Protein processing and other modifications analyzed by diagonal peptide chromatography

Biochimica et Biophysica Acta 1764 (2006) 1801 – 1810 www.elsevier.com/locate/bbapap

Review

Protein processing and other modifications analyzed by diagonal peptide chromatography Kris Gevaert ⁎, Petra Van Damme, Bart Ghesquière, Joël Vandekerckhove Department of Medical Protein Research and Biochemistry, Flanders Interuniversity Institute for Biotechnology and Faculty of Medicine and Health Sciences, Ghent University, A. Baertsoenkaai 3, B-9000 Ghent, Belgium Received 6 June 2006; received in revised form 6 September 2006; accepted 6 September 2006 Available online 14 September 2006

Abstract Diagonal peptide chromatography consists of two consecutive, identical peptide separations with in between an enzymatic or chemical alteration of the side-chain structure of selected peptides. Such selected and altered peptides acquire different chromatographic properties thereby segregating from non-altered peptides in a series of secondary peptide separations. Originally described by Brown and Hartley in 1966, we have modified the technique such that it can be used for higher throughput gel-free proteomics. Our technique is termed COmbined FRActional DIagonal Chromatography (COFRADIC) and exploits evoked differences of the hydrophobicity of peptides in reverse-phase liquid chromatography. One important advantage of COFRADIC is its versatility: by changing the alteration reaction, different classes of peptides are sorted and finally analyzed. We previously published protocols and applications for separating methionyl, cysteinyl, amino terminal and phosphorylated peptides. In this review, we assess the potential of COFRADIC for the analysis of several posttranslational modifications emphasizing on in vivo protein processing events. Additional modifications that can be analyzed include phosphorylation and N-glycosylation. The potential of COFRADIC for isolating peptides holding such modified amino acids are discussed here. © 2006 Elsevier B.V. All rights reserved. Keywords: Diagonal chromatography; Protein processing; Protein phosphorylation; N-glycosylation; Proteomics

1. Introduction Two-dimensional polyacrylamide gels (2D-PAGE, [1,2]) have been critical for proteomics. 2D-PAGE underwent several technical improvements, most notably the introduction of immobilized pH gradients as the medium for isoelectric focusing [3] and, more recently, parallel labeling of proteomes with different fluorescent dyes (DIGE, [4]). Matured over more than three decades, 2D-PAGE is now routinely used for miscellaneous research topics. Nevertheless, and, obviously like any other analytical technique, 2D-PAGE holds several drawbacks making it impossible to completely visualize a proteome. Especially cumbersome are proteins only present in low copy numbers, hydrophobic proteins, very small and very large proteins and highly basic proteins.

⁎ Corresponding author. Tel.: +32 92649274; fax: +32 92649496. E-mail address: [email protected] (K. Gevaert). 1570-9639/$ - see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.bbapap.2006.09.003

Because of the limited loading capacity only proteins present in more than 1000 copies per cell (abundant proteins) may be analyzed in un-fractionated proteomes [5,6]. One solution to this “protein abundance problem” is focusing on the proteome of specialized sub-cellular structures (e.g. discrete organelles) leading to the identification of proteins present in low copy numbers present in such structures (reviewed in [7]). Even so, the rather limited protein concentration range that may be visualized by common protein stains – maximally about 4 orders of magnitude using fluorescent dyes – still leads to an under-representation of low copy number proteins. Hydrophobic integral membrane proteins remain challenging for 2DPAGE since they are difficult to fully extract from their hydrophobic environment, and once in solution tend to precipitate, especially near their isoelectric point during isoelectric focusing. The work of Thierry Rabilloud on analyzing membrane proteins is very well appreciated in the field, though it drove him to using lyrical manuscript titles, maybe illustrating the “protein hell” he went through when

