Chapter 5 Phosphorylation-specific analysis strategies for mass spectrometry: enhanced detection of phosphorylated proteins and peptides

Chapter 5

Phosphorylation-specific analysis strategies for mass spectrometry: enhanced detection of phosphorylated proteins and peptides Allan Stensballe and Richard J. Simpson

5.1

INTRODUCTION

Protein phosphorylation is one of the most widespread and arguably the best understood post-translational modifications (PTMs). Virtually all cellular processes are regulated in one or more ways through protein phosphorylation and dephosphorylation, and the identification of kinases, their substrates and the specific sites of phosphorylation are vital to a molecular understanding of signal transduction. The modification of side chains of specific amino acids by a phosphate moiety (H3PO4) is associated with regulation of a range of basal cellular processes like metabolic pathways, signal transduction by activation of kinase cascades, membrane transport, cell growth, division, differentiation and memory [50,51,97,99]. In mammals, more than 80% of known oncoproteins are receptor or cytoplasmic tyrosine kinases which underscores the essential role phosphorylation plays in regulating cell function [98]. Advances in cell signaling and knowledge of protein phosphorylation will therefore enable new strategies for attacking diseases that are caused or exacerbated by faulty signaling in cells––among them are cancer, diabetes and disorders of the immune system. In the human genome, it is now hypothesized that up to 30% of all gene products may be targets for protein kinases [50,97]. Localization of the phosphorylation site(s) in a protein identifies important regulatory domains and may also reveal which protein kinases and Comprehensive Analytical Chemistry 46 Marko-Varga (Ed) Volume 46 ISSN: 0166-526X DOI: 10.1016/S0166-526X(05)46005-5 r 2005 Elsevier B.V. All rights reserved.

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phosphatases regulate their activity and thereby help elucidate the biological function and significance. From a physiological stance, the determination of the regulatory in vivo phosphorylation sites naturally occurring in cells, tissue or cell culture provides most information, however, such studies are generally more technically demanding. This is mainly due to in vivo phosphorylated proteins involved in signaltransduction pathways and therefore are of low abundance in cells due to a low copy number of interacting proteins and only a fraction of the protein population (generally less than 5%) being phosphorylated in response to stimuli in vivo [97]. The global identification of protein phosphorylation has become one of the major challenging sub proteomes that are now starting to be scrutinized in laboratories around the world. The ‘‘phosphoproteome’’ is a snapshot of the entire assemblage of protein phosphorylations in a given organelle, tissue or organism at a given time point. From single protein analysis to maybe characterization of many hundred posttranslationally modified proteins in large scale phosphoproteome investigation must be identified, characterized and quantified in order to describe the cellular dynamics and eventually understand the biological system investigated. Direct phosphoproteome analysis presents specialized analytical problems due to a large dynamic expression range (protein abundance) and a huge diversity of protein expression profiles (multiple protein forms, variable stoichiometry). In addition to protein phosphorylation, O-glycosylation and O-sulfonation are also used to regulate protein function in cellular processes [82,114,158]. O-linked b-N-acetylglucosamine (O-GlcNAc) is a nucleocytoplasmic modification more analogous to phosphorylation than to classical complex O-glycosylation. In several instances, O-GlcNAc maps to the same or adjacent sites as phosphorylation, termed the ‘‘yin-yang’’ modification [55,259]. The close chemical relation of the two modifications makes distinguishing between O-phosphate and O-GlcNAc difficult when mapping sites of serine and threonine PTM using b-elimination/ Michael addition methods [245]. O-sulfonation is the transfer of a sulfonate group (SO1
3 ) to tyrosine, threonine and serine residues, which is implicated in multiple functions including protein assembly, protein–protein interactions and signal transduction. Phosphate and sulfate moieties have almost identical masses differing by merely 9.4 mmu (sulfate addition 79.9568; phosphate addition 79.96633) whereby only high mass-accuracy mass spectrometers can measure the molecular mass with the appropriate mass accuracy to distinguish the two molecular 276

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species. Inevitably, analysis strategies must include discriminatory methods that can be used to unambiguously differentiate between O-phosphorylation, O-sulfonation and O-glycosylation. 5.1.1

