Advances in the Analysis of Protein Phosphorylation

Advances in the Analysis of Protein Phosphorylation

Alberto Paradela* and Juan Pablo Albar

Departamento de Proteómica, Centro Nacional de Biotecnologia, Consejo Superior de InvestigacionesCientíficas, c/ Darwin 3, 28049 Madrid, Spain

Received October 10, 2007

Phosphorylation is one of the most relevant and ubiquitous post-translational modifications. Despiteits relevance, the analysis of protein phosphorylation has been revealed as one of the most challengingtasks due to its highly dynamic nature and low stoichiometry. However, the development andintroduction of new analytical methods are modifying rapidly and substantially this field. Especiallyimportant has been the introduction of more sensitive and specific methods for phosphoprotein andphosphopeptide purification as well as the use of more sensitive and accurate MS-based analyticalmethods. The integration of both approaches has enabled large-scale phosphoproteome studies to beperformed, an unimaginable task few years ago. Additionally, methods originally developed fordifferential proteomics have been adapted making the study of the highly dynamic nature of proteinphosphorylation feasible. This review aims at offering an overview on the most frequently used methodsin phosphoprotein and phosphopeptide enrichment as well as on the most recent MS-based analysisstrategies. Current strategies for quantitative phosphoproteomics and the study of the dynamics ofprotein phosphorylation are highlighted.

Keywords: IMAC • Mass spectrometry • Phosphopeptides • Quantitative phosphoproteomics • Proteinphosphorylation

Introduction

Since the isolation in 1932 by Phoebus A. Levene and FritzA. Lipmann of phosphoserine (first described as serine phos-phoric acid), protein phosphorylation has turned to be one ofthe most biologically relevant and ubiquitous post-translationalprotein modifications. Phosphorylation is a reversible modifi-cation affecting both the folding and function of proteins,regulating essential functions such as cell division, signaltransduction, enzymatic activity, and so forth. According tocomprehensive databases, the estimated number of phosphor-ylation sites in the mammalian proteome could be as high as105 (www.phosphosite.org). Other values calculate at 30–50%the percentage of proteins supposed to be phosphorylated atsome point.1 The relevance of phosphorylation is underlinedby the fact that the number of genes involved in phosphory-lation processes may constitute as much as 2–3% of the entireeukaryotic genome.2 For example, about 2% of the human andmouse genomes encode protein kinases (PK) with 518 and 540distinct PKs determined in human3 and mouse,4 respectively.The analysis of the genome of Saccharomyces cerevisiae hasrevealed the presence of 123 putative protein kinases and 40protein phosphatases, respectively, constituting approximately2% of expressed yeast proteins.5,6 Finally, 251 protein kinasesand 86 protein phosphatases have been identified in theDrosophila melanogaster genome.7 Both counteracting enzy-matic systems, kinases and phosphatases, regulate preciselyprotein phosphorylation and dephosphorylation, and differ in

their kinetic properties, substrate specificities, and cellular ortissue distribution. Among the amino acids that can be phos-phorylated, O-phosphates are by far the most abundant, mostlyattached to serine, threonine, and tyrosine residues. Theoccurrence of phosphorylation on Ser and Thr residues is morefrequent than on Tyr residues, with the ratio of pSer/pThr/pTyr in the order of 1800:200:1.8 The phosphoramidates ofarginine, histidine, and lysine also occur as do acyl derivativesof aspartic and glutamic acid, although they are less abundant.

For the aforementioned reasons, the analysis of proteinphosphorylation is of paramount importance. A comprehensivestudy of protein phosphorylation should include the identifica-tion of phosphoproteins and sites of phosphorylation (phos-phoproteomics), the identification of the proteins (kinases andphosphatases) involved in the phosphorylation process, and adescription of the biological events following the phosphory-lation events. Mass spectrometry (MS) has become a powerfultechnology for proteomics and a method of choice for unbiasedanalysis of protein phosphorylation.9 However, phosphopro-teomics faces the challenge of low-abundance proteins and theoften low ratio of phosphorylated versus nonphosphorylatedproteins found in vivo. Additionally, while some residues areconstitutively phosphorylated, others are only transiently phos-phorylated, in some cases at very low levels. It is also importantto note that often MS-analysis is unable to identify unambigu-ously the phosphorylation site(s) within a peptide. The resultsobtained in collaborative studies focused on the ability ofproteomic laboratories to identify the phosphorylation sitespresent in a relatively simple mixture of phosphoproteins haveclearly demonstrated that phosphorylation site analysis still