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searching the ultimate remedy for hydrophobic proteins [8]? Its size also determines whether or not a protein is present on a 2Dgel with small proteins (typically less than 10 kDa) running off the gel and large proteins (more than 150 kDa) not properly entering the polyacrylamide gel(s). Which alternatives do we have for analyzing complex (both in terms of absolute numbers and in terms of concentration differences) proteomes? In fact, many different suggestions have been described, the one being more successful than the other … Abandoning the 2D-gel phase and focusing on the composite peptides of proteins – techniques generally known as non-gel, gel-free or peptide-centric proteomics – have proven to be well suited for the job. These techniques combine peptide liquid chromatography (LC), tandem mass spectrometry (MS/ MS) and database search robots, thereby enabling the identification of thousands of proteins in one sample: consider for example the recent identification of 7,792 proteins in a murine brain sample [9]. Non-gel approaches start by digesting the complex proteome typically using a specific protease such as trypsin, thus generating an even more complex peptide mixture. Two different approaches deal with these peptide mixtures, often encompassing tens of thousands of different peptides. The first approach tries to separate the proteome digest as exhaustive as possible by combining orthogonal (i.e. based on different physical peptide parameters such as charge and hydrophobicity) LC techniques. In the gel-free proteome world, these multi-dimensional LC separations were introduced by the group of John Yates [10] and became later on known as MudPIT (multidimensional protein identification technology, [11]). A second series of approaches first reduces the complexity of the peptide sample by (affinity) isolating a specific class of peptides prior to analysis, the general idea being that when mass spectrometers are not flooded by peptides, more peptides ions are finally fragmented and identified. The, already archetypical, example of these approaches is the ICAT (isotope-coded affinity tag) technique introduced by the group of Ruedi Aebersold [12]. Here, cysteinyl peptides are tagged with a biotin label enabling their (strept)avidin-based affinity isolation from complex mixtures. Cysteine is a rather scarce amino acid, but evenly distributed over proteomes whereby chances are substantially high that every protein in a given mixture is finally represented by at least one peptide [13,14]. Combinations of these two approaches – isolation of a subset of peptides and multidimensional LC – have been published and lead to increased proteome coverage [15]. Our lab has modified the previously described technique of diagonal electrophoresis and diagonal chromatography [16,17] for gel-free proteomics. Diagonal chromatography essentially consists of two identical peptide separations step with a chemical or enzymatic reaction (sorting reaction) in between. This reaction specifically alters the side-chain of a specific type of amino acids thereby changing the chromatographic properties of peptides holding the targeted (altered) amino acids. In our approach, peptides are separated by reverse-phase (RP) HPLC and the sorting reaction changes the column retention of a selected class of peptides by altering their overall hydrophobicity: it thus induces either

hydrophilic or hydrophobic shifts of the subset of altered peptides. Furthermore, we increased the throughput of diagonal chromatography by combining several altered primary fractions prior to the secondary separations. This explains the acronym COFRADIC as it stands for COmbined FRActional DIagonal Chromatography. By its nature and unlike other approaches, diagonal chromatography is not restricted towards the analysis of one given class of peptides such as for example cysteinyl peptides when using ICAT. Indeed, changing the COFRADIC sorting reaction selects for different classes of peptides. In our original publication [18] we used controlled oxidation of the sidechain of methionine to its sulfoxide counterpart. Peptides carrying the latter amino acid are more hydrophilic because of the induced dipole and, compared to their original retention during the primary COFRADIC run, elute in front of peptides devoid of methionine and are thereby specifically separated and isolated for further analysis. A somewhat different sorting strategy leads to the isolation of cysteinyl peptides. Here, proteins are first reduced and then reacted with Ellman's reagent, making a heterodisulfide bridge between the thiol group of cysteine and a hydrophobic nitrobenzoic acid group [19]. Following protein digestion and a primary peptide separation, the sorting step discerning cysteinyl peptides from all other peptides is a reduction by tris(2-carboxyethyl) phosphine which removes the nitrobenzoic acid group and renders cysteinyl peptides more hydrophilic [20]. Isolating methionyl or cysteinyl peptides overall reduces the complexity of the analyte mixture with a factor of about five. However, when analyzing complex proteomes (e.g. whole lysates of cells or tissues from higher eukaryotes) this complexity reduction may be insufficient still withholding several thousands of peptides for analysis. One obvious way to cope with such complex mixtures is placing an orthogonal, additional peptide separation step in front of the actual COFRADIC step. Therefore, we recently combined strong cation exchange (SCX) chromatography with the COFRADIC procedure for isolating methionyl peptides and applied it to a proteome preparation of human adult stem cells. In this way, we have identified 2,151 different proteins using MALDI-TOF/ TOF sequencing [21]. In our hands, this orthogonal peptide fractionation strategy leads to an improved proteome coverage, the trade-off being the concomitant increase in overall MS/MS analysis time as about six times more peptide fractions required analysis. A second approach for reducing the complexity of the final peptide mixture is isolating peptides that hold the extremity of a protein (e.g. its amino (N) terminus). For this reason, we adapted COFRADIC such that only peptides holding the Nterminus of a protein (N-terminal peptides) are isolated [22]. Here, the sorting reagent is 2,4,6-trinitrobenzenesulfonic acid (TNBS) which distinguishes N-terminal peptides that were either in vivo or in vitro blocked and internal peptides carrying a free α-amine (see below). Such N-terminal peptides lead to the highest possible reduction in sample complexity since every protein is finally represented by a single peptide: its N-terminal one. As the final analyte mixtures are less complex, under-