Chemical properties and abundances of phosphoamino acids

The specificity of the complement of protein kinases and protein phosphatases in prokaryote and eukaryotic organisms differ significantly, although pervasive parallels between homologous families of enzymes exist [120]. Bacteria favor the use of histidine and the carboxyamino acids, aspartic acid and glutamic acid, as phosphoacceptors whereas the typical phosphoacceptors in multicellular organisms are the hydroxyamino acids serine, threonine and, less often, also tyrosine residues. Lysine, arginine and cysteine have also been identified as phosphate acceptors in both prokaryotes and eukaryotes [255]. In vertebrates the ratio of phosphoamino acids vary significantly. The ratio of phosphorylation on serine/threonine/tyrosine is 1800:200:1 [77], whereas phosphohistidine is currently speculated to be 10–100-fold more abundant than phosphotyrosine, and recently a human histidine kinase (HHK) was classified as an oncodevelopmental marker kinase [152,238]. The chemical stability of phosphorylated amino acids depends on the amino acid donor as shown in Fig. 5.1. The chemical properties of the

Fig. 5.1. Chemical stability of phosphorylated amino acids (+, stable phosphoamino acid;7, less stable phosphoamino acid;—labile phosphoamino acid). Adapted from Klumpp and Krieglstein[126]. 277

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individual phosphoamino acids, the biological origin and cellular abundance of phosphoproteins adds serious constraints to sample handling, as well as the set of analytical methods available for isolating and characterizing phosphoproteins. Only the O-phosphates (pS, pT, pY) are stable under acidic conditions while the N-phosphates (pR, pH, pK), S-phosphates (pC) and acyl-phosphates (pD, pE) are acid-labile [222]. Acid treatment of putative phosphoproteins of the latter class will therefore be detrimental and hinder detection. As such, the existence of acid-labile phosphates are currently suspected to be largely overlooked [126]. In contrast, the chemical stability of the phosphomonoester bond of O-phosphates under acidic conditions is considerably higher than the rate of hydrolysis of peptide bonds in the protein backbone [58]. Under alkaline conditions, only phosphotyrosine, phosphohistidine, phospholysine and to a lesser extent phosphothreonine remain stable. At the same time analytical methods have been developed that rely on the base lability of phosphoserine and phosphothreonine to form the thioacyl derivative of the dehydroamino acid and subsequent derivatives [73,161,176,230]. Mass spectrometry (MS)-based strategies for analyzing O-phosphates are now becoming reasonably robust while the analysis of N- and acyl-phosphates can only be analyzed indirectly by MS [222]. In this chapter, only the analysis of O-phosphorylated proteins will be presented. The property of a double-negative charge and the capacity for forming extensive hydrogen-bond networks with the four phosphoryl oxygens confer special characteristics to phosphopeptides. The introduction of one or often several phosphoryl groups in a peptide shifts the hydrophobicity/hydrophilicity indices for the polypeptide to an increased hydrophilicity [32,195]. The Bull and Breese index is a measure of a peptide’s hydrophobicity, a positive value being associated with hydrophobic peptides and a negative value with hydrophilic peptides. For phosphopeptides, the Bull and Breeze index can be mimicked by mutating phosphoserine or phosphothreonine residues to glutamic acid residues [91]. 5.1.2 Toolbox for micro-characterization of phosphoproteins by mass spectrometry

The analytical power of MS for identification and characterization of posttranslationally modified proteins has been pivotal for recent major advances strategies for characterization of phosphoproteins. MS 278

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measures the mass-to-charge ratio (m/z) of biomolecules with very high mass accuracy enabling detection of modification-specific mass increments or mass losses compared to the unmodified biomolecule. Different configurations of mass spectrometers provide diverse means to identify and characterize phosphoproteins at full-length protein, polypeptide or amino acid residue level. Mass spectrometric detection of phosphoproteins and phosphopeptides is facilitated by the 80 Da increase per phosphate group present in the protein. Hence, the phosphorylation of a tyrosine residue increases the nominal molecular mass of this residue from 163 to 243 Da. This small mass increment can in principle be detected by accurate mass determination of the intact protein, however, more often measured at the peptide level. Analysis of protein phosphorylation is one of the major analytical challenges of protein characterization. Numerous MS-based strategies have been devised and applied for the discovery and characterization of phosphoproteins and phosphoproteomes (reviewed in [110,150,151,155,200,222,223,255]). Most strategies seek to obtain experimental evidence for protein phosphorylation by taking advantage of combinations of many different technologies [12,144]. The ‘‘tool box’’ of phosphoprotein analysis includes techniques for specific detection and visualization of phosphoproteins; enrichment/purification approaches to isolate phosphoproteomes or phosphoproteins; chromatography techniques for reducing complexity of protein mixtures and sample preparation before MS analysis, chemical derivatization strategies to eliminate and convert phosphoamino acid residues into more stable and tractable species; and MS techniques for analysis of full-length proteins and protein digests for identification of the phosphorylated residues. Novel or improved technologies continued to be developed due to the increasing demand for more efficient, sensitive and comprehensive strategies. The far majority of phosphoprotein and phosphoproteome studies are currently carried out by combinations of gel-based and/or gel-free separation methodologies and a range of affinity enrichment methodologies for capturing the subset of phosphorylated proteins and peptides prior to MS analysis [109,144,150,151]. Strategies for identification of phosphorylation sites in proteins by MS typically include the following sequential stages: (I) (II)

