Advances in the Analysis of Protein Phosphorylation - ACS Publications

Kalume , D. E.; Molina , H.; Pandey , A. Tackling the phosphoproteome: tools and ...... Monroe , David G. Camp , II , Richard D. Smith , H. Steven Wil...
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Advances in the Analysis of Protein Phosphorylation Alberto Paradela* and Juan Pablo Albar Departamento de Proteómica, Centro Nacional de Biotecnologia, Consejo Superior de Investigaciones Científicas, c/ Darwin 3, 28049 Madrid, Spain Received October 10, 2007

Phosphorylation is one of the most relevant and ubiquitous post-translational modifications. Despite its relevance, the analysis of protein phosphorylation has been revealed as one of the most challenging tasks due to its highly dynamic nature and low stoichiometry. However, the development and introduction of new analytical methods are modifying rapidly and substantially this field. Especially important has been the introduction of more sensitive and specific methods for phosphoprotein and phosphopeptide purification as well as the use of more sensitive and accurate MS-based analytical methods. The integration of both approaches has enabled large-scale phosphoproteome studies to be performed, an unimaginable task few years ago. Additionally, methods originally developed for differential proteomics have been adapted making the study of the highly dynamic nature of protein phosphorylation feasible. This review aims at offering an overview on the most frequently used methods in phosphoprotein and phosphopeptide enrichment as well as on the most recent MS-based analysis strategies. Current strategies for quantitative phosphoproteomics and the study of the dynamics of protein phosphorylation are highlighted. Keywords: IMAC • Mass spectrometry • Phosphopeptides • Quantitative phosphoproteomics • Protein phosphorylation

Introduction Since the isolation in 1932 by Phoebus A. Levene and Fritz A. Lipmann of phosphoserine (first described as serine phosphoric acid), protein phosphorylation has turned to be one of the most biologically relevant and ubiquitous post-translational protein modifications. Phosphorylation is a reversible modification affecting both the folding and function of proteins, regulating essential functions such as cell division, signal transduction, enzymatic activity, and so forth. According to comprehensive databases, the estimated number of phosphorylation sites in the mammalian proteome could be as high as 105 (www.phosphosite.org). Other values calculate at 30–50% the percentage of proteins supposed to be phosphorylated at some point.1 The relevance of phosphorylation is underlined by the fact that the number of genes involved in phosphorylation processes may constitute as much as 2–3% of the entire eukaryotic genome.2 For example, about 2% of the human and mouse genomes encode protein kinases (PK) with 518 and 540 distinct PKs determined in human3 and mouse,4 respectively. The analysis of the genome of Saccharomyces cerevisiae has revealed the presence of 123 putative protein kinases and 40 protein phosphatases, respectively, constituting approximately 2% of expressed yeast proteins.5,6 Finally, 251 protein kinases and 86 protein phosphatases have been identified in the Drosophila melanogaster genome.7 Both counteracting enzymatic systems, kinases and phosphatases, regulate precisely protein phosphorylation and dephosphorylation, and differ in * To whom correspondence should be addressed. Phone, 34-915854696; fax, 34-915854506; e-mail, [email protected]. 10.1021/pr7006544 CCC: $40.75

 2008 American Chemical Society

their kinetic properties, substrate specificities, and cellular or tissue distribution. Among the amino acids that can be phosphorylated, O-phosphates are by far the most abundant, mostly attached to serine, threonine, and tyrosine residues. The occurrence of phosphorylation on Ser and Thr residues is more frequent than on Tyr residues, with the ratio of pSer/pThr/ pTyr in the order of 1800:200:1.8 The phosphoramidates of arginine, histidine, and lysine also occur as do acyl derivatives of aspartic and glutamic acid, although they are less abundant. For the aforementioned reasons, the analysis of protein phosphorylation is of paramount importance. A comprehensive study of protein phosphorylation should include the identification of phosphoproteins and sites of phosphorylation (phosphoproteomics), the identification of the proteins (kinases and phosphatases) involved in the phosphorylation process, and a description of the biological events following the phosphorylation events. Mass spectrometry (MS) has become a powerful technology for proteomics and a method of choice for unbiased analysis of protein phosphorylation.9 However, phosphoproteomics faces the challenge of low-abundance proteins and the often low ratio of phosphorylated versus nonphosphorylated proteins found in vivo. Additionally, while some residues are constitutively phosphorylated, others are only transiently phosphorylated, in some cases at very low levels. It is also important to note that often MS-analysis is unable to identify unambiguously the phosphorylation site(s) within a peptide. The results obtained in collaborative studies focused on the ability of proteomic laboratories to identify the phosphorylation sites present in a relatively simple mixture of phosphoproteins have clearly demonstrated that phosphorylation site analysis still Journal of Proteome Research 2008, 7, 1809–1818 1809 Published on Web 03/08/2008