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sampling effects, leading to different protein maps when analyzing the same sample repeatedly, as observed in other gel-free approaches [23,24], do not affect the overall analysis outcome. Clearly, one of the major benefits of COFRADIC is its versatility: changing the sorting reaction leads to the isolation of a different set of peptides [14,25]. Below we focus on posttranslational protein modifications that can be analyzed by COFRADIC and emphasize on the global analysis of in vivo protein processing events since until now high-throughput proteomic techniques are not available for analyzing this important, irreversible protein modification. 2. Targeted analysis of in vivo protein processing sites Selective and restricted proteolytic protein cleavage, which we will further call protein processing, is vital for many processes in development, health [26,27] and disease [28–30]. Although protein processing is an important and wide-spread protein modification, until recently, the field of degradomics – a term coined by Chris Overall [31] – suffered from the lack of high-throughput and sensitive analytical techniques for studying protein processing on a proteome-wide scale. Indeed, when studying protein processing on the protein level, 2D polyacrylamide gel systems have typically been used since upon protein processing, the spot volume of the intact protein fades whereas novel spots at apparent lower molecular weights emerge, if the produced protein fragments are not fully degraded by the proteasome machinery. Gelbased studies have led to the discovery of several protease substrates (e.g. [32–34]) however they all suffer from the intrinsic 2D-PAGE drawbacks discussed above. Furthermore, when aimed not only at identifying protease substrates but also at mapping the actual sites of processing, gel-based studies are not at all apt. Indeed, in these systems defining protease processing sites is hardly an educated guess based on the migration of protein fragment(s) in 2D-gels, sequence coverage by MS-generated data and, if known, the specificity of the protease(s). Peptide-centric or gel-free proteomic approaches have been described that alleviate most of the disadvantages of 2D-gels for characterizing protein processing events. Recent examples include the combined use of ICAT and multidimensional LC for the characterization of substrates of the human MT1-MMP metalloproteinase [35] and SILAC labeling of Escherichia coli, identifying proteins trapped on the ClpXP bacterial protease complex following DNA damage [36]. However, these gel-free approaches do not grant immediate access to the actual site of processing since physical (solubility in aqueous solvents) and chemical (ease of ionization) parameters as well as the amount of peptides available for analysis finally determine if a peptide carrying the actual amino terminus of a protein (fragment) will be identified. Clearly, both gel-based and the gel-free proteome analyses mentioned above are not suited for clearly defining protein processing sites. An “in-between solution” allowing a more direct view on the (in vitro) substrate specificity of proteases uses peptides

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synthesized on miniaturized carriers (peptide arrays) and peptide or protein libraries presented on phages (reviewed in [37]). Examples of peptide arrays for determining protease substrate specificities include a small, 400-mer collection of peptides for studying the P1 and P′1 amino acids preferentially recognized by chymotrypsin and papain [38], a fluorescence-quenched peptide library for characterizing a recombinant cysteine protease of Leishmania mexicana [39] and a 722-mer solution-phase peptide library for screening various blood serine proteases [40]. Phage displays have been used for characterizing substrate features of human factor Xa and neurolysin [41]. Although both fluorogenic peptide arrays and phage display libraries allow parallel and rapid screening of a wide variety of potential protease substrates they hold a number of important drawbacks. Not only are these analyses done in an in vitro, rather artificial environment, the complexity that can be dealt with may well be below the complexity required for a comprehensive study of the structural confinements of protease substrates. For example, both granzyme A [42] and granzyme B [43,44] display an extended substrate specificity spanning more than six subsites on the substrate. This implies that both peptide libraries and phage display libraries should at least hold 64.106 different modules which, especially for the synthetic peptide libraries, currently remains a technical challenge to handle by many labs. Intuitively, since one processing event produces two novel protein forms, peptide-centric techniques targeted to the extremities of proteins (and their fragments) would be ideally suited for determining the exact processing site. Protein processing leads to an N-terminal and a C-terminal protein fragment, shown in red and blue respectively in Fig. 1. Compared to the intact precursor, the N-terminal protein fragment holds a novel C-terminus, whereas the C-terminal fragments holds a novel N-terminus (Fig. 1). Specific isolation and analysis of one of the peptides holding these “novel” protein extremities straightly points to the exact processing site in the precursor protein.