* 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 Journal of Proteome Research 2008, 7, 1809–1818 1809Published on Web 03/08/2008

represents a challenge for many laboratories. The wide rangeof results suggested that analytical methods are far from beingwell-established and that a significative percentage of the dataabout protein phosphorylation published to date should bereconsidered carefully (ABRF sPRG Study 2007; www.abrf.org).10 Despite these major drawbacks, recent advances inphosphoproteomics technologies, including sample enrichmentat the phosphopeptide and phosphoprotein levels, MS analysis,phosphorylation site mapping, and quantitative phosphopro-teomics, have made feasible large-scale phosphoproteomicsanalysis 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-basedphosphoproteomics analysis, emphasizing recent advancessuch as phosphopeptide enrichment using TiO2-based station-ary phases, ECD- and ETD-based phosphopeptide MS-analysis,and quantitative phosphoproteomics. Classical approaches areclearly depicted and reviewed elsewhere in more detail.1,8,19

Selective Enrichment of Phosphoproteins andPhosphopeptides

Site-specific analysis of post-translational modifications isusually performed by MS approaches, requiring that themodified protein (phosphoprotein) first be cleaved enzymati-cally or chemically (less frequently) into peptides of a sizesuitable for sequence analysis. However, substoichiometricphosphorylation, wich reduces phosphoanalyte abundancescompared to corresponding unphosphorylated forms, phos-phopeptide inefficient ionization, and specific losses occurringby adsorption to metal or plastics, make highly advisable theuse of phosphoprotein and/or phosphopeptide-specific enrich-ment methods. On most occasions, enrichment of the samplein phosphoproteins followed of protease-specific digestion andMS-analysis is not sufficient to identify the sites of phospho-rylation present in a complex sample and requires a secondenrichment step at the phosphopeptide level. Commercial kitsmake the enrichment process easy, fast, and reproducible,although it has been clearly demonstrated that the differentmethods available differ in the specificity of isolation and inthe set of phosphoproteins and phosphopeptides isolated,20

strongly suggesting that no single method is sufficient for acomprehensive phosphoproteome analysis (Figure 1).

Immunoprecipitation. Antibodies specific to phosphory-lated residues are used to immunoprecipitate total proteinsrather than phosphopeptides. Immunoprecipitation of phos-photyrosine-containing proteins is more frequent than immu-noprecipitation using phosphoserine- or phosphothreonine-specific antibodies as the former are most reliable thanantibodies specific to Ser/Thr-phosphorylated proteins. Thisfact explains why phosphorylation in Tyr residues has beenstudied so intensively in the last years despite its low abun-dance compared to phosphorylation in Ser and Thr residues.21–24

Specific phosphotyrosine binding domains (PTB) have provideda useful tool to profile the global tyrosine phosphorylation stateof the cell.25,26 The reasons that explain why most phospho-serine- and phosphothreonine-specific antibodies are hardlysuitable for phosphoprotein immunoprecipitation are notknown, but it is well-known that frequently these antibodiesare very specific to certain consensus motifs. In practice,immunoprecipitation of phosphoproteins phosphorylated inthese residues usually requires an expensive mixture of different

antibodies. Therefore, their use has only been reported in alimited number of studies.27