reviews represents a challenge for many laboratories. The wide range of results suggested that analytical methods are far from being well-established and that a significative percentage of the data about protein phosphorylation published to date should be reconsidered carefully (ABRF sPRG Study 2007; www.abrf. org).10 Despite these major drawbacks, recent advances in phosphoproteomics technologies, including sample enrichment at the phosphopeptide and phosphoprotein levels, MS analysis, phosphorylation site mapping, and quantitative phosphoproteomics, have made feasible large-scale phosphoproteomics analysis in a wide set of biological models: bacteria,11 plants,12,13 yeast,14,15 and metazoa.16–18 This review aims to offer a detailed overview on MS-based phosphoproteomics analysis, emphasizing recent advances such as phosphopeptide enrichment using TiO2-based stationary phases, ECD- and ETD-based phosphopeptide MS-analysis, and quantitative phosphoproteomics. Classical approaches are clearly depicted and reviewed elsewhere in more detail.1,8,19

Selective Enrichment of Phosphoproteins and Phosphopeptides Site-specific analysis of post-translational modifications is usually performed by MS approaches, requiring that the modified protein (phosphoprotein) first be cleaved enzymatically or chemically (less frequently) into peptides of a size suitable for sequence analysis. However, substoichiometric phosphorylation, wich reduces phosphoanalyte abundances compared to corresponding unphosphorylated forms, phosphopeptide inefficient ionization, and specific losses occurring by adsorption to metal or plastics, make highly advisable the use of phosphoprotein and/or phosphopeptide-specific enrichment methods. On most occasions, enrichment of the sample in phosphoproteins followed of protease-specific digestion and MS-analysis is not sufficient to identify the sites of phosphorylation present in a complex sample and requires a second enrichment step at the phosphopeptide level. Commercial kits make the enrichment process easy, fast, and reproducible, although it has been clearly demonstrated that the different methods available differ in the specificity of isolation and in the set of phosphoproteins and phosphopeptides isolated,20 strongly suggesting that no single method is sufficient for a comprehensive phosphoproteome analysis (Figure 1). Immunoprecipitation. Antibodies specific to phosphorylated residues are used to immunoprecipitate total proteins rather than phosphopeptides. Immunoprecipitation of phosphotyrosine-containing proteins is more frequent than immunoprecipitation using phosphoserine- or phosphothreoninespecific antibodies as the former are most reliable than antibodies specific to Ser/Thr-phosphorylated proteins. This fact explains why phosphorylation in Tyr residues has been studied so intensively in the last years despite its low abundance compared to phosphorylation in Ser and Thr residues.21–24 Specific phosphotyrosine binding domains (PTB) have provided a useful tool to profile the global tyrosine phosphorylation state of the cell.25,26 The reasons that explain why most phosphoserine- and phosphothreonine-specific antibodies are hardly suitable for phosphoprotein immunoprecipitation are not known, but it is well-known that frequently these antibodies are very specific to certain consensus motifs. In practice, immunoprecipitation of phosphoproteins phosphorylated in these residues usually requires an expensive mixture of different 1810