Fig. 1. Protein processing produces novel protein termini. In its most simple form, protein processing gives rise to an N- (shown in red) and C-terminal (shown in blue) protein fragment. Compared to the unprocessed protein form, these fragments produce two novel peptides: the N-terminal fragment carries a novel C-terminal end and the C-terminal fragment carries a novel N-terminal end. Possible techniques for isolating peptides carrying the N- or C-termini of proteins are indicated (numbers correspond to references in the main text).

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Although several approaches for the specific analysis of peptides holding the C-terminal part of a protein have been described [45–48], none was adapted for large-scale analysis. Likewise, next to the isolation of N-terminal by COFRADIC [22], various techniques have been described for isolating peptides holding the N-terminal part of proteins [49–51]. Whereas some of these approaches have been used for proteome-wide analysis [52–54], only the N-terminal COFRADIC technology has been used to map in vivo protein processing sites in a global, cellular context [55]. Central for the analysis of protein processing events was the combination of the COFRADIC procedure for isolating Nterminal peptides [22] with differential labeling of peptides from living and apoptotic Jurkat T-lymphocytes by trypsinmediated incorporation of two heavy oxygen-18 atoms in peptides ending on arginine [56]. Here, we schematize the isolation of N-termini (Fig. 2) and would like to refer the reader to our previous papers for further technical details [22,55,56]. Briefly, prior to trypsin digestion, all cystines are reduced and free thiol groups are alkylated (for instance using iodoacetamide). Next, all free α- and ε-amines are blocked by an acetylation step using an N-hydroxysuccinimide ester of trideuteroacetic acid [57]. Following trypsin digestion, which now only cleaves C-terminal to arginines, two types of peptides are generated: peptides carrying a free α-amine and peptides with a blocked α-amine. The latter amines are either in vivo acetylated, in vitro trideuteroacetylated or start with proline, pyrrolidone carboxylic acid (derived from an N-terminal glutamine or, to a lesser extent, from an N-terminal glutamic acid residue) or pyro- S-carbamoylmethylcysteine. This peptide mixture is separated a first time by RP-HPLC and typically resolves into twelve to fifteen so-called primary fractions. Then, each primary fraction is reacted with TNBS attaching a hydrophobic trinitrophenyl (TNP) group to free α-amines. When such an altered primary fraction is separated a second time using identical chromatographic conditions, TNP-peptides

Fig. 2. Scheme for the isolation of peptides holding the N-terminus of proteins.