Immobilized Metal-Ion Affinity Chromatography (IMAC).IMAC is the most frequently used technique for phosphopep-tide and phosphoprotein enrichment, although it was originallyintroduced for purification of His-tagged proteins.28 Thistechnique allows a higher success rate in phosphopeptideanalysis because it reduces ion suppression effects that wouldotherwise occur with untreated complex mixtures.29 Phosphor-ylated peptides or proteins are bound to the IMAC stationaryphase by electrostatic interactions of its negatively chargedphosphate group with positively charged metal ions bound tothe column material via nitriloacetic acid (NTA), iminodiaceticacid (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 highdensity of histidines. However, immobilized metal ions of Fe3+,Ga3+, and Al3+ have been demonstrated to show better bindingcharacteristics with phosphopeptides. Recently, immobilizedZr4+ has been reported to bind phosphopeptides with highspecificity.30 One of the major drawbacks of IMAC-basedstrategies is the nonspecific binding of peptides containingacidic amino acids, that is, Glu and Asp, and the strong bindingof multiply phosphorylated peptides. Despite following anapparently simple common schema (binding-washing-eluting),IMAC experimental conditions are very variable and careshould be taken as small variations in the experimentalconditions (for example, pH, ionic strength, or organic com-position of the solvents) could drastically affect the selectivityof the IMAC stationary phase. Nonspecific binding of acidicpeptides can be diminished by esterification of carboxylic acidsto methyl esters using HCl-saturated, dried methanol.15 Reac-tion conditions have to be chosen carefully to avoid bothincomplete esterification and side reactions because theyincrease sample complexity. IMAC procedures have becomevery popular rapidly due to its good compatibility with sub-sequent separation and detection techniques such as LC-ESI-MS/MS and MALDI MS.14,17,31

On the basis of measurements of 32P or 33P-radioactivity inwhole cell extracts and in phosphoprotein samples afterenrichment, IMAC-based techniques have been reported torecover up to 70–90% of total phosphoproteins.32 Recently,Machida et al. optimized the conditions for IMAC to enrichfor phosphoproteins.33 According to the authors, Ga3+ was thebest option among the metal ions tested. About 1/10 of the totalprotein was recovered in the eluate when whole cell lysateswere analyzed. Most interestingly, specific phosphoproteinscould be tracked along the enrichment procedure, demonstrat-ing the efficiency of this method.

In addition to IMAC, strong cation exchange chromatography(SCX) has been used in the enrichment of phosphorylatedpeptides. This procedure is based on the fact that under acidicconditions (pH 2.7) tryptic phosphorylated peptides are singlepositively charged and amenable to further separation fromnonphosphorylated tryptic peptides that usually have a netcharge of 2+ at low pH. Although this strategy does not havehigh specificity and the fractions enriched in phosphopeptidesalso contain a high percentage of contaminants, SCX enrich-ment has been used for massive phosphoprotein profiling indeveloping mouse brain18 and in nuclear extracts of HeLa celllysates.16 One of the main advantages of this method is thatcomplex tryptic mixtures can be analyzed directly34 since the

reviews Paradela and Albar

1810 Journal of Proteome Research • Vol. 7, No. 5, 2008

widely adopted multidimensional LC strategy in shotgunproteomics uses a similar SCX-RP approach.

Titanium Dioxide. A promising alternative to the use ofIMAC for the enrichment of phosphorylated peptides was firstdescribed by Pinkse et al.35 The approach is based on theselective interaction of water-soluble phosphates with poroustitanium dioxide microspheres via bidentate binding at the TiO2

surface. Phosphopeptides are trapped in a TiO2 precolumnunder acidic conditions and desorbed under alkaline condi-tions. An increased specificity for phosphopeptides has beenreported, although TiO2-based columns still retain nonphos-phorylated acidic peptides. Peptide loading in 2,5-dihydroxy-benzoic acid (DHB) has been described to efficiently reducethe binding of nonphosphorylated peptides to TiO2 whileretaining high binding affinity for phosphorylated peptides.36