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Paradela and Albar antibodies. Therefore, their use has only been reported in a limited number of studies.27 Immobilized Metal-Ion Affinity Chromatography (IMAC). IMAC is the most frequently used technique for phosphopeptide and phosphoprotein enrichment, although it was originally introduced for purification of His-tagged proteins.28 This technique allows a higher success rate in phosphopeptide analysis because it reduces ion suppression effects that would otherwise occur with untreated complex mixtures.29 Phosphorylated peptides or proteins are bound to the IMAC stationary phase by electrostatic interactions of its negatively charged phosphate group with positively charged metal ions bound to the column material via nitriloacetic acid (NTA), iminodiacetic acid (IDA), and Tris(carboxymethyl)ethylenediamine (TED) linkers. Immobilized metal ions such as Ni2+, Co2+, or Mn2+ were initially shown to bind strongly to proteins with a high density of histidines. However, immobilized metal ions of Fe3+, Ga3+, and Al3+ have been demonstrated to show better binding characteristics with phosphopeptides. Recently, immobilized Zr4+ has been reported to bind phosphopeptides with high specificity.30 One of the major drawbacks of IMAC-based strategies is the nonspecific binding of peptides containing acidic amino acids, that is, Glu and Asp, and the strong binding of multiply phosphorylated peptides. Despite following an apparently simple common schema (binding-washing-eluting), IMAC experimental conditions are very variable and care should be taken as small variations in the experimental conditions (for example, pH, ionic strength, or organic composition of the solvents) could drastically affect the selectivity of the IMAC stationary phase. Nonspecific binding of acidic peptides can be diminished by esterification of carboxylic acids to methyl esters using HCl-saturated, dried methanol.15 Reaction conditions have to be chosen carefully to avoid both incomplete esterification and side reactions because they increase sample complexity. IMAC procedures have become very popular rapidly due to its good compatibility with subsequent separation and detection techniques such as LC-ESIMS/MS and MALDI MS.14,17,31 On the basis of measurements of 32P or 33P-radioactivity in whole cell extracts and in phosphoprotein samples after enrichment, IMAC-based techniques have been reported to recover up to 70–90% of total phosphoproteins.32 Recently, Machida et al. optimized the conditions for IMAC to enrich for phosphoproteins.33 According to the authors, Ga3+ was the best option among the metal ions tested. About 1/10 of the total protein was recovered in the eluate when whole cell lysates were analyzed. Most interestingly, specific phosphoproteins could be tracked along the enrichment procedure, demonstrating the efficiency of this method. In addition to IMAC, strong cation exchange chromatography (SCX) has been used in the enrichment of phosphorylated peptides. This procedure is based on the fact that under acidic conditions (pH 2.7) tryptic phosphorylated peptides are single positively charged and amenable to further separation from nonphosphorylated tryptic peptides that usually have a net charge of 2+ at low pH. Although this strategy does not have high specificity and the fractions enriched in phosphopeptides also contain a high percentage of contaminants, SCX enrichment has been used for massive phosphoprotein profiling in developing mouse brain18 and in nuclear extracts of HeLa cell lysates.16 One of the main advantages of this method is that complex tryptic mixtures can be analyzed directly34 since the

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Figure 1. Summary of the experimental strategies commonly used in the analysis of phosphoproteins and phosphopeptides.

widely adopted multidimensional LC strategy in shotgun proteomics uses a similar SCX-RP approach. Titanium Dioxide. A promising alternative to the use of IMAC for the enrichment of phosphorylated peptides was first described by Pinkse et al.35 The approach is based on the selective interaction of water-soluble phosphates with porous titanium dioxide microspheres via bidentate binding at the TiO2 surface. Phosphopeptides are trapped in a TiO2 precolumn under acidic conditions and desorbed under alkaline conditions. An increased specificity for phosphopeptides has been reported, although TiO2-based columns still retain nonphosphorylated acidic peptides. Peptide loading in 2,5-dihydroxybenzoic acid (DHB) has been described to efficiently reduce the binding of nonphosphorylated peptides to TiO2 while retaining high binding affinity for phosphorylated peptides.36 This improved TiO2 procedure was found to be more selective than IMAC. An exhaustive analysis has been recently reported that describes the relationship between the occurrence of some amino acids and the phospho-specific and nonspecific binding of peptides using TiO2-based enrichment.37 Two well-charac-

terized peptide mixtures consisting of either 33 or 8 synthetic phosphopeptides or their nonphosphorylated counterparts and differing in charge and hydrophobicity were tested. The results confirmed the high selectivity of titanium dioxide for phosphorylated sequences. Interestingly, drastically reduced recovery was observed for phosphopeptides with multiple basic amino acids. The importance of phosphopeptide enrichment was highlighted by the fact that 50–75% of the phosphopeptides from both mixtures were not detected by MALDI MS without previous enrichment. One of the main advantages of this approach is that it can be easily coupled with a LC-ESI-MS/ MS or LC-MALDI MS/MS workflow. Recently, the use of zirconium dioxide micro tips for phosphopeptide isolation has been described. These micro tips displayed similar overall performance as TiO2 columns, although more selective isolation of singly phosphorylated peptides was observed with ZrO2, whereas TiO2 preferentially enriched multiply phosphorylated peptides.38 This approach has been used to selectively isolate phosphopeptides from the Journal of Proteome Research • Vol. 7, No. 5, 2008 1811