are significantly longer retained on the RP column (i.e. they undergo a hydrophobic shift) and are thereby separated from the blocked peptides. These blocked peptides are thus collected in a number of secondary fractions which are finally analyzed by automated LC-MS/MS. To further distinguish between pre-existing (prior to processing events) N-terminal peptides (i.e. peptides holding the mature N-termini of unprocessed proteins) and novel N-terminal peptides (originating from processing events) an isotopic label on the isolated peptides is needed. In the original apoptosis study of Van Damme et al. [55] isotopic labeling was done postmetabolically by trypsin-driven incorporation of 18O isotopes into arginine-ending peptides. While this labeling procedure is applicable irrespective of the nature of the proteome sample (from a biopsy, a body fluid or cultured cells), the spacing introduced between the “light” and “heavy” peptide forms is only 4 atomic mass units and thus “on the edge” for a straightforward (and automated) statistical assessment of peptide concentration fluxes. This is especially unwieldy for large peptides with overlapping isotopes from their light and heavy variants. Furthermore, in mixtures of light and heavy peptides, stable incorporation of 18O isotopes can only be achieved when trypsin is completely inhibited since residual trypsin activity, even at a pH below 5, steadily exchanges the carboxyl oxygen atoms on C-terminal arginines [56]. Likewise, extremes of pH, as used for many RP-HPLC separations (e.g. [58]), result in gradual elimination of the 18O-label [56]. Therefore, and whenever possible, we suggest to metabolically label the proteins for peptide-centric proteomics. This type of labeling may be performed on various ways but, especially for animal cells, most conveniently and accurately the SILAC (stable isotope labeling by amino acids in cell culture) technology originally described by the group of Matthias Mann is used [59]. In our experience with different human cell lines, SILAC labeling is absolute after 5 to 6 cell population doublings and besides the increased segregation of the isotopic envelopes of light and heavy peptide variants (e.g. 6 Da when using 13C6-Arg or 13C515N-Met), differently treated cells may be blended prior to or following cell lysis, keeping further experimental variance to a minimum. A typical setup for characterizing in vivo protein processing is shown in Fig. 3. Here, stimulated (apoptotic) cells are labeled with natural isotopes whereas control (living) cells are grown in a medium in which natural arginine is replaced by 13 C6-arginine. Following cell lysis, the two proteomes are mixed and all further steps for the isolation of N-terminal peptides (Fig. 2) are performed on this mixed proteomic sample. An example of the final MS and MS/MS analyses is shown in Fig. 4. Using the setup depicted in Figs. 2 and 3, Fas-induced apoptosis was studied in human Jurkat T-cells. Following isolation of N-terminal peptides, LC-MS/MS analysis of the isolated peptides by a Q-TOF mass spectrometer generally reveals complex mass spectra as shown in the upper spectrum in Fig. 4: note that this (small part of a) spectrum constitutes a collection of 3 successive MS spectra merged over an interval of 36 s, while the complete elution range of one secondary fraction containing N-terminal

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Fig. 3. Typical setup for studying in vivo protein processing by N-terminal COFRADIC. One batch of cells is labeled with a naturally occurring essential amino acid (here, 12C6-arginine) and another batch of cells is labeled with a heavy variant of this essential amino acid (13C6-arginine). Following stimulation (here induction of apoptosis), the cells are lysed, mixed together and the mix of proteomes is processed for specific isolation of N-terminal peptides. In this setup, protein processing events are represented by trideuteroacetylated peptides carrying but 12C6-arginine.

peptides is generally in the range of 10 min. As expected, the majority of peptides appears as light and heavy N-terminal peptide ions having an average ratio of 1/1 and spaced by 6 amu. This is evident from the insets (red spectra) in the MS spectrum of the peptide couple at 663.41 Th and 666.43 Th and the couple at 699.42 Th and 672.43 Th (both peptide ions are doubly charged and thus spaced by 3 amu). When specifically analyzing processing events, such peptide couples are at first glance of minimal interest as they point to the Ntermini of the unprocessed form of the protein or its Nterminal fragment (see Fig. 1). However, after all observed ratios of such couples are determined, one may perform a statistical analysis pointing to N-terminal peptides enriched in a given sample (i.e. peptides with ratio values significantly differing from the measured distribution of values). In turn, such peptides can hint to proteins that were enriched in one cell lysate – an example are the histone proteins which are known to be more efficiently extracted using non-charged detergents from apoptotic cells [60] – or they hold protease recognition sites that were not completely processed in the apoptotic cells [55]. The peptide at 675.47 Th is only present in its light, 12C form and should thus point to a processing event (see Fig. 3). Indeed, following MS/MS analysis, Mascot [61] identified this peptide as AGM*AMAGQSPVLR (M* indicates a methionine-sulfoxide) holding a trideuteroacetylated N-terminus indicating that in vivo, this α-amino group was free. This peptide spans amino acids 173 to 185 in the polypyrimidine tract-binding protein 1 (PTB1, SwissProt accession number P26599) and is headed by the sequence AAVD. Previous studies indicated that PTB1 can be cleaved by caspases [62] and revealed that in vitro, caspase 3 cleaved PTB1 at the noncanonical sites at asparagines 7, 139 and 172 [63]. We here show that PTB1 is indeed processed in vivo in apoptotic Jurkat T-cells at Asp-172 and our previous analyses pointed to further processing at Asp-2, Asp-7 and, somewhat more surprisingly, Met-175 [55]. The fact that processing at Asp-2 was characterized points to one of the