This improved TiO2 procedure was found to be more selectivethan IMAC. An exhaustive analysis has been recently reportedthat describes the relationship between the occurrence of someamino acids and the phospho-specific and nonspecific bindingof peptides using TiO2-based enrichment.37 Two well-charac-

terized peptide mixtures consisting of either 33 or 8 syntheticphosphopeptides or their nonphosphorylated counterparts anddiffering in charge and hydrophobicity were tested. The resultsconfirmed the high selectivity of titanium dioxide for phos-phorylated sequences. Interestingly, drastically reduced recov-ery was observed for phosphopeptides with multiple basicamino acids. The importance of phosphopeptide enrichmentwas highlighted by the fact that 50–75% of the phosphopeptidesfrom both mixtures were not detected by MALDI MS withoutprevious enrichment. One of the main advantages of thisapproach 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 phos-phopeptide isolation has been described. These micro tipsdisplayed similar overall performance as TiO2 columns, al-though more selective isolation of singly phosphorylated pep-tides was observed with ZrO2, whereas TiO2 preferentiallyenriched multiply phosphorylated peptides.38 This approachhas been used to selectively isolate phosphopeptides from the

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

Advances in the Analysis of Protein Phosphorylation reviews

Journal of Proteome Research • Vol. 7, No. 5, 2008 1811

tryptic digestion of a mouse liver lysate. Overall, 248 phosphor-ylation sites and 140 phosphorylated peptides were identified.39

Chemical Modification of Phosphate Groups. Several meth-ods for enrichment of phosphoproteins and phosphopeptidesare based on the specific chemical modification of phosphategroups. Oda et al. described a method for enriching phospho-serine/threonine-containing proteins and for subsequent iden-tification of the phosphoproteins and sites of phosphorylation.The method involved chemical replacement of the phosphatemoieties by an affinity tag and required free sulfhydryls to beblocked before the tagging step.40 One of the main disadvan-tages of this method is that O-linked sugar moieties may alsoundergo �-elimination along the process. Thus, cross-reactivi-ties with glycosylations should be tested carefully. A seconddisadvantage is that �-elimination is not applicable to tyrosinephosphorylation. A different approach reported by Zhou et al.bound phosphopeptides to sulfhydryl-containing compoundsvia phosphoamidate-bonds.41 Modified phosphopeptides arefurther linked to a solid support with immobilized iodoacetyl-groups and obtained as native phosphopeptides after acidelution. This method is applicable to phosphotyrosine-contain-ing peptides as well as those containing phosphoserine andphosphothreonine. However, the amino and carboxyl groupsof the peptides have to be protected to avoid undesiredreactions. Recent modifications of this method used an amino-derivatized dendrimer42 or controlled pore glass derivatizedwith 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. Oneof the major drawbacks of these methods is that reactionconditions have to be monitored very carefully to avoid sidereactions and undesired modifications. Unwanted and partialreactions increase sample complexity while diminishing thetotal yield of the purification. As a result, these methods requirelarge amounts of starting sample with the result that onlyabundant proteins are easily identified. A list of the majorimprovements in phosphopeptide derivatization methods canbe found elsewhere.19

As a summary, it is important to stress that large-scalephosphoproteomics studies should use several phosphopeptideisolation methods. Although most of the methods describedto date are reproducible, it is also clear that they preferentiallyisolate different but somewhat overlapping subsets of thephosphoproteome.20 In other words, no single method issuitable for the isolation of the entire phosphoproteome as themixture 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 onephosphorylation site, while IMAC methods preferentially isolatemultiple phosphorylated peptides. Finally, TiO2 stationaryphases are biased toward acidic peptides with one phosphor-ylation site, although multiphosphorylated peptides have alsobeen eluted from TiO2 columns.44 The differential selectivitycould also explain the variability of the results obtained in somecomparative studies (ABRF sPRG study; www.abrf.org).

Identification of Phosphorylation Sites

Phosphoproteins are usually phosphorylated on a numberof different sites throughout the protein with individual sitesbeing phosphorylated to various degrees. Determination ofphosphorylation sites is a challenging task, but not impossible.As only state-of-the-art mass spectrometers (e.g., FT ICR-MS

