Simple chemical tools to expand the range of proteomics applications

J O U RN A L OF P R O TE O MI CS 74 ( 20 1 1 ) 1 3 7–1 5 0

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Review

Simple chemical tools to expand the range of proteomics applications M a Jesús García-Murria, M a Luz Valero, Manuel M. Sánchez del Pino⁎ Laboratorio de Proteómica, Centro de Investigación Príncipe Felipe, Avda, Autopista del Saler 16, 46012 Valencia, Spain

AR TIC LE I N FO

ABS TR ACT

Article history:

Proteomics is an expanding technology with potential applications in many research fields.

Received 10 August 2010

Even though many research groups do not have direct access to its main analytical

Accepted 3 November 2010

technique, mass spectrometry, they can interact with proteomics core facilities to incorporate this technology into their projects. Protein identification is the analysis most frequently performed in core facilities and is, probably, the most robust procedure.

Keywords:

Here we discuss a few chemical reactions that are easily implemented within

Chemical modification

the conventional protein identification workflow. Chemical modification of proteins with

Protein structure

N-hydroxysuccinimide esters, 4-sulfophenyl isothiocyanate, O-methylisourea or through

De novo sequencing

β-elimination/Michael addition can be easily performed in any laboratory. The reactions

Phosphorylation

are quite specific with almost no side reactions. These chemical tools increase considerably

O-glycosylation

the number of applications and have been applied to characterize protein–protein interactions, to determine the N-terminal residues of proteins, to identify proteins with non-sequenced genomes or to locate phosphorylated and O-glycosylated. © 2010 Elsevier B.V. All rights reserved.

Contents 1. 2.

3.

Introduction . . . . . . . . . . . . . . . . . . . . . . . Chemical tools . . . . . . . . . . . . . . . . . . . . . . 2.1. Labelling with N-hydroxysuccinimide esters . 2.2. Sulfonation with 4-sulfophenyl isothiocyanate 2.3. Guanidination . . . . . . . . . . . . . . . . . . 2.4. β-elimination/Michael addition . . . . . . . . . Applications . . . . . . . . . . . . . . . . . . . . . . . 3.1. Protein structural information . . . . . . . . . 3.2. Aids for de novo sequencing . . . . . . . . . .

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⁎ Corresponding author. Centro de Investigación Príncipe Felipe, Avda. Autopista del Saler, 16, E-46013 Valencia, Spain. Tel.: +34 963289680; fax: +34 963289701. E-mail address: [email protected] (M.M. Sánchez del Pino). 1874-3919/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.jprot.2010.11.002

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3.3. Enhancing the ionization efficiency of lysine 3.4. PTM: phosphorylation and O-glycosylation . 4. Conclusions . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . .

1.

containing peptides. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Introduction

Proteomics is a very active and expanding field of research. The combination of bioinformatics tools, the increasing number of sequenced organisms and advances in mass spectrometry technology have made proteomics possible, as we know it nowadays. There are plenty of research groups that are doing an excellent work pushing forward the development of methodologies and applications of proteomics. As a result, many researchers could benefit from the great potential of proteomic techniques either in their own laboratories or in collaboration with other laboratories, such as core facilities, equipped with the required infrastructure. The core facilities currently available in many research centres and universities play an important role in the widespread use of proteomics because they make it accessible to many researchers. However, a quick survey of proteomics core facilities web pages show that virtually all of them provide services for protein identification, many of them also provide quantitative information but not many provide regular services for other type of analysis. An important reason for this may be the difficulty to provide results in the shortest period of time possible from a large number of highly diverse samples. Working under these conditions requires the adjustment of very robust and wellestablished procedures to handle samples from different tissues, experimental procedures or protein concentration. Protein identification is the proteomic analysis most frequently performed in core facilities and is an example of a very well established procedure. These experiments are carried out very successfully in most laboratories where the performance is determined mainly by the equipment rather than by the procedure. Thus, it would be useful to widen the range of proteomic applications by just introducing minor modifications to the conventional and robust workflows used for protein identification. Chemical modification of proteins could be considered one of the strategies to pursue this goal. Although protein chemistry laboratories have been modifying proteins for many years, its combination with current proteomics techniques (mass spectrometry and bioinformatics tools) has allowed researchers to exploit the potential of these strategies more easily than it was in the past. There are many reasons to modify peptides or proteins chemically and to take advantage of the new properties introduced by the modification in processes such as purification, stabilization or identification. The introduction of a tag can modify the chromatographic behaviour, alter the spectroscopic properties or introduce a reporter ion to allow its identification. For example, the introduction of a biotin tag can be used for affinity purification of the peptides of interest. Similarly, the identification of the tag can be used to obtain information about the modified residue such as its position within the

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protein sequence or its accessibility/exposure to the solvent. Of course, all these additional information have a cost. Each chemical reaction introduced in the workflow implies the use of more reagents and time. Moreover, the overall yield of the procedure would probably decrease so that a larger amount of samples would be required, which is often difficult or not possible. In addition, since there are always some nondesirable reactions, the complexity of the results is increased and, hence, the analysis may be more difficult and time consuming. The main goal of this work is to review a small number of chemical reactions (chemical tools) that are relatively easy to integrate in the conventional proteomic workflow. They are simple well-established and quite specific reactions that usually have a reduce number of side-reactions. Some of them are very versatile since they can introduce many chemical functionalities. The procedures can be performed in any laboratory, not just proteomic laboratories, before its incorporation into the regular proteomics workflow. The experimental strategies where they are used and some of their applications to obtain additional information without too much additional work are also discuss.