Isolation, detection and visualization of intact phosphoproteins Generation of peptides by sequence-specific proteolytic cleavage of phosphoproteins 279

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(III)

(IV)

Characterization of the peptide mixtures by phosphopeptide mass mapping, selective mass spectrometric detection methods (precursor ion scanning or constant neutral loss scanning) and schemes for the selective retention/affinity purification of the phosphorylated species, identification of the phosphopeptides by MS and/or phosphopeptide sequencing by tandem mass spectrometry (MS/MS); and Quantitative analysis of the state of phosphorylation and data mining.

In this chapter, we will outline the techniques used in MS-driven strategies at each stage of current phosphoprotein and phosphoproteomics research.

5.2

GLOBAL GEL-BASED PHOSPHOPROTEIN ANALYSIS

The global analysis of protein phosphorylation may be probed by twodimensional gel electrophoresis, 2-DE monitoring phosphorylated proteins by either radiolabeling or by Western blot using anti-phosphoamino acid-directed monoclonal antibodies. So far the only technique able to efficiently separate thousands of proteins or posttranslationally modified proteins from one single sample is 2-DE. Since its introduction in 1975, 2-DE has evolved as a powerful and sensitive separation method [61,125,174,201]. The high resolving power of 2-DE enables the separation of the high number of protein species likely to be present in the phosphoproteome. Owing to the zwitterionic character of proteins, the electrophoretic mobility of each protein is a characteristic value (i.e., isoelctric point and molecular mass). Nearly all PTMs and protein degradations change the molecular mass and may affect the isoelectric point (pI) of proteins, so that the observed pI of a protein deviates from that calculated from the DNA sequence. The orthogonal separation of proteins by pl followed by molecular weight enables separation of similar but differentially modified forms of a given protein. A ‘‘train-of-spots’’ pattern observed by 2-DE indicates protein phosphorylation. The negatively charged phosphoryl moiety of phosphoproteins enable phosphorylated forms of a protein to be visualized by 2-DE, where phosphorylated species focus on the acidic side of the non-phosphorylated fraction. Similarly, the introduction of charge heterogeneity by glycosylation, oxidation, formylation, acetylation, methylation, ubiquitination and deamidation also 280

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generate such a pattern which necessitates further validation of phosphorylation events [23,149,211,240]. At the level of full-length proteins, two-dimensional polyacrylamide gels can reveal phosphoproteins in expressed proteomes or multi-subunit complexes by a differential display using protein phosphatases [102,254]. Phosphatase treatment can be applied as a convenient readout by enzymatic treatment of proteins with alkaline phosphatase or lambda phosphatase. The protein samples are then divided into two aliquots, one of which is dephosphorylated using the protein phosphatase and the other is not treated with the enzyme. Each of the two samples is then subjected to parallel 2-DE (Fig. 5.2). By comparing the resulting protein pattern of the two electrophoretic gels the phosphoproteins in the treated aliquot can be identified due to a shift to more basic positions on the gel. Two-dimensional blue-native electrophoresis is a tool for functional proteomics of signaling complexes [31,35,213,214]. In the first dimension, blue-native PAGE employs Coomassie dyes to introduce charge shifts on

2-DE gel Acidic

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Small Before PPase treatment

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Fig. 5.2. 2-DE pattern before and after phosphatase treatment. Courtesy of Dr. D. B. Kristensen [254]. 281