or LTQ-Orbitrap) reach the level of resolution required for theanalysis of entire proteins,45 “bottom-up” approaches dealingwith peptides obtained from protein digests are preferred over“top down” techniques dealing with whole proteins. Theproteome to be analyzed is cleaved into peptides of size suitablefor MS analysis, ideally between 700 and 3000 Da, yieldingpeptide mixtures containing both phosphorylated and non-phosphorylated peptides. Enzymatic digestion of phosphop-roteins generate peptides (phosphopeptides) that could containmore than one potential phosphorylation site, making acomplete structural analysis of the phosphopeptides manda-tory. Analysis is performed by tandem mass spectrometry (MS/MS) to unambiguously establish the sequence and to determinewhich residues are phosphorylated.46 There are a number ofdifferent MS methods for determining what residues arephosphorylated in a peptide, although the preferred ones arethose that make use of the specific fragmentation behavior ofphosphopeptides such as triple-quads or Q-TOF instruments.One of the major drawbacks of most of the MS-based methodsis that phosphoester bonds are very labile in the experimentalconditions (CID, collision-induced dissociation) habituallyused, resulting in a loss of phosphoric acid from the peptideand complicating the localization of the site at which themodification was originally attached.

As mentioned previously, phosphopeptides in complexmixtures show decreased ionization rates due to suppressioneffects. Measuring phosphopeptides in negative ion mode canreduce this effect.46,47 However, negative-polarity MS/MSspectra are frequently difficult to decipher, reducing drasticallythe rate of peptide identification. Negative-mode MS/MS ofphosphorylated serine, threonine, and tyrosine residues yieldfragments of -79 Da (PO3

-) and -63 Da (PO2-). Selective

monitoring of phosphopeptide parent ions in negative modebased on their -79 Da ion signature,followed by polarityswitching to obtain positive-ion MS/MS spectra, is a commonlyused method. Howewer, it should be considered that polarityswitching between positive and negative ion modes resultsunavoidably in decreased scanning rates.

The best way to reduce the negative ion suppression effectsis a reduction of sample complexity. This is achieved first byenriching the sample in phosphopeptides, followed by frac-tionation of the enriched sample using powerful separationtechniques such as nanoHPLC or capillary electrophoresis (CE).These separation techniques can be easily coupled online witha wide array of mass spectrometers.48 Capillary electrophoresiscoupled with mass spectrometry (CE-MS) has proven to meetthe requirements of high-throughput, exceptional resolution,and outstanding certainty in protein/peptide identification.49

However, the method requires further improvement to becomea widely used technique as it is limited by the small samplevolumes that can be applied. Therefore, separation of thephosphopeptide-enriched sample is done preferentially byLC-MS and especially nano-LC-MS as these approaches showhigh sensitivity (in the low fentomolar-high attomolar range)and good reproducibility.

Precursor ion scanning is particularly useful for analysis ofphosphorylations that are stable during MS and generatespecific fragment ions that can be monitored, such as tyrosinephosphorylations. Triple quadrupole mass spectrometersequipped with ion-counting detectors provide the highestsensitivity of any mass analyzer for precursor-ion scanning dueto their capability to signal-time average very weak data andreject noise. The first quadrupole (Q1) scans through the entire



mass range, and the peptides passing through are fragmentedin the second quadrupole (Q2). The third quadrupole (Q3) isused for monitoring a specific fragment ion characteristic forthe residue of interest (e.g., m/z 79 for phosphopeptides innegative ion mode or m/z 216.04 for the immonium ion ofphosphotyrosine).

In neutral loss scanning, Q1 and Q3 are simultaneouslyscanned over two different m/z ranges, which differencecorresponds to the m/z value for the neutral molecule that isbeing lost (for a [M + 2H]2 phosphopeptide, this value is 49m/z). This approach is not applied to phosphorylations thatare stable during MS such as tyrosine phosphorylations. 3D-Ion traps do not perform “real” precursor ion scanning orneutral loss analysis. However, the prominent fragment iongenerated after CID-induced phosphate loss in MS/MS mode(MS2 mode) is used for further fragmentation in MS/MS/MSmode (MS3 mode). MS3 spectrum yields more sequenceinformation than the MS2 spectrum, although this informationis not useful for identifying unambiguously where the phos-phorylation site is located. Despite these drawbacks, thisstrategy was used to identify more than 2000 phosphorylationsites from HeLa cells in a large-scale phosphoproteomicsanalysis using a 3D ion trap with software-controlled, neutral,loss-dependent MS3 capabilities.16