2.

Chemical tools

2.1.

Labelling with N-hydroxysuccinimide esters

N-hydroxysuccinimide (NHS) esters are the most widely used amine-reactive compounds. They are used in most of the commercially available reagents to label, modify or crosslink amines, including iTRAQ and DIGE labelling reagents. Nhydroxysuccinimide can be considered as a cargo carrier to transfer chemical labels to amino groups in proteins. The NHS esters derivatives react at physiological pH with nucleophiles to form acylated products according to the reaction (Fig. 1A). The reaction can take place with several functional groups but only amines form stable conjugates [1]. Thus, in proteins, mostly Nterminal and lysine residues react with NHS esters to form stable amides. The unwanted modifications of serine, threonine, and tyrosine residues that have sometimes been observed [2–4] may be reverted by treatment with hydroxylamine at alkaline pH [4] or by boiling the sample [5]. The reaction with amines competes with the hydrolysis of the NHS ester in aqueous solutions, which increases at basic pH. Thus, a working pH range of 7–9 is widely used to balance between unprotonated reactive amines and hydrolysis. Since this pH range preserves native conformation of proteins, it is convenient to perform structural studies. The pKa value for the ε-ammonium group of a lysine residue is approximately 11 while that of an N-terminal α-ammonium group is more frequently below 8 [6]. Thus, some

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A

O

O Protein NH2 + Label

Protein

O N

N

+

Label

HO N

H

O

B

O

O

O

O -

O S

N C S

S

O -

O

O S

N

N Peptide

O

H

H

+ Peptide NH2

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O

O NH2

R2 R1

R1

+ O

NH2

N

R2

NH2

NH2

+ H3C OH

NH

D

O R1

N X

O

O R2

O

Base

R1

N

R2

Label

SH

R1

N X

X

PTM

R2 S Label

Fig. 1 – Reaction schemes of the chemical tools. A. Labelling reaction with N-hydroxysuccinimide (NHS) esters. The NHS ester activated label is transferred to an amino group of the protein. B. Sulfonation reaction with 4-sulfophenyl isothiocyanate (SPITC). The α-amino of the peptide reacts with SPITC to form a thiocarbamoyl derivative. C. Guanidination reaction. The α-amino of a lysine residue reacts with O-methylisourea to produce a homoarginine residue. D, β-elimination and Michael addition reactions. Under basic conditions, the phosphoryl or O-glycosyl moieties (indicated as PTM) are eliminated to form an α,β unsaturated residue. This residue easily reacts with thiol-containing labels, which are incorporated where the modification was initially located. X stands for H (serine residues) or CH3 (threonine residues).

selectivity may be achieved by selecting the appropriate reaction pH. For instance, a reaction buffered at pH 8.0–8.2 has been described to specifically modify N-terminal amino groups [7]. Parameters such as pH, temperature, time and the NHS ester/amine ratio may require optimization for each reaction. Many NHS esters are relatively insoluble in aqueous buffers so that concentrated stock solutions in organic solvents such as DMF or DMSO are usually prepared. To avoid these inconveniences, the more water-soluble sulfo-NHS derivatives are frequently used nowadays. In addition, these compounds have a longer life in aqueous buffers and keep similar reactivity and specificity [8]. There are a large variety of compounds that contains NHS esters to react with protein amines. As mentioned, iTRAQ and DIGE reagents react with N-terminal and lysine residues of peptides and proteins, but there are also activated beads of different materials designed to bind proteins and peptides using NHS esters. Biotin tags, crosslinkers, fluorinated tags, etc. can be used for many purposes. Protocols for performing the reaction in solid-phase (C18 μZipTip) have also been described [4].

2.2.

Sulfonation with 4-sulfophenyl isothiocyanate

Phenylisothiocyanate (PITC, Edman's reagent) have been used in protein chemistry for a long time to perform peptide sequencing. It reacts with amino groups to form a thiocarbamoyl derivative. In the case of the α-amino group there are further rearrangements with the carbonyl group of the first peptide bond that leads to the cleavage of the N-terminal amino acid. The mechanism involved in Edman degradation is very similar to that of the formation of b-type ions during peptide fragmentation in a mass spectrometer [9]. Since the presence of a negative charge in the modifying reagent has some advantages for the analysis of the derivatized peptides by mass spectrometry, 4-sulfophenyl isothiocyanate (SPITC) is more frequently used in proteomics applications (Fig. 1B). The initial description of its use to aid the interpretation of MALDIPSD spectra was made by Gevaert et al. [10]. Although the fragmentation spectra of the derivatized peptides were much simpler and easy to interpret, the authors did not recommend its general use mainly because of an overall low sensitivity.