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proteins in order to separate intact protein complexes according to their size followed by Tricine SDS-PAGE in the second dimension (Fig. 5.3A). Bykova and co-workers demonstrated the isolation of protein complexes in the matrix fraction isolated from potato tuber mitochondria. The bluenative PAGE allowed the separation of multi-subunit respiratory chain

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Fig. 5.3. Two-dimensional resolution of protein complexes by blue-native/ tricine SDS-PAGE. (A) Blue-native/SDS-PAGE; (B) left Coomassie staining; right, phosphor image (Formate dehydrogenase (FDH); pyruvate dehydrogenase E1a-subunit (PDH)).Courtesy of Dr. N. V. Bykova [36]. 282

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complexes of inner mitochondrial membrane complexes from potato within one single gel and allowed a size determination of membrane proteins and membrane-bound protein complexes (I to IV;[164]. Next, the phosphoproteins could be identified based on 32P labeling (Fig. 5.3B) and phosphoproteins characterized by combinations of immobilized metal affinity chromatography (IMAC) enrichment and liquid chromatography–MS/MS (LC-MS/MS) analysis [35]. In studies of biological systems the incorporation of 32/33P is widely applicable to positively identify phosphorylation events [35,38,107,138,227]. This radioactive labeling of proteins is the most sensitive method for determining which proteins or peptides are phosphorylated. A common approach is the in vivo or in vitro labeling of kinase substrates followed by isolation by polyacrylamide gel electrophoresis and then visualized by autoradiography. For proteome studies using 2-DE, the incorporation of the radioisotopes 32/33P enables global studies of this subset of post translationally modified proteins in response to stimulation of cells with mitogens, etc., [138]. However, the in vivo incorporation of radioactive isotopes is inefficient because of the presence of endogenous ATP pools within cells. Therefore, almost 100–1000-fold higher amounts of radioactively labeled ATP are required to achieve a degree of in vivo phosphorylation that is sufficient for sensitive detection in comparison to in vitro kinase assays [227]. The radioactive labeling provides an efficient means for tracking phosphorylated proteins and peptides during sample handling and means for quantification. By radiolabelling the phosphoproteome (i.e., the spatiotemporal complement of phosphorylated proteins) with [32/33P]phosphate and comparing the radioactivity pattern of 2-DE gels before and after stimulation, the proteins involved in a signal transduction pathway can be mapped [79,110]. The use of radioisotopes has several important limitations including the health hazards associated with working with radioactivity, and incomplete incorporation of the radioisotope. Specific visualization of phosphoproteins isolated by 2-DE gels and SDS–PAGE is possible by using antibodies for multiple antiphosphoamino acids enabling immunoblotting [61,71]. Fluorescent dyes specific for staining of phosphopthreonine-, phosphoserine- and phosphotyrosine-containing proteins in electrophoretic gels are commercially available [72,186,217]. This stain helped Collins et al. [54] evaluate the depletion of phosphoproteins from protein mixtures using Ga(IMAC) and Hayduk et al. [85] identify phosphoproteins in a twodimensional electrophoresis map of Chinese hamster ovary cells. 283

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MS-based characterization of proteins visualized in 2-DE gels by using phosphorylation-specific dyes or radiolabeling may be obscured by the presence of abundant, unmodified or co-migrating proteins. A reduction of the complexity of protein samples to be analyzed by 2-DE may be advantageous, e.g. by prior purification or enrichment of modified proteins or protein complexes. Also the sensitivity level of 32/33P and the limited amount of protein load exceeds the sensitivity level of common MS method for phosphorylation mapping explaining why the number of novel phosphorylation sites mapped from 2-DE are currently very limited [35,94,165,232]. Although, gel-based approaches enable identification of many differentially phosphorylated proteins by 2D-PAGE, this approach excludes most membrane proteins due to their hydrophobicity and low abundance [116,253].