reviews tryptic digestion of a mouse liver lysate. Overall, 248 phosphorylation sites and 140 phosphorylated peptides were identified.39 Chemical Modification of Phosphate Groups. Several methods for enrichment of phosphoproteins and phosphopeptides are based on the specific chemical modification of phosphate groups. Oda et al. described a method for enriching phosphoserine/threonine-containing proteins and for subsequent identification of the phosphoproteins and sites of phosphorylation. The method involved chemical replacement of the phosphate moieties by an affinity tag and required free sulfhydryls to be blocked before the tagging step.40 One of the main disadvantages of this method is that O-linked sugar moieties may also undergo β-elimination along the process. Thus, cross-reactivities with glycosylations should be tested carefully. A second disadvantage is that β-elimination is not applicable to tyrosine phosphorylation. A different approach reported by Zhou et al. bound phosphopeptides to sulfhydryl-containing compounds via phosphoamidate-bonds.41 Modified phosphopeptides are further linked to a solid support with immobilized iodoacetylgroups and obtained as native phosphopeptides after acid elution. This method is applicable to phosphotyrosine-containing peptides as well as those containing phosphoserine and phosphothreonine. However, the amino and carboxyl groups of the peptides have to be protected to avoid undesired reactions. Recent modifications of this method used an aminoderivatized dendrimer42 or controlled pore glass derivatized with maleimide for phosphopeptide isolation.43 Interestingly, in the methods based on phosphoramidate chemistry (PAC), the phosphate group remains bound to the phosphopeptide, facilitating the identification of the site of phosphorylation. One of the major drawbacks of these methods is that reaction conditions have to be monitored very carefully to avoid side reactions and undesired modifications. Unwanted and partial reactions increase sample complexity while diminishing the total yield of the purification. As a result, these methods require large amounts of starting sample with the result that only abundant proteins are easily identified. A list of the major improvements in phosphopeptide derivatization methods can be found elsewhere.19 As a summary, it is important to stress that large-scale phosphoproteomics studies should use several phosphopeptide isolation methods. Although most of the methods described to date are reproducible, it is also clear that they preferentially isolate different but somewhat overlapping subsets of the phosphoproteome.20 In other words, no single method is suitable for the isolation of the entire phosphoproteome as the mixture of phosphopeptides purified differ in terms of size, charge, and number of phosphorylation sites. For example, PAC-based methods typically tend to isolate peptides with one phosphorylation site, while IMAC methods preferentially isolate multiple phosphorylated peptides. Finally, TiO2 stationary phases are biased toward acidic peptides with one phosphorylation site, although multiphosphorylated peptides have also been eluted from TiO2 columns.44 The differential selectivity could also explain the variability of the results obtained in some comparative studies (ABRF sPRG study; www.abrf.org).

Identification of Phosphorylation Sites Phosphoproteins are usually phosphorylated on a number of different sites throughout the protein with individual sites being phosphorylated to various degrees. Determination of phosphorylation sites is a challenging task, but not impossible. As only state-of-the-art mass spectrometers (e.g., FT ICR-MS 1812