Two recent fragmentation techniques have been recentlydeveloped to avoid the prominent loss of the phosphate groupseen during CID. Both fragmentation techniques, electroncapture dissociation (ECD)50 and electron transfer dissociation(ETD),51 have been implemented on FT-ICR, LIT, and 3D ITinstruments. ECD and ETD are applicable to large peptides andsmall proteins and are particularly useful in the analysis ofmultiply charged peptides and in the identification of post-translational modifications. The reason is that post-transla-tional modifications easily lost in the case of CID analysisremain intact when ECD or ETD fragmentation is used, makingthe assignment of phosphorylation sites more precise52 (Figure2). The principle of ECD is to react multiply protonatedpolypeptide cations with low-energy electrons. This inducesfragmentation at the amide (N-CR) bond to produce c-typeand z-type fragment ions. In the case of ETD, the electrontransfer from the radical anions (e.g., singly charged anthraceneanions) to multiply protonated peptides induces fragmentationof the peptide backbone along pathways analogous to thosedescribed for ECD. Recently, Molina et al. have evaluated theuse of ETD for a global phosphoproteome analysis. A total of1435 phosphorylation sites were identified in TiO2-enrichedsamples obtained from human embryonic kidney cells 293T.A comparison of ETD and CID modes demonstrated that CIDyielded 60% less phosphorylation site identifications with anaverage of 40% more fragment ions per fragmentation spec-trum.53 However, although the number of phosphorylation sitesidentified was lower, a significant set of phosphopeptidesidentified when CID was used were specific for this method,indicating that both CID and ETD should be combined for amore comprehensive analysis. Similarly, ETD for peptidefragmentation allowed the identification of 1252 phosphory-lation sites from an IMAC-enriched sample obtained from 30µg of total yeast protein.54 Several examples have been pub-lished demonstrating the feasibility of ECD in the identificationof phosphorylation sites,45,55,56 although large-scale studieshave not yet been published.

Quantitative PhosphoproteomicsQuantitative proteomics is particularly useful to elucidate

highly dynamic processes such as phosphorylation. 2D-PAGEdifferential phosphoproteomics, using either DIGE- or silverstain-based gels, has provided a significative number of results.Although specific patterns seen in 2D-PAGE gels alert aboutthe presence of putative phosphorylated forms of a givenprotein, the use of phosphorylation-specific stains such as Pro-QDPS57 is strongly recommended due to two main reasons: (a)Pro-Q DPS binds directly to the phosphate moiety of phosphop-roteins with high sensitivity and linearity, and (b) the stain is fullycompatible with other staining methods (e.g., DIGE) and modernMS-based analysis. The use of phosphorylation-specific fluores-cent stains such as Pro-Q DPS in 2D-PAGE gels has been reportedfor various biological models.11,12,58–62For example, this approachhas been used in the analysis of the regulation of endosome-specific phosphoproteins obtained after subcellular fraction-ation from EpH4 mammary epithelial cells and followingstimulation by epidermal growth factor (EGF).63 Similarly, ithas been reported that stimulation of human polymorpho-nuclear neutrophils (PMS) through the formyl peptide receptorlike-1 (FPRL-1) alters the protein pattern of PMNs and thatseveral phosphoproteins such as L-plastin, moesin, cofilin, andstathmin are up-regulated or down-regulated under thesecircumstances.64 One of the major drawbacks of 2D-PAGE-based quantitative phosphoproteomics is that only a relativelylimited number of proteins is amenable to detection andidentification. Moreover, large-scale quantitative profiling ofphosphoproteins requires a significative number of high-resolution 2D-PAGE maps to achieve statistically significantdata. This approach has been used to map the changesoccurring in the phosphoproteome during seed filling in oilseedrape. Up to 103 phosphoproteins (70 nonredundant) wereidentified and quantified.65 More details about 2D-PAGE-basedquantitative phosphoproteomics as well as other classicalapproaches are clearly depicted and reviewed elsewhere.66