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However, several improvements of the original protocol have been described that allow the implementation of this derivatization reaction in regular proteomics workflows [11–14]. Initially, Marekov et al. used organic solvents in the media and obtained a quantitative modification of the peptides extracted from gel bands [13]. However, it was later reported that even though the organic compounds improve the yield of the reaction, a higher total intensity of the modified peptides could be obtained without them [14]. Wang et al. studied the influence of different components on the reaction and the best results were obtained using 20 mM sodium bicarbonate at pH 9.5 [14]. The pH of the reaction mixture and the SPITC/peptide ratio are also important. Specific modification of the Nterminal residues, regardless of the presence of lysine at the C-terminal end of the peptide, was achieved controlling the reaction pH at around 8.0 and using a relatively low SPITC/ peptide ratio (5–15 fold SPITC excess) [12]. Derivatization of large peptides is sometimes incomplete, probably due to intramolecular solvation of the α-amino. There are other protocols to perform the reaction with the sample immobilized in C18 μZipTip [11] or directly on AnchorChip targets [15] that could be convenient in some applications.

2.3.

Guanidination

Guanidination has been used in protein chemistry for long time [16] and a role in stabilizing proteins has been documented [17]. Although it was initially reported some reaction with α-amino groups (see for instance Ref. [18]) and a slight reactivity toward N-terminal glycine residues have been detected [19,20], the reaction is virtually specific for ε-amino groups of lysine residues. Guanidination reaction takes place with high yield in aqueous solutions in a single step so that it can be easily implemented in the regular proteomics workflow (Fig. 1C). The reaction is quite simple; usually a concentrated solution of O-methylisourea at basic pH is added to the peptides or proteins and incubated for certain time. Older protocols used low temperature and the reaction was performed over several days because the protein activity had to be preserved. However, in proteomics experiments where maintaining protein activity is not a concern, more stringent conditions are used so that the reaction is completed in 10– 20 min in the presence of about 1 M O-methylisourea at 65 °C [19]. The basic pH is maintained by adding ammonia to a final concentration of about 3.5 M. The reaction has also been adapted to derivatize proteins contained in gel pieces [21] as well as peptides bound to C18 ZipTips [22]. There are reports where guanidination of proteins reduced trypsin cleavage efficiency [21,23]. However, it does not seem to be a serious drawback because you can still obtain valuable information while avoiding the desalting step to remove guanidination reagents. Thus, this approach can be very convenient for some experimental strategies. The basic pH used in the guanidination reaction may enhance deamidation of asparagine and glutamine residues [24].

2.4.

β-elimination/Michael addition

It is well established that under strong alkaline conditions, substituted serine and threonine residues undergo β-elimination

to form dehydroalanine (ΔSer) or dehydro-2-aminobutiryc acid (ΔThr), respectively [25]. The resultant α,β unsaturated residues are potent Michael acceptors, which can readily react with a nucleophile [25–27] (Fig. 1D). This reaction allows the specific labelling of serine and threonine residues containing posttranslational modifications with a large variety of molecules. This analytical approach has been called chemically targeted identification (CTID) [28], and was first introduced in proteomics by Meyer et al. to sequence phosphoproteins by Edman degradation [29]. Since then, many different nucleophilic agents and chemical conditions have been used to modify peptides in order to improve the analysis of the products by mass spectrometry. The specific conditions of the reaction depend on which structure is the focus of the analysis because the requirements to preserve the integrity of the peptide backbone or the O-glycan moiety are different. Initial conditions using very high alkaline media with long incubations times and high temperature produced some peptide degradation, mainly under reductive conditions needed to preserve O-glycans. Thus, several improvements have been described to moderate these parameters and to adapt the reaction to proteomics requirements. Other important factors to take into account are the base and the buffer composition. The use of Ba(OH)2 was introduced by Byford [30] to specifically catalyze β-elimination of phosphopeptides. However, this selectivity toward phosphopeptides has been shown to depend on the reaction conditions. Thus, with high Ba(OH)2 concentration there is no difference between phospho- and glycopeptides at room temperature [28] whereas more than 12fold selectivity toward phosphopeptides has been achieved using 0.015 M Ba(OH)2 for 10 min at 45 °C [31]. Nevertheless, Ba(OH)2 is frequently used to perform a more specific β-elimination of phosphopeptides [28,30–46]. Reaction conditions have also been described where β-elimination of glycopeptides takes place at a much faster rate than phosphopeptides [40,47]. When using Ba (OH)2 care should be taken to minimize the presence of counterions in the sample that could either precipitate or neutralize the base [28]. Using a C18 reversed-phase micropipette is a convenient and practical way to desalt the peptides before the elimination reaction. After the β-elimination reaction, the excess of base can be removed by adding ammonium sulfate [41] or solid carbon dioxide where no peptides could be detected in the precipitate [34]. The solvent composition also plays an important role and a decrease in the yield has been reported with increasing amounts of polar, hydroxylic components [40]. On the contrary, aprotic co-solvents such as DMSO have been frequently used to improve the yield of the modification and to prevent peptide hydrolysis [32,37,40,41]. The Michael addition is more frequently performed simultaneously with the β-elimination reaction but it can also be carried out as consecutive reactions [28,34,37,48]. Many labelling tags have been used for the derivatization of the α,β unsaturated residues, mainly thiols. Most of them are relatively small neutral alkanethiols like mercaptoethanol, [29,49], ethanedithiol [32,50–54], dithiothreitol [34,44,46,47,55,56], and others [29,48,57,58]. Charged thiols such as aminoethanethiol (AET) [28,37,39,41], dimethylaminoethanethiol [59], mercaptoethylpyridine [35] or guanidinoethanethiol (GET) [33,60] have also been used. Although the β-elimination/addition reaction is most frequently performed in solution after protein digestion, depending on the strategy, it can also take place on immobilized peptides [41,60] or at the protein