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advantages N-terminal COFRADIC possesses over gel-based techniques aimed at identifying processing events. Asp-2 processing only removes two amino acids from the 57 kDa PTB1 protein and depending upon the separation potential of the chosen 2D-gel system such small sequence alterations may easily remain unnoticed. Clearly, processing at Met-175 is atypical for caspases and, as was observed for other proteins in dying Jurkat cells – e.g. processing of vimentin at the known caspase cleavage site Asp-84 was observed next to further processing at Phe-85 and Leu-87 [55] – this hints to the action of other proteases on the unprotected N-terminal end of the protein fragment formed by the initial caspase cut (e.g. [64]). We could not characterize the in vivo cleavage of PTB1 at Asp139 since the protein fragment created this way holds an N-

Fig. 4. MS and MS/MS based characterization of in vivo processing of the polypyrimidine tract-binding protein 1 at Asp-172. Using the setup shown in Fig. 3, the proteomes of living and Fas-induced apoptotic human Jurkat cells were compared based on their N-terminal peptides. The upper MS spectrum is only a small part of the m/z range normally covered by the Q-TOF mass spectrometer and clearly holds a high number of peptide ions. Zoomed regions show two peptide couples (red spectra) with light peptides at 663.41 Th and 669.42 Th respectively. Since these couples are spaced by 6 amu (note that the peptides are present as doubly charged precursors) and present in ratio (light/ heavy) values close to one, most probably they point to the mature N-termini of intact proteins. Next to a large number of such peptide couples, a peptide at 675.47 Th is present in one form (the light form) only. MS/MS analysis of this ion followed by Mascot database searching, identified the peptide as Ac (D3)-173AGM*AMAGQSPVLR185 of the polypyrimidine tract-binding protein 1 which is the result of caspase 3 mediated processing as was shown by previous in vitro studies (for the sake of clarity only the observed yn-type of fragment ions are indicated and the sequence tag that could be manually read from these fragments is shown in bold).

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terminal peptide that, if sorted, is only seven amino acids long and, in our experience, such small peptides are extremely burdensome to identify by Mascot because of the lack of a sufficient number of peptide backbone fragments. PTB1 binds to the polypyrimidine zone of introns and is involved in pre-mRNA splicing. Actually, the spliceosome appeared as a favored target for proteolytic degradation in Fasinduced dying Jurkat cells since 14 spliceosomal core or associated proteins were in vivo processed [55]. Clearly, both because of its nature (selecting for N-terminal peptides), the fact that SILAC may be tuned such that every sorted peptide carries an isotopic marker (12C6 or 13C6 arginine) and trideuteroacetate labeling of free α-amines (which are normally expected following a protein processing event), we strongly believe that N-terminal COFRADIC is currently the most promising proteomic technique for studying in vitro and in vivo processing events. 3. Other posttranslational modifications

Fig. 5. A hydrophobic shift is evoked by phosphatase treatment of phosphopeptides. Shown are the UV-absorption chromatograms (at 214 nm). The black line shows the chromatogram of a synthetic phosphorylated peptide baring the sequence ADLpYAHQGVEANK (phosphorylated on tyrosine). This peptide elutes at 41.23 min corresponding to a concentration of 21.8% (v/v) acetonitrile. Upon phosphatase treatment, the phosphate moiety is removed and the peptide undergoes a hydrophobic shift and when re-run on the same column and under identical conditions: it elutes at 47.60 min (or 26.3% (v/v) acetonitrile) (chromatogram shown by the grey line).