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Paradela and Albar or LTQ-Orbitrap) reach the level of resolution required for the analysis of entire proteins,45 “bottom-up” approaches dealing with peptides obtained from protein digests are preferred over “top down” techniques dealing with whole proteins. The proteome to be analyzed is cleaved into peptides of size suitable for MS analysis, ideally between 700 and 3000 Da, yielding peptide mixtures containing both phosphorylated and nonphosphorylated peptides. Enzymatic digestion of phosphoproteins generate peptides (phosphopeptides) that could contain more than one potential phosphorylation site, making a complete structural analysis of the phosphopeptides mandatory. Analysis is performed by tandem mass spectrometry (MS/ MS) to unambiguously establish the sequence and to determine which residues are phosphorylated.46 There are a number of different MS methods for determining what residues are phosphorylated in a peptide, although the preferred ones are those that make use of the specific fragmentation behavior of phosphopeptides such as triple-quads or Q-TOF instruments. One of the major drawbacks of most of the MS-based methods is that phosphoester bonds are very labile in the experimental conditions (CID, collision-induced dissociation) habitually used, resulting in a loss of phosphoric acid from the peptide and complicating the localization of the site at which the modification was originally attached. As mentioned previously, phosphopeptides in complex mixtures show decreased ionization rates due to suppression effects. Measuring phosphopeptides in negative ion mode can reduce this effect.46,47 However, negative-polarity MS/MS spectra are frequently difficult to decipher, reducing drastically the rate of peptide identification. Negative-mode MS/MS of phosphorylated serine, threonine, and tyrosine residues yield fragments of -79 Da (PO3-) and -63 Da (PO2-). Selective monitoring of phosphopeptide parent ions in negative mode based on their -79 Da ion signature,followed by polarity switching to obtain positive-ion MS/MS spectra, is a commonly used method. Howewer, it should be considered that polarity switching between positive and negative ion modes results unavoidably in decreased scanning rates. The best way to reduce the negative ion suppression effects is a reduction of sample complexity. This is achieved first by enriching the sample in phosphopeptides, followed by fractionation of the enriched sample using powerful separation techniques such as nanoHPLC or capillary electrophoresis (CE). These separation techniques can be easily coupled online with a wide array of mass spectrometers.48 Capillary electrophoresis coupled with mass spectrometry (CE-MS) has proven to meet the requirements of high-throughput, exceptional resolution, and outstanding certainty in protein/peptide identification.49 However, the method requires further improvement to become a widely used technique as it is limited by the small sample volumes that can be applied. Therefore, separation of the phosphopeptide-enriched sample is done preferentially by LC-MS and especially nano-LC-MS as these approaches show high sensitivity (in the low fentomolar-high attomolar range) and good reproducibility. Precursor ion scanning is particularly useful for analysis of phosphorylations that are stable during MS and generate specific fragment ions that can be monitored, such as tyrosine phosphorylations. Triple quadrupole mass spectrometers equipped with ion-counting detectors provide the highest sensitivity of any mass analyzer for precursor-ion scanning due to their capability to signal-time average very weak data and reject noise. The first quadrupole (Q1) scans through the entire

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Advances in the Analysis of Protein Phosphorylation mass range, and the peptides passing through are fragmented in the second quadrupole (Q2). The third quadrupole (Q3) is used for monitoring a specific fragment ion characteristic for the residue of interest (e.g., m/z 79 for phosphopeptides in negative ion mode or m/z 216.04 for the immonium ion of phosphotyrosine). In neutral loss scanning, Q1 and Q3 are simultaneously scanned over two different m/z ranges, which difference corresponds to the m/z value for the neutral molecule that is being lost (for a [M + 2H]2 phosphopeptide, this value is 49 m/z). This approach is not applied to phosphorylations that are stable during MS such as tyrosine phosphorylations. 3DIon traps do not perform “real” precursor ion scanning or neutral loss analysis. However, the prominent fragment ion generated after CID-induced phosphate loss in MS/MS mode (MS2 mode) is used for further fragmentation in MS/MS/MS mode (MS3 mode). MS3 spectrum yields more sequence information than the MS2 spectrum, although this information is not useful for identifying unambiguously where the phosphorylation site is located. Despite these drawbacks, this strategy was used to identify more than 2000 phosphorylation sites from HeLa cells in a large-scale phosphoproteomics analysis using a 3D ion trap with software-controlled, neutral, loss-dependent MS3 capabilities.16 Two recent fragmentation techniques have been recently developed to avoid the prominent loss of the phosphate group seen during CID. Both fragmentation techniques, electron capture dissociation (ECD)50 and electron transfer dissociation (ETD),51 have been implemented on FT-ICR, LIT, and 3D IT instruments. ECD and ETD are applicable to large peptides and small proteins and are particularly useful in the analysis of multiply charged peptides and in the identification of posttranslational modifications. The reason is that post-translational modifications easily lost in the case of CID analysis remain intact when ECD or ETD fragmentation is used, making the assignment of phosphorylation sites more precise52 (Figure 2). The principle of ECD is to react multiply protonated polypeptide cations with low-energy electrons. This induces fragmentation at the amide (N-CR) bond to produce c-type and z-type fragment ions. In the case of ETD, the electron transfer from the radical anions (e.g., singly charged anthracene anions) to multiply protonated peptides induces fragmentation of the peptide backbone along pathways analogous to those described for ECD. Recently, Molina et al. have evaluated the use of ETD for a global phosphoproteome analysis. A total of 1435 phosphorylation sites were identified in TiO2-enriched samples obtained from human embryonic kidney cells 293T. A comparison of ETD and CID modes demonstrated that CID yielded 60% less phosphorylation site identifications with an average of 40% more fragment ions per fragmentation spectrum.53 However, although the number of phosphorylation sites identified was lower, a significant set of phosphopeptides identified when CID was used were specific for this method, indicating that both CID and ETD should be combined for a more comprehensive analysis. Similarly, ETD for peptide fragmentation allowed the identification of 1252 phosphorylation sites from an IMAC-enriched sample obtained from 30 µg of total yeast protein.54 Several examples have been published demonstrating the feasibility of ECD in the identification of phosphorylation sites,45,55,56 although large-scale studies have not yet been published.