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level, either in solution [32,37,51,53,54,58] or in gel bands [35,48,59,61,62]. There are some potential problems associated with β-elimination/addition reactions that should be considered. Cysteine residues can also react and have to be blocked before the β-elimination reaction. However, this is not a serious inconvenient since reduction and alkylation of cysteines are usually included in any conventional proteomics workflow. In some cases the diastereoisomers formed after the Michael addition can be resolved in the RP-HPLC prior to the mass spectrometry analysis [63]. Although this is an undesired side effect due to the increase in complexity and the probable decrease of the signal, it also has the positive effect of increasing the chances to identify the peptide due to the longer analysis time [58]. The reaction conditions usually favour deamidation of asparagines and glutamine residues [64] so that this modification should be included in the searching parameters. A small percentage (1–2%) of β-elimination of free serine residues has also been reported [52]. Overall, the advantages of the β-elimination/addition procedures overcome their disadvantages and it is, for instance, the only way to extract O-glycans without artefacts. Thus, β-elimination is the most frequently used experimental strategy to study the O-glycosylation of IgA1 within the Human Proteome Organisation Human Disease Glycomics/Proteome Initiative [65].

3.

Applications

3.1.

Protein structural information

Since NHS esters are the most widely used reagents to modify amino groups they have been used in many different applications. A very common application is protein quantitation. Many of the reagents used for quantitation, including the most popular commercial reagents such as iTRAQ, CyDyes used in DIGE, TMT, or ICPL, are NHS esters. Other important but less common application of NHS containing compounds is the structural characterization of proteins. To understand protein function it is important to know its structure and the surface interacting with other proteins or molecules such as nucleic acids. X-ray crystallography and NMR spectroscopy are the most powerful techniques to determine protein structure. However, many proteins are not amenable to these approaches. There are several reasons for this but protein solubility at the required concentration is an important issue. In these cases, alternative methodologies capable of providing structural information may be very valuable. The combination of old protein chemistry strategies with mass spectrometry has been proven useful as a low-resolution three-dimensional methodology [66]. The use of cross-linking molecules allows placing spatial restrictions to unknown structures. Although they could, in principle, provide substantial information, data analysis is complicated because of the considerable number of possible combinations and side product reactions. Thus, several approaches to cope with these difficulties are being developed both technical as well as computational. There are some excellent reviews on crosslinking proteomics [67,68]. A simpler scenario is when the goal is not determining the three-dimensional structure of the protein but rather characterizing their interactions with other

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molecules. In these cases, the accessible surface of a protein or complex can be probed using reagents that modify specific residues with a simple experimental approach. Lysine residues are very appropriate targets because of their abundance and because they are usually located at the surface of the protein due to their polarity. Due to their specificity and versatility, and because they can be used in conditions that preserve native protein structure, NHS esters are the most frequently used reagents for this purpose. The simplest strategy consists in labelling the protein in the presence and in the absence of the interacting protein. Thus, in the presence of the interacting protein, the contact surface will be inaccessible to the reagent and will not be labelled (Fig. 2). Depending on the strategy, the properties of the label can be selected to simplify the purification or the detection of the

Fig. 2 – Outline of a basic strategy to identify protein–protein interaction surfaces. A. There are a number of lysine residues, generally located on the protein surface due to their polarity. Upon interaction with other proteins some of these residues may become occluded or less accessible. B. NHS esters can be used to label lysine residues under experimental conditions that preserve native protein structure. Only accessible residues are labelled. C. A conventional proteomic analysis (digestion and MS analysis) shows the degree of modification of lysine residues. Comparison between experiments performed with isolated proteins and with protein complexes reveals the relative accessibility to the solvent of each residue.