3.1. Protein phosphorylation Protein phosphorylation is the result of an enzymatic reaction of kinases acting on proteins, although recently, non-enzymatic phosphorylation by inositol pyrophosphates has been described [65]. Several non-gel technologies have been developed for mapping phosphorylation events on a global (cell or tissue) scale and the some of the most popular employ IMAC [66], SCX [67,68] or antibodies [69] enriching phosphorylated peptides or proteins prior to further analysis. We choose here not to discuss all possible techniques for phosphoproteomicis in detail and would like to refer to several other contributions in this special BBA issue. It is known that during reverse-phase separations, phosphorylated peptides display different chromatographic retention as compared to their non-phosphorylated counterparts [70]. Keeping this differential chromatographic behavior in mind, we adapted the COFRADIC sorting reaction in such a way that in vivo phosphorylation events can be studied [71]. Here, a blend of commercially available phosphatases (calf intestinal phosphatase, lambda protein phosphatase and the alkaline Escherichia coli phosphatase) holding a broad specificity towards phosphorylated amino acids was used to dephosphorylate peptides between the primary and secondary COFRADIC separation. After fine-tuning the RP-HPLC buffers systems (ammonium acetate (pH 5.5) in water/acetonitrile (see [71]), we noticed that, unlike previous reports [70], all phosphopeptides undergo a hydrophobic shift following dephosphorylation thus segregating from non-phosphorylated peptides. This sorting principle is illustrated in Fig. 5: following enzymatic dephosphorylation of a synthetic peptide phosphorylated on a tyrosine residue, a hydrophobic shift of more than 6 min on a conventional RP-HPLC system is observed. The extent of this shift depends on the nature of the phosphorylated amino acid (phosphorylated tyrosines showing larger shifts as compared to phosphorylated serines or threonines) and on the number of

phosphorylated residues (multiple phosphorylated peptides showing larger shifts than singly phosphorylated ones). One inherent disadvantage of analyzing phosphorylated peptides this way is that the actual information on the phosphorylated residue is lost during the sorting step. In fact, this knowledge is crucial not only to distinguish dephosphorylated peptides from artificially shifting peptides (e.g. due to methionine oxidation or deamidation, both resulting in hydrophilic shifts), but might also hint to the kinase involved since, at least for some kinases, recognized sequence motifs have been mapped (see for example [72]). To distinguish peptides shifting due to dephosphorylation from peptides shifting due to non-enzymatic reactions we applied an isotope-tagging procedure (see Fig. 6). In brief, a peptide mixture enriched for phosphopeptides by an IMAC step was split in two equal parts. One part (control peptides) was treated with a cocktail of phosphatases and then labeled with two oxygen-18 isotopes. The other part (phosphopeptides) was neither dephosphorylated nor tagged with isotopes. The primary COFRADIC separation was performed on the reassembled mixture of control and phosphopeptides. Upon dephosphorylation of peptides in the isolated primary fractions and a series of secondary COFRADIC separations, peptides dephosphorylated by this second phosphatase treatment undergo hydrophobic shifts. In fact, following the isotope tagging strategy, such peptides may not carry any oxygen-18 isotopes and are thereby discriminated from peptides shifting artificially since these will be present as couples of oxygen-16 and oxygen18 labeled peptides in a 1/1 ratio [71]. If such sorted, dephosphorylated peptides carry only one serine, threonine or tyrosine, one can assume that in vivo phosphorylation occurred on this amino acid. However, whenever more than one phosphorylation site is present, additional information is necessary to characterize the site(s) that was (or were) phosphorylated in vivo. One obvious way is to re-examine the primary fractions (thus prior to the sorting

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Fig. 6. Scheme depicting the COFRADIC isolation of phosphorylated peptides. Following trypsin digestion of a proteome, phosphorylated peptides are enriched by Fe3+-IMAC. After several washing steps, retained peptides (mainly phosphorylated peptides) are eluted and split into two equal parts. One part is untreated whereas the other part is dephosphorylated and labeled with two oxygen-18 isotopes. As such, peptides containing a phosphorylated amino acid are only present in their light form and following diagonal chromatography including a blend of phosphatases between two consecutive RP separations (see Fig. 5), they are easily distinguished from artificially shifting peptides that now appear as doublets separated by 4 Da.

reaction) using the data obtained by analyzing the secondary fractions (dephosphorylated peptides). In fact, m/z inclusion lists can be generated based on the sequence and number of potential phosphorylation sites residing in the identified dephosphorylated peptides. In turn, these lists are used to specifically fragment phosphorylated peptides present in the primary fractions. Applying this COFRADIC technology to a human cell line (HepG2 cells), we were able to identify 190 phosphorylated peptides from 152 different proteins. The majority of these peptides carried phosphorylation sites that had been detected in earlier studies, next to 38 novel sites, illustrating the potential of COFRADIC for analyzing in vivo phosphorylation events.