In addition to 2D-PAGE-based approaches, large-scale quan-titative phosphoproteomics studies are performed by (2D)LC-MS/MS analysis of phosphopeptide-enriched samples. Thesamples to be analyzed are tagged previously by stable-isotopelabeling to make them distinguishable from each other. It isvery important to note that labeling should not affect thechemical properties of identical (phospho) peptides obtainedfrom the samples to be compared (for example, sample A andB) (Figure 3). In other words, “light” and “heavy” phospho-peptides must have identical LC elution times ensuring thatthe ionic environment is equal, and thus, the ratio of therelative intensities can be used to estimate the relative phos-phopeptide abundance in each sample. There are various waysto incorporate stable isotopes into (phospho) peptides. Meta-bolic labeling of proteins using amino acids labeled with 15Nversus 14N, or 13C versus 12C, is an effective approach to stableisotope labeling. This method (called SILAC, stable isotope-derivatized amino acids in cell culture) was originally developedas a simple and accurate procedure that can be used as aquantitative proteomic approach in any cell culture system.67

In the most effective implementation of the method, theaddition of 13C6-arginine and/or 13C6-lysine to the cell growthmedium ensures that all tryptic cleavage products carry at leastone C-terminal labeled amino acid. Protein labeling in excessof 90% is tipically obtained after a small number (6–8) ofpassages. A small number of studies, for example in plants,68

have demonstrated the feasibility of in vivo total 15N metabolic



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

lysine. More than 700 phosphopeptides were identified and 139were found to be differentially regulated at least 2-fold inresponse to pheromone.70 A similar approach has been de-scribed recently to perform a quantitative analysis of ninedifferent phosphorylation sites of the epidermal growth factor.71

As SILAC-based approaches are only applicable to growingcells, several methods have been designed to label peptides orproteins in vitro. These include isotopically distinguishablecommercial reagents specific for different functional groups

Figure 2. Direct comparison of MS/MS spectra obtained when the phosphopeptide SNQRSpSHSpSTLDDIL (M + 3H 607.25) was subjectedto 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 thelow number of fragments and poor sequence coverage obtained when CID was used.



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

cally reacts with free cysteine residues.72 Subsequent ICATversions included changes, for example, incorporation of 13C/12C isotopes, to improve the chromatographic properties of the

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



reagent. As cysteine is a rare amino acid (occurrence in proteinsof ∼1.8%), ICAT is not suitable for quantifying proteins withnone (∼8% in humans) or few cysteines. In practical terms, itsuse should be avoided in quantitative phosphoproteomics, asthe number of (tryptic) peptides containing both a phospho-rylation site and a cysteine residue is extremely low. This keyissue has been solved by the development of reagents thatspecifically label the peptide at the N-terminus and the ε-aminogroup of lysine residues, such as iTRAQ73 and ICPL.74 ICPL andiTRAQ allow multiplexed quantitation of 3 and 8 differentsamples, respectively, and are particularly useful in the analysisof biological systems over multiple time points. Although theyhave been extensively used in differential proteomics, onlyiTRAQ has been recently used to obtain quantitative phos-phoproteomics data.75 Kim et al. reported a method based onIMAC purification of iTRAQ-labeled phosphopeptides whichallowed depicting precisely the phosphorylation events follow-ing CD3 and/or CD28 stimulation in Jurkat cells. In all, 101tyrosine 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 developedfor quantitative phosphoproteomics analysis.77,78 Althoughpromising, the number of reports published to date using thesereagents are not enough to figure out their feasibility inquantitative phosphoproteomics accross different biologicalmodels.

A completely different approach for stable isotope labelingis the incorporation of 16O/18O during the hydrolysis reactioncatalyzed by trypsin or Glu-C. Incorporation of 18O intoC-termini of peptides results in a mass shift of 2 Da per 18Oatom. As the hydrolisis catalyzed by trypsin and Glu-C intro-duces two oxygen atoms, it results in a 4 Da mass shift wich isgenerally sufficient to differentiate labeled from unlabeledsamples. This strategy combined with IMAC-based purificationof phosphopeptides allowed the quantification of the changesin the phosphoproteome of the cell body and pseudopodiumof chemotactic cells. A total of 228 unique phosphopeptidescorresponding to 197 proteins were identified.79