Quantitative Phosphoproteomics Quantitative proteomics is particularly useful to elucidate highly dynamic processes such as phosphorylation. 2D-PAGE differential phosphoproteomics, using either DIGE- or silver stain-based gels, has provided a significative number of results. Although specific patterns seen in 2D-PAGE gels alert about the presence of putative phosphorylated forms of a given protein, the use of phosphorylation-specific stains such as Pro-Q DPS57 is strongly recommended due to two main reasons: (a) Pro-Q DPS binds directly to the phosphate moiety of phosphoproteins with high sensitivity and linearity, and (b) the stain is fully compatible with other staining methods (e.g., DIGE) and modern MS-based analysis. The use of phosphorylation-specific fluorescent stains such as Pro-Q DPS in 2D-PAGE gels has been reported for various biological models.11,12,58–62For example, this approach has been used in the analysis of the regulation of endosomespecific phosphoproteins obtained after subcellular fractionation from EpH4 mammary epithelial cells and following stimulation by epidermal growth factor (EGF).63 Similarly, it has been reported that stimulation of human polymorphonuclear neutrophils (PMS) through the formyl peptide receptor like-1 (FPRL-1) alters the protein pattern of PMNs and that several phosphoproteins such as L-plastin, moesin, cofilin, and stathmin are up-regulated or down-regulated under these circumstances.64 One of the major drawbacks of 2D-PAGEbased quantitative phosphoproteomics is that only a relatively limited number of proteins is amenable to detection and identification. Moreover, large-scale quantitative profiling of phosphoproteins requires a significative number of highresolution 2D-PAGE maps to achieve statistically significant data. This approach has been used to map the changes occurring in the phosphoproteome during seed filling in oilseed rape. Up to 103 phosphoproteins (70 nonredundant) were identified and quantified.65 More details about 2D-PAGE-based quantitative phosphoproteomics as well as other classical approaches are clearly depicted and reviewed elsewhere.66 In addition to 2D-PAGE-based approaches, large-scale quantitative phosphoproteomics studies are performed by (2D) LC-MS/MS analysis of phosphopeptide-enriched samples. The samples to be analyzed are tagged previously by stable-isotope labeling to make them distinguishable from each other. It is very important to note that labeling should not affect the chemical properties of identical (phospho) peptides obtained from the samples to be compared (for example, sample A and B) (Figure 3). In other words, “light” and “heavy” phosphopeptides must have identical LC elution times ensuring that the ionic environment is equal, and thus, the ratio of the relative intensities can be used to estimate the relative phosphopeptide abundance in each sample. There are various ways to incorporate stable isotopes into (phospho) peptides. Metabolic labeling of proteins using amino acids labeled with 15N versus 14N, or 13C versus 12C, is an effective approach to stable isotope labeling. This method (called SILAC, stable isotopederivatized amino acids in cell culture) was originally developed as a simple and accurate procedure that can be used as a quantitative proteomic approach in any cell culture system.67 In the most effective implementation of the method, the addition of 13C6-arginine and/or 13C6-lysine to the cell growth medium ensures that all tryptic cleavage products carry at least one C-terminal labeled amino acid. Protein labeling in excess of 90% is tipically obtained after a small number (6–8) of passages. A small number of studies, for example in plants,68 have demonstrated the feasibility of in vivo total 15N metabolic Journal of Proteome Research • Vol. 7, No. 5, 2008 1813