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peptides of interest. This approach has been used to investigate protein:nucleic acid interactions introducing biotin tags [69,70] or protein:protein interactions acetylating lysine residues via NHS- or sulfoNHS-acetate [71,72]. Peptides containing acetylated lysine can be easily identified by monitoring the presence of the immonium ion of 126.1 Da produced by the modified residue [71]. Similarly, the solvent accessibility of lysine residues in proteins has been studied using NHSacetate to acetylate amino groups [73]. A dual labelling strategy was developed to identify heparin-binding sites in proteins [74]. In this approach, proteins are bound to heparin beads and lysine residues are acetylated with sulfoNHSacetate. Subsequently, proteins are released from heparin beads and labelled with NHS-biotin. The biotin tag is used to purify the peptides from the binding site. In this strategy, the acetylation step is the key to increase the signal to noise ratio of the binding site labelling. Another application of NHS esters is terminal [75] or positional [76] proteomics, where the N- or C-terminal sequences of proteins are determined. The basic strategy usually combines a first labelling step of amino groups at the protein level, followed by digestion and a second labelling step of the newly generated α-amino groups (Fig. 3). As usual, reduction and alkylation of cysteine residues are included in the initial steps of the procedure. After that, if the strategy is exclusively based on the modification of α-amino groups, a lysine blocking step can be introduced, which is generally done by guanidination. The properties of the tag introduced at the amino groups are usually carefully selected to provide a clear advantage in the purification or the identification of the peptides of interest. In the most intuitive and simplest strategy, the amino groups of proteins or fragments (after a limited enzymatic or CNBr proteolysis experiment) are modified with Tag1. Since the purpose of this tag is, usually, to block the amino groups of the protein, a simple acetylation procedure using sulfoNHS-acetate is frequently used [77–79]. Sometimes, the tag is also selected to improve fragmentation in the mass spectrometer and the peptide identification rate [80–82]. Proteins are then digested to generate peptides containing new free α-amino groups, which can be separated from the blocked N-terminal peptides by reacting with amino scavenger beads [78,80,81,83,84]. Alternatively, labelling free amino groups with a tag that alters their chromatographic behaviour [77,79] or that allows affinity purification [76] has also been used. A modification of this basic strategy was developed by Kuyama et al. that allowed them to purify N-terminal or C-terminal peptides of proteins by just changing the protease [83,84]. In this approach, proteins are digested with Lys-C or Lys-N before any treatment. After digestion, free α-amino groups are selectively labelled with NHS-Ac-TMPP. Under the reaction conditions used by the authors, lysine residues were not labelled. In the case of cleavage by Lys-C, all peptides will have lysine residues at their C-terminal positions except the C-terminal peptide of the protein. In the case of Lys-N cleavage the N-terminal peptide of the protein will be the only peptide without a lysine residue at the N-terminal side. Internal lysine-containing peptides are then removed using DITC (p-Phenylenediisothiocyanate) resin. In all these approaches, endogenous N-terminally modified peptides are also included in the analysis because the purification proce-

Fig. 3 – Outline of a general procedure for the identification of protein N-terminal residues. A. Proteins have an N-terminal residue that could be free or modified. The ε-amino groups of lysine residues may interfere with the labelling of the α-amino so that they are usually blocked by guanidination. B. α-amino groups can now be labelled. The labelling reagent (frequently containing NHS esters) can be selected to differentiate from naturally occurring modifications such as acetylation. C. Protein digestion with proteases produces peptides with new free α-amino groups, except for the peptide containing the protein N-terminal. D. A second labelling reaction of the newly formed α-amino groups can be performed with a different reagent. This reaction may not be necessary depending on the separation procedure to be used. E. The properties of the different N-terminal labels are used to separate N-terminal from internal peptides. Analysis by mass spectrometry of the isolated peptides containing the first label provides the identity of the protein and its N-terminal sequence.

dures were not based on the tag introduced at the protein level. In fact, these peptides can be specifically isolated with a similar approach by just omitting the first labelling step [85].

3.2.

