubiquitin isomer with (A) or without (B) chemical compounds, leading to subsequent cleavage of diubiquitin into monoubiquitin. The reaction is stopped by the addition of trifluoroacetic acid (TFA) and 15N-labelled ubiquitin is added as an internal standard. The sample is then spotted onto a MALDI target and analysed by MALDI-TOF MS, and the ratio of light and 15N-labelled ubiquitin can be used to determine differences in DUB activity between control and inhibitor-treated reactions. (C) For drug discovery, all these steps are performed by liquid handling robots in a 1536-well format.

Conclusion

Ubiquitin regulates most biological processes of eukaryotic life, including proteasomal and lysosomal protein degradation, cell division, autophagy and cell signalling. Aberrant regulation of these processes lead to many different diseases including cancer, autoimmunity and neurodegeneration. The complexity of the ubiquitin system with eight different ubiquitin chain types that can appear in mixed or branched populations can only be partly unravelled by classical biochemical tools. Current MS methods provide a powerful toolsets to generate significant insights into molecular functions of the ubiquitin system. It will be important to utilise these methods to answer important biological questions. However, it will be equally important to develop novel methods. Novel approaches could employ ion mobility MS to study the structures and formation of different chain types and mixed chains, exploit subcellular proteomics and MALDI imaging MS to study the role of ubiquitin in vesicle trafficking and further advance high-throughput MALDI-TOF MS for the development of inhibitors for DUBs and E3 ligases. It is expected that the potent combination of biochemical, structural, cell biological and MS-based approaches will further our understanding of the role of ubiquitin in biology and will be fundamental in advancing the treatment of human disease.

Abbreviations

     
  • ABP

    activity-based probe

  •  
  • AQUA

    absolute quantification

  •  
  • DUB

    deubiquitylases

  •  
  • ESI

    electrospray ionisation

  •  
  • HPLC

    High-Performance Liquid Chromatography

  •  
  • LC

    liquid chromatography

  •  
  • MALDI

    matrix-assisted laser desorption/ionisation

  •  
  • MS

    mass spectrometry

  •  
  • PRM

    parallel reaction monitoring

  •  
  • SRM

    selected reaction monitoring

  •  
  • TOF

    time of flight

  •  
  • TUBEs

    tandem ubiquitin-binding entities

  •  
  • UBL

    ubiquitin-like modifier

Funding

R.E.H. and M.S.G. are funded by BBSRC CASE studentships with Bruker Daltonics and Thermo-Fisher Scientific, respectively. F.L. is a European Molecular Biology Organization long-term fellow [ALTF 841-2016]. J.P. and M.T. are funded through a generous start-up of Newcastle University.

Acknowledgments

We thank Katharina Trunk, Anetta Härtlova and Paul Dean for critical proof reading.

Competing Interests

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

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