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Figure 2. Direct comparison of MS/MS spectra obtained when the phosphopeptide SNQRSpSHSpSTLDDIL (M + 3H 607.25) was subjected to alternating ETD (A) and CID (B). Experiments were performed in a HCTultra ion trap equipped with an ETD device (Bruker Daltonics, Bremen, Germany). For simplicity purposes, only the most significative single-charged fragments are indicated in each case. Note the low number of fragments and poor sequence coverage obtained when CID was used.

protein labeling of higher organisms. However, the cost and time required recommends avoiding this approach. In practical terms, in vivo metabolic labeling is restricted to situations where cells may be grown on labeled media. Despite this limitation, the method has been adapted for quantitative phosphoproteomics studies.69 Gruhler et al. combined SILAC, SCX, and IMAC for phosphopeptide enrichment and LC-MS in a quantitative analysis of the yeast pheromone response. SILAC was achieved by incorporation of 13C6-arginine and 13C61814

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lysine. More than 700 phosphopeptides were identified and 139 were found to be differentially regulated at least 2-fold in response to pheromone.70 A similar approach has been described recently to perform a quantitative analysis of nine different phosphorylation sites of the epidermal growth factor.71 As SILAC-based approaches are only applicable to growing cells, several methods have been designed to label peptides or proteins in vitro. These include isotopically distinguishable commercial reagents specific for different functional groups

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Figure 3. Summary of a stable isotope labeling-based method adapted for quantitative phosphoproteomics. In vitro labelling methods have been optimized for comparison of three (ICPL, Bruker Daltonics) or more samples (iTRAQ, Applied Biosystems). Samples are labelled and combined prior to digestion and purification steps, although some methods (iTRAQ) require proteolysis previously to sample labelling. Phosphopeptide enrichment methods are highly recommended prior to LC-ESI-MS/MS analysis. The relative signal intensities of “light” and “heavy” peptides are used to quantify the relative amounts of a given protein in samples A and B.

such as amines (ICPL, iTRAQ) or sulfhydryls (ICAT). Isotopecoded affinity tag (ICAT) was originally developed as a reagent containing either zero or eight deuterium atoms that specifi-

cally reacts with free cysteine residues.72 Subsequent ICAT versions included changes, for example, incorporation of 13C/ 12 C isotopes, to improve the chromatographic properties of the Journal of Proteome Research • Vol. 7, No. 5, 2008 1815

reviews reagent. As cysteine is a rare amino acid (occurrence in proteins of ∼1.8%), ICAT is not suitable for quantifying proteins with none (∼8% in humans) or few cysteines. In practical terms, its use should be avoided in quantitative phosphoproteomics, as the number of (tryptic) peptides containing both a phosphorylation site and a cysteine residue is extremely low. This key issue has been solved by the development of reagents that specifically label the peptide at the N-terminus and the -amino group of lysine residues, such as iTRAQ73 and ICPL.74 ICPL and iTRAQ allow multiplexed quantitation of 3 and 8 different samples, respectively, and are particularly useful in the analysis of biological systems over multiple time points. Although they have been extensively used in differential proteomics, only iTRAQ has been recently used to obtain quantitative phosphoproteomics data.75 Kim et al. reported a method based on IMAC purification of iTRAQ-labeled phosphopeptides which allowed depicting precisely the phosphorylation events following CD3 and/or CD28 stimulation in Jurkat cells. In all, 101 tyrosine and 3 threonine phosphorylation sites were identified, while 87 sites were quantified across four different cell states.76 Other amino-reactive isotope tags have been also developed for quantitative phosphoproteomics analysis.77,78 Although promising, the number of reports published to date using these reagents are not enough to figure out their feasibility in quantitative phosphoproteomics accross different biological models. A completely different approach for stable isotope labeling is the incorporation of 16O/18O during the hydrolysis reaction catalyzed by trypsin or Glu-C. Incorporation of 18O into C-termini of peptides results in a mass shift of 2 Da per 18O atom. As the hydrolisis catalyzed by trypsin and Glu-C introduces two oxygen atoms, it results in a 4 Da mass shift wich is generally sufficient to differentiate labeled from unlabeled samples. This strategy combined with IMAC-based purification of phosphopeptides allowed the quantification of the changes in the phosphoproteome of the cell body and pseudopodium of chemotactic cells. A total of 228 unique phosphopeptides corresponding to 197 proteins were identified.79 Finally, label-free methods for quantitative analysis have also been described as an alternative to expensive isotope-based labeling techniques. As with isotopic labeling, these methods are based on the measurement and comparison of the mass spectrometric signal intensities of peptide precursor ions. However, as precursor ions are not isotopically distinguisable, samples to be compared have to be analyzed in independent runs. Afterward, the ion chromatogram for every peptide is extracted from an LC-MS/MS run and its mass spectrometric peak area integrated over the chromatographic time scale. In practical terms, label-free comparison of integrated peak intensities obtained in different LC-MS/MS runs requires a combination of accurate mass and stringent reproducibility in sample processing and chromatography. Special software has been developed to align LC-MS/MS runs prior to identifying corresponding peptides between different experiments.80,81 To date, one report successfully applied label-free quantification of IMAC-purified of phosphopeptides. A total of 714 phosphorylation sites on 223 phosphoproteins were identified. A number of proteins that significantly changed phosphorylation state in response to vasopressin treatment was identified.82 The final goal of quantitative phosphoproteomic studies is to elucidate the dynamics of protein phosphorylation across different cellular physiological and pathological states. In other words, the objective is to obtain a picture as complete as 1816