Finally, label-free methods for quantitative analysis have alsobeen described as an alternative to expensive isotope-basedlabeling techniques. As with isotopic labeling, these methodsare based on the measurement and comparison of the massspectrometric signal intensities of peptide precursor ions.However, as precursor ions are not isotopically distinguisable,samples to be compared have to be analyzed in independentruns. Afterward, the ion chromatogram for every peptide isextracted from an LC-MS/MS run and its mass spectrometricpeak area integrated over the chromatographic time scale. Inpractical terms, label-free comparison of integrated peakintensities obtained in different LC-MS/MS runs requires acombination of accurate mass and stringent reproducibility insample processing and chromatography. Special software hasbeen developed to align LC-MS/MS runs prior to identifyingcorresponding peptides between different experiments.80,81 Todate, one report successfully applied label-free quantificationof IMAC-purified of phosphopeptides. A total of 714 phosphor-ylation sites on 223 phosphoproteins were identified. A numberof proteins that significantly changed phosphorylation state inresponse to vasopressin treatment was identified.82

The final goal of quantitative phosphoproteomic studies isto elucidate the dynamics of protein phosphorylation acrossdifferent cellular physiological and pathological states. In otherwords, the objective is to obtain a picture as complete as

possible of the evolution of the phosphoproteome as a functionof time, stimulus, and other parameters. For this purpose, it isessential to develop and integrate global and quantitativemethods. Blagoev et al. described the dynamic profile of 81affinity-purified tyrosine-phosphorylated proteins after stimu-lation by epidermal growth factor (EGF) of three different cellpopulations.83 More recently, the temporal dynamics of 6600phosphorylation sites on 2244 proteins after stimulating HeLacells with epidermal growth factor has been published.84

Interestingly, results showed that phosphorylation is regulateddifferently on different sites within the same protein suggestingthat the dynamics of phosphorylation should be measured sitespecifically rather than for the protein as a whole. Consideringthe complexity of the phosphoproteome dynamics, integrativeapproaches like this are essential for a systematic understand-ing of cellular behavior.

Summary

The analysis of the phosphoproteome is one of the mostexciting and challenging tasks in current proteomics research.Large-scale phosphoproteome analysis is now an affordabletask since the development of methods for the purification ofphosphoproteins and phosphopeptides as well as the new massspectrometry technologies developed for the structural analysisof the enriched samples. However, many challenges lie ahead.Purification methods are far from rendering homogeneousresults as it has been demonstrated that they tend to isolate pre-ferentially specific subsets of phosphopeptides. Comparativestudies have concluded that different proteomic strategies arecomplementary to each other. Therefore, the analysis of thephosphoproteome requires a combination of multiple techniques.

In the last years, the interest has turned on the study of thephosphoproteome as a highly dynamic environment. Classicalproteomic strategies such as 2D-PAGE combined with the useof phosphoprotein-specific stains such as Pro-Q DPS are nowcomplemented with the combined use of stable isotope labelingand phosphopeptide enrichment followed by LC-MS/MS(MS)analysis. These strategies will permit the analysis of theevolution of the phosphoproteome across different cellularstates. The huge amount of data generated will require thedevelopment of new and more robust and specific softwaretools.

Abbreviations: CID, collision-induced dissociation; ECD,electron capture dissociation; ETD, electron transfer dissocia-tion; IMAC, immobilized metal ion affinity chromatography;LC-MS/MS, liquid chromatography tandem mass spectrom-etry; LC, liquid chromatography; LIT, linear ion trap; MALDITOF, matrix-assisted laser desorption/ionization time-of-flight;MS, mass spectrometry; PK, protein kinase; SCX, strong cationexchange; TiO2, titanium dioxide; PAC, phosphoramidate chem-istry; PTM, post-translational modifications.

Acknowledgment. This work was partially supportedby S-GEN-0166-2006 grant from the Comunidad Autonomade Madrid.

Note Added after ASAP Publication. Acknowledg-ment paragraph was missing from the version published ASAPon 3/8/2008; the correct version was published 3/18/2008.

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Advances in the Analysis of Protein Phosphorylation

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