Aids for de novo sequencing

Many proteomics experiments are conducted with organisms whose genome is not sequenced. In these cases, the most frequently used approach for protein identification is to obtain partial sequences from MS data and to identify similar proteins in databases by sequence homology. Correct interpretation of MS/MS spectra to obtain partial sequences is essential in this critical process. However, this is many times a difficult and

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time-consuming task despite the available software packages that help in this process with different success. Thus, a procedure that could help in the interpretation of MS/MS spectra would be very useful. Such experimental procedures exist and they can be implemented without too much interference in normal proteomics workflows. The use of charged labels to analyze peptides by mass spectrometry has been used for a long time [86]. Keough et al. clearly showed the usefulness of sulfonated compounds to facilitate fragmentation in MALDI-PSD analysis. Singly charged peptides need much higher energy for fragmentation than doubly or triply charged peptides. The reason for this is that, according to the mobile proton model [87], one proton is localized at the basic residue (lysine or arginine) while the other is mobile along the peptide backbone to assist fragmentation [88]. In the case of a singly charged peptide the only proton is anchored to the basic residue and more energy is required to fragment the peptide. However, two protons are required to obtain a singly charged peptide containing a fixed negative charge. One is required to neutralize the negative charge and the other to ionize the peptide. Thus, it is similar to doubly charged peptides so that fragmentation is facilitated by the presence of the negative charge. The negative charge at the N-terminal side provides an additional advantage. The N-terminal fragments retaining one proton, such as b-type ions, become neutral and are not detected by the mass spectrometer. Hence, the fragmentation spectrum of these derivatized peptides contains only C-terminal fragments like y-type ions, which makes the interpretation much simpler. Since this advantage is not observed in doubly charged peptides, this derivatization is more useful for MALDI-PSD or MALDI TOF/TOF analysis. After testing several compounds, Keough et al. found that 3-sulfopropionic acid NHS had the best properties for the derivatization of peptides to facilitate fragmentation. Although it is now discontinued, GE Healthcare commercialized this reagent as Ettan CAF (Chemically Assisted Fragmentation). There have been reports using alternative reagents to introduce a negatively charged label at the amino terminal site [89,90]. However, the reagent most frequently used is SPITC because it is economical, stable, commercially available, and produces similar or even better results than other reagents [22]. As mentioned before, this reagent promotes an Edman-type degradation that cleaves very efficiently the N-terminal residue. Thus, the most intense signal of the spectra usually corresponds to the yn-1 ion. The other y-ions are also easily recognized even though their intensity may be lower (Fig. 4). In the worst scenario, where only the yn-1 ion would be observed, the knowledge of the N-terminal residue would reduce the number of potential matches in a database search and, thus, would help protein identification [12]. After derivatization with CAF or SPITC a lower intensity of the MS spectra is usually observed. However, it does not affect the intensity of the MS/MS spectra, which is higher [22]. Since the advantages of introducing a negative charge in the peptide are fully exploited by MALDI mass spectrometry, the strategy is specially suited for the analysis of protein spots from 2D gels. There have been several reports where SPITC derivatization has been coupled to the conventional proteomics workflow where proteins were separated by 2D electrophoresis and the protein spots were identified by MALDI-MS. There are even protocols where peptides extracted from a

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Fig. 4 – De novo sequencing using SPITC. MALDI TOF/TOF spectra of a Lys-C peptide of Cyclin A2 before (A) and after (B) reaction with SPITC. Even though the spectrum of the unmodified peptide contains enough information to determine the sequence (APQHAQQSIREK), its interpretation is difficult. The spectrum of the peptide labelled with SPITC is quite simple and the sequence can be easily obtained from the y-ions. Fragmentation of SPITC-modified peptides produces the loss of the modifying group (i.e. removal of SPITC and sulfanilic acid) and the formation of yn-1 (y11 for this peptide). Usually, these three ions are the most intense peaks of the spectrum as shown in the right side of B.

silver stained 2D gel are bound to ZipTips to perform all the reactions [11]. Hjerno et al. [91] performed a DIGE experiment with strawberry samples and obtained a 58% success rate even though no genomic sequences were available. Their strategy to identify the spots of interest included the double derivatization of the spots were no positive identification was obtained with the conventional protocol. The peptides extracted from the 2D gel spots were guanidinated to block lysine residues before labelling the amino terminal groups with SPITC. Similarly, Oehlers et al. [92] performed a DIGE experiment with medaka (Oryzias latipes) to determine hypoxia biomarkers in the brain. Even though the same group had described a protocol to perform the SPITC reaction directly on the AnchorChip targets [93], they labelled the peptides in solution and were able to obtain enough peptide sequences to allow positive identification by BLAST for most of the spots of interest. Leon et al. [94] also reported an increase of up to 78% in the identification rate of proteins from a nematode with an unknown genome by labelling peptides with SPITC. They derivatized tryptic peptides of protein spots from a preparative 2D gel. As expected, SPITC labelling did not improve Mascot identification success rate. However, they were able to identify more proteins by homology search after manual interpretation of the MS/MS spectra. Almost no lysine containing peptides were identified, probably because they did not

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block lysine amino groups. Moreover, in some cases where no positive identification was obtained, the high confidence sequence data led the authors to discover new proteins. In this regard, the sequence information obtained with SPITC allowed Rinalducci et al. [95] to determine over 65% of the sequence of phycocyanin subunits isoforms from an organism whose genome was unsequenced. New applications for the use of SPITC, in combination with other proteomic techniques, have been described. Thus, although it is not commercially available, SPITC can be isotopically labelled to allow simultaneous quantitation of the identified peptides [96]. Also, a protocol to carry out ontissue derivatization has been developed to improve protein identification in the emerging field of MALDI imaging [97]. However, 3-sulfobenzoic acid succinimidyl ester performed better than SPITC in this application. In yet another application, SPITC has been shown useful to perform SRM experiments [98]. In this approach, it is easier to select specific transitions for peptides because SPITC-derivatized peptides produce almost complete y-ion series.