N-glycosylation is known (Asn-X-Ser/Thr), one may exploit this further to access glycosylated peptides following MS/MS analysis and database searching. When applied to 10 μL of affinity-depleted mouse serum, we characterized 127 different N-glycosylation sites in 82 serum proteins [74]. For the overall majority of these sites (117) earlier experiments and/or in silico predictions classified them as known targets for glycosylation illustrating the potential of COFRADIC for studying this protein modification. At this point, we would like to refer the reader to our original paper for full technical details [74] and to other chapters in this special issue dealing with other techniques for analyzing glycosylation events.

3.2. N-glycosylation

4. Conclusions and perspectives

As was speculated by the group of Fred Regnier [70], diagonal reverse-phase chromatography could be used to isolate N-glycosylated peptides. Recently, we developed a COFRADIC procedure for sorting out such peptides. After optimizing the chromatographic conditions (ammonium acetate at pH 5.5 was used as the ion pairing reagent in RP-HPLC) we noted that enzymatic deglycosylation by peptide N-glycosidase F (PNGase F), which removes glycan structures conjugated to asparagine [73], strongly alters the column retention of Nglycosylated peptides. Importantly, the evoked shits can be hydropholic or hydrophobic and this most probably reflects the charged nature of the sugar chain, with highly charged sugars (e.g. due to sialic acids and sulfate groups) resulting in a hydrophobic shift of the deglycosylated peptides. Furthermore, since deglycosylation is accompanied by conversion of the conjugated asparagine to an aspartic acid and a general motif for

One of the major advantages COFRADIC holds over other gel-free techniques is its versatility. As indicated above, changing the alteration reaction between the two consecutive RP-HPLC separations leads to the isolation of a different set of peptides. While our first applications focused on mapping the proteins present in cells or tissues [18,20,22], recently we exploited the possibility of analyzing protein modifications with our technology. Protein processing is an irreparable protein modification for which no straightforward techniques were available that allowed high-throughput and detailed characterization on a proteome-wide level (see above). Combining isolated Nterminal protein ends with isotope-tagging of peptides or proteins, we now dispose of a peptide-centric technique for mapping in vivo processing events ([55] and Figs. 2–4). We utterly believe that this technique may be of major interest for

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those working on the biology of proteases and protease inhibitors since we have not seen any analytical counterpart identifying a protease substrate and the actual in vivo cleavage site in a single analysis. The latter information appears very interesting as these data may be used to train algorithms for predicting sites recognized in vivo by a given protease (see for example [75]). Furthermore, we are currently introducing an orthogonal peptide separation step prior to N-terminal COFRADIC (analogous to coupling SCX to methionine-COFRADIC; [21]) thus increasing the number of N-termini (and thus potential protease cleavage events) identified in a proteome analysis. The other posttranslational modifications described above are all in vivo reversible and this was exploited for both protein phosphorylation and N-glycosylation. For the former a cocktail of phosphatases dephosphorylates phosphopeptides in between the two COFRADIC runs and results in a hydrophobic shift of the targeted peptides (Fig. 5). Peptides carrying N-glycans may be sorted following the action of PNGase F. Interestingly, here, such deglycosylated peptides may undergo both hydrophilic and hydrophobic shifts, depending upon the nature of the glycan structure and, more in particular, the number of charged sugar blocks [74]. It should be clear that eventually every type of modification that may be specifically converted or reverted, either enzymatically or chemically is, in principle, open to sorting by COFRADIC. Hence, we expect that the already substantial repertoire of peptide classes that may be studied by COFRADIC will continue to grow in the near future. Acknowledgements K.G. is a Postdoctoral Fellow of the Fund for Scientific Research-Flanders (Belgium) (F.W.O.-Vlaanderen). The project was supported by research grants from the Fund for Scientific Research-Flanders (Belgium) (project number G.0008.03), the GBOU-research initiative (project number 20204) of the Flanders Institute of Science and Technology (IWT), the Concerted Research Actions (GOA) from the Ghent University, the Inter University Attraction Poles (IUAP05) and the European Union Interaction Proteome (6th Framework Program).

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