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Paradela and Albar possible of the evolution of the phosphoproteome as a function of time, stimulus, and other parameters. For this purpose, it is essential to develop and integrate global and quantitative methods. Blagoev et al. described the dynamic profile of 81 affinity-purified tyrosine-phosphorylated proteins after stimulation by epidermal growth factor (EGF) of three different cell populations.83 More recently, the temporal dynamics of 6600 phosphorylation sites on 2244 proteins after stimulating HeLa cells with epidermal growth factor has been published.84 Interestingly, results showed that phosphorylation is regulated differently on different sites within the same protein suggesting that the dynamics of phosphorylation should be measured site specifically rather than for the protein as a whole. Considering the complexity of the phosphoproteome dynamics, integrative approaches like this are essential for a systematic understanding of cellular behavior.

Summary The analysis of the phosphoproteome is one of the most exciting and challenging tasks in current proteomics research. Large-scale phosphoproteome analysis is now an affordable task since the development of methods for the purification of phosphoproteins and phosphopeptides as well as the new mass spectrometry technologies developed for the structural analysis of the enriched samples. However, many challenges lie ahead. Purification methods are far from rendering homogeneous results as it has been demonstrated that they tend to isolate preferentially specific subsets of phosphopeptides. Comparative studies have concluded that different proteomic strategies are complementary to each other. Therefore, the analysis of the phosphoproteome requires a combination of multiple techniques. In the last years, the interest has turned on the study of the phosphoproteome as a highly dynamic environment. Classical proteomic strategies such as 2D-PAGE combined with the use of phosphoprotein-specific stains such as Pro-Q DPS are now complemented with the combined use of stable isotope labeling and phosphopeptide enrichment followed by LC-MS/MS(MS) analysis. These strategies will permit the analysis of the evolution of the phosphoproteome across different cellular states. The huge amount of data generated will require the development of new and more robust and specific software tools. Abbreviations: CID, collision-induced dissociation; ECD, electron capture dissociation; ETD, electron transfer dissociation; IMAC, immobilized metal ion affinity chromatography; LC-MS/MS, liquid chromatography tandem mass spectrometry; LC, liquid chromatography; LIT, linear ion trap; MALDI TOF, matrix-assisted laser desorption/ionization time-of-flight; MS, mass spectrometry; PK, protein kinase; SCX, strong cation exchange; TiO2, titanium dioxide; PAC, phosphoramidate chemistry; PTM, post-translational modifications.

Acknowledgment. This work was partially supported by S-GEN-0166-2006 grant from the Comunidad Autonoma de Madrid. Note Added after ASAP Publication. Acknowledgment paragraph was missing from the version published ASAP on 3/8/2008; the correct version was published 3/18/2008. References (1) Kalume, D. E.; Molina, H.; Pandey, A. Tackling the phosphoproteome: tools and strategies. Curr. Opin. Chem. Biol. 2003, 7 (1), 64–69.

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