3.3. Enhancing the ionization efficiency of lysine containing peptides It has been reported that the most intense peaks in MALDI mass spectra correspond to peptides having C-terminal arginine residues [99,100]. The high proton affinity of arginine relative

to that of lysine may produce screening effects that would prevent the detection of lysine-terminated peptides when arginine-terminated peptides are present in the mixture. Increasing the basicity of lysine residues would increase the ionization efficiency of lysine-containing peptides and, thus, their intensity in MALDI spectra (Fig. 5). This was the rational to improve detection of lysine-containing peptides by converting lysine residues to the more basic amino acid homoarginine through guanidination [101–103]. Since then, it has been widely used to improve protein identification by MALDI mass spectrometry either alone [104,105] or in combination with other labelling reagents [20–22,81,82,106]. Although guanidination was reported to promote fragmentation in plasma desorption mass spectrometers [107], this effect was not observed later with MALDI instruments [106]. Nevertheless, guanidination of lysines improves the MALDI spectroscopic properties of peptide mixtures so that the number of peptides, the protein coverage and the identification scores are increased [105]. Most compounds used to facilitate de novo sequencing are amine-specific reagents designed to react with the N-terminal residue of peptides. Guanidination is frequently used to confer α-amino specificity to these reagents by blocking lysine residues. In addition to facilitate detection of lysine containing peptides by MALDI mass spectrometry, guanidination has some other advantages that help to improve protein identification. The increase in 42 Da due to guanidination allows discrimination between glutamine and lysine residues. Another advantage of the reaction is that the y1 ion of homoarginine (m/z 189) appears to be enhanced in both MALDI MS/MS and ESI MS/MS spectra compared with the y1 ion from unmodified lysine (m/z 147) [20]. When peptides are analyzed by electrospray ionization, the effect of guanidination apparently depends on the fragmentation procedure. Thus, guanidination enhance fragmentation by electron transfer dissociation (ETD), resulting in simpler spectra with higher peak intensities [108]. On the contrary, charge and ionization properties of peptides and, thus, the efficiency of ionization and overall signal intensity are unaffected by guanidination in ESI-LC/MS/MS if fragmentation is achieved via collision induced dissociation (CID) [109]. Indeed, this lack of effect has been exploited in a quantitation and de novo sequencing strategy based on the comparison of unmodified and guanidinated peptide spectra named mass-coded abundance tagging (MCAT) [109].

3.4.

Fig. 5 – Enhancing the ionization efficiency of lysine containing peptides by guanidination. A gel band of Cyclin A2 was digested with trypsin. The resulting peptides were analyzed by MALDI TOF directly (A) or after guanidination (B). Peptides that appear in both spectra are indicated with the broken lines. Triangles indicate peptides that contain lysine but not arginine residues. The increase in the intensity of the lysine containing peptides is clear. The median intensity of the lysine containing peptides is 6.3 and 25.4% of the maximum intensity in spectra A and B, respectively.

PTM: phosphorylation and O-glycosylation

Phosphorylation and O-glycosylation are among the most frequent and relevant posttranslational modifications of proteins in biology. Thus, there is a lot of interest in the study and characterization of these modifications in biological systems. The protein that carries the modification, the modified residue, and the amount of modified protein should be known to fully describe the process. Mass spectrometry is probably the technique most widely used to provide this information. However, there are difficulties to analyze phosphorylation and O-glycosylation by mass spectrometry mainly derived from the low abundance of modified peptides compare with non-modified ones and the instability of the linkage between the peptide and the modification. Identification of peptides and proteins containing both types of

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modifications has been facilitated by the development of specific enrichment and purification procedures. However, even in such cases, it is difficult to determine the precise location of the modification due to several factors. The highly acidic phosphate moiety may be the reason for the lower ionization efficiency of these peptides. In addition, phosphopeptides [110] and glycopeptides [111] show a high propensity towards neutral loss of the side chain of the modified amino acid so that the MS/MS analysis of these peptides yields little more than the neutral loss of the precursor ion. Thus, in many occasions the fragmentation spectra of these peptides do not contain enough information to determine the site of the modification, which is an important information to understand the biology of the modification. The main approach to circumvent the later difficulty is to devise a chemical strategy to stabilize the peptides and to obtain the required informa-

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tion. The procedure most frequently used is the removal of the attached moiety by β-elimination. Afterwards, a tag is introduced where the initial modification was located. The tag, unlike the phosphate or glycan moiety, is stable under the mass spectrometer and produce useful sequence information to determine the location of the modification [58,112]. The addition reaction is highly flexible in terms of the molecule that is being attached and, thus, to devise many experimental strategies with different purposes such as purification, quantitation, identification or combinations of them (Fig. 6). Identification of the modified residue using β-elimination has been carried out using different experimental strategies. One approach is to create a new protease cleavage site. This has been achieved by introducing 2-aminoethanethiol, which mimics a lysine residue. This approach has been successfully applied for phospho- and glycopeptides [37,39]. The strategy

Fig. 6 – Identification of phosphorylated and O-glycosylated residues. A. Peptides containing these modifications (circle) usually provide little sequence information by MS/MS spectrometry, frequently due to neutral loss of the modification. After β-elimination and Michael addition, these modifications are substituted by new chemical entities (diamond) with beneficial properties some of which are depicted in the figure. B. The more stable bond of the new label produces highly informative fragmentation spectra in the mass spectrometer. C. There are labels that produce specific reporter ions with high signal intensity. The modified peptides are then easier to identify even in the presence of unmodified peptides. Specific monitoring can be achieved with precursor ion scanning experiments. D. The properties of the label can be exploited to devise a specific purification procedure for modified peptides.

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requires previous affinity purification or specific enzymatic treatment to achieve specificity and a lysine-blocking step. After these steps, specific cleavage can be obtained with proteolytic enzymes such as Lys-C. The protease cleavage efficiency, at least with trypsin, is reduced in these samples because cleavage only occurs in one of the diastereoisomers produced during the addition reaction. However, this potential drawback is considered as an advantage for peptide mapping because the generated miscleavages allow the assignment of successive phosphorylation sites [37,39]. In addition to the specific cleavage site, the modified residues produce a highly abundant characteristic y1-ion at m/z of 165.07, which may increase the sensitivity to detect the peptides of interest in complex mixtures by precursor ion scanning. The presence of this ion allows the direct identification of both the peptide and the position of the modification even in the case of low MS/MS information. In other approaches, the tag is selected to produce a specific reporter ion that can be used to identify phosphopeptides [35,59]. Similarly, the introduction of a tag that mimics arginine improves the detection by MALDI because the intensity of the modified peptides is highly increased [33]. Other strategies have used β-elimination/addition to tag the modified residues in such a way that they could be affinity purified in subsequent steps. Initial procedures of this approach used dithiol molecules after blocking cysteine residues. In this way, new thiol groups were introduced that could subsequently be labelled with biotin or other compounds for affinity purification [36,53]. Although this strategy was successfully applied, biotin tags interfered with the analysis by mass spectrometry and were later substituted by purification approaches based on thiol exchange mechanisms. This simpler strategy does not require the second tagging reaction. Moreover, the tag does not interfere with the fragmentation process so that it is usually possible to obtain enough information to identify the position of the modification [34,47,52]. The covalent attachment to functionalized beads with 2-mercaptoacetyl-hexanoyl has also been described to specifically purify phosphopeptides [42]. In this approach phosphopeptides are covalently bound to the resin and, after extensive washing, the bound peptides are released with acid. Even though phosphotyrosine residues do not experiment β-elimination, the reaction can still be useful to identify these residues [113]. In this case, the phosphate groups from serine and threonine residues are removed by β-elimination so that after the treatment only phosphotyrosine containing peptides are purified by IMAC. Finally, the use of tags containing stable isotopes allows quantification. Thus, in addition to identify and purify, the simultaneous quantitation of phosphopeptides or glycopeptides is also possible [32,34,44,51,54,57].

4.

Conclusions

Protein identification is the most well established procedure in proteomics with a robust and reliable workflow. Besides protein identification, some simple chemical tools can be incorporated into this conventional workflow to obtain additional information. These chemical tools are very specific reactions used in protein chemistry laboratories for many years. Although they may

require some adjustment for the implementation as a routine procedure in the laboratory, they have been used widely in a variety of applications in a relatively simple way. Overall, their benefits compensate for the drawbacks that any additional chemical reaction may introduce. The range of applications goes from improvements of the spectral properties to aid de novo sequencing for working with organism with non-sequenced genomes to the identification of the precise location of posttranslational modifications or the structural characterization of proteins. These chemical tools may be of interest to many of the researchers that use proteomics as one of the available technical tools in their projects. The procedures could be performed either in the proteomic user laboratory or in a proteomics core facility as a routine service.

Acknowledgements The work has been funded by grants GV05/211 from the Generalitat Valenciana and PI061718 from the Instituto de Salud Carlos III. The Laboratorio de Proteómica from the Centro de Investigación Príncipe Felipe belongs to the Spanish National Institute of Proteomics, ProteoRed.

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