5.3

NEO-CLASSICAL STRATEGIES FOR PHOSPHOPROTEIN ANALYSIS

Prior to the emerging MS-based methods, examination of protein phosphorylation by classical biochemical strategies usually involved degrading the phosphoprotein chemically or enzymatically into small peptides. This was followed up by several rounds of reverse-phase highperformance loquid chromatography (RP-HPLC) purification in order to isolate a single peptide and preparation for composition and sequence analysis [141]. Sequence analysis was commonly accomplished by Edman degradation of peptides, where phosphorylation sites are determined by monitoring the release of the radioactivity during the Edman cycles or release of chemically modified phosphoamino acids [2,37,43,86,146]. Phosphopeptide sequencing by MS has largely replaced the traditional Edman sequencing due to a higher sensitivity and the ability to sequence directly from peptide mixtures (discussed in [183]). For visualizing low amounts of phosphoproteins and low stoichiometry phosphorylation the sensitivity of radioactive labeling is unparalleled. Presently, the 32/33P labeling enables the possibility to determine the sites of phosphorylation in a peptide by solid-phase Edman degradation. This simple, efficient and very sensitive method was first demonstrated by Wettenhall and co-workers and later refined by the Morrice laboratory [38,247]. In vivo or in vitro 32/33P-labeled proteins are prepared for solid-phase Edman degradation by enzymatic 284

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digestion and fractionation of peptides by HPLC. In each fraction the radioactivity can be measured by Cerenkov counting, online radio nucleotide monitor or spotted onto a PVDF membrane for subsequent autoradiography or phosphoimaging. The HPLC effluent can be collected for comprehensive analysis of 32/33P-containing fractions by MS followed by sequencing of candidate phosphopeptides by solid-phase Edman degradation or tandem MS [38,45]. The site of phosphorylation is revealed by monitoring the cycles at which the free phosphoamino acids elute corresponding to an elevated level of activity (Fig. 5.4). This elegant approach has recently enabled the comprehensive analysis of the human Insulin Receptor Substrate-1(IRS1), identifying at least 18 sites of serine and threonine phosphorylation [75]. Solid-phase Edman degradation is a powerful and very sensitive technique, especially when combined with MS; however, it has the disadvantages that require the use of radioactively labeled material and either prior knowledge of the protein sequence or the ability to make some assumptions about the peptide sequence. Alternatively, radiolabeled phosphopeptides or phosphoamino acids can be separated by two-dimensional thin-layer chromatography (2D-TLC) [30,58] and identified by autoradiography (see Fig. 5.5). 2D-TLC/thin-layer electrophoresis (TLE) analysis is a highly usable

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Fig. 5.4. Solid-phase Edman sequence analysis of phosphopeptides. An aliquots of 32P-labelled peptide derived from NUAK2 was covalently coupled to a Sequelon arylamine membrane and analyzed on an sequenator. 32P radioactivity was measured after each cycle of Edman degradation. The site of phosphorylation is revealed by a major release of radioactivity. Courtesy of Dr. N. Morrice.

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and a common supplement to gel electrophoresis or MS-based methods [108,171,241]. Phosphoamino acid analysis by 2D-TLC can reveal the presence and abundance of the phosphoamino acids phosphothreonine, phosphoserine and phosphotyrosine (Fig. 5.5A). Radiolabeled phosphopeptide and phosphoprotein mixtures can be visualized by 2D-TLC/ TLE [170,187]. Nu ¨ hse and co-workers have demonstrated the applicability of 2D-TLC to display differential cellular responses (Fig. 5.5B) [171]. A number of the radioactive peptides are equally abundant in both samples, indicating that the differences indeed are a result of a biological response and not from losses during sample handling. Elution of the phosphopeptides from the 2D-TLC plated also enable sequencing by Edman degradation or mass spectrometry [3,239]. However, labeling of phosphoproteins with radioactive [32/33P]phosphate is

Fig. 5.5. Two-dimensional TLC/TLE of phosphopeptides and phosphoamino acids. (Upper panel) Phosphoamino acid analysis of serine phosphorylated protein by two-dimensional TLC. Theoretical locations of phosphothreonine (pT), phosphoserine (pS) and phosphotyrosine (pY) are indicated. Adapted from Stensballe [231]. (Lower panel) Two-dimensional TLC/TLE of stimulated versus control phosphopeptide samples (arrows indicate upregulated or unregulated expression levels). The origin (site of sample application) is indicated by ‘‘+’’. Adapted from [Nuhse et al.] [171]. 286

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required and the amount of protein needed for the latter experiments far exceeds the levels needed for MS-based strategies.

5.4

MASS SPECTROMETRY-DRIVEN PHOSPHOPEPTIDE MAPPING AND SEQUENCING

MS was first applied for assignment of phosphorylation sites using fastatom bombardment ionization MS (FAB-MS); 252Cf plasma desorption MS (PD-MS); liquid secondary ion MS (LSI-MS) and sector mass analyzers [41,52,60,95,191,195,218]. Sample size requirements of mass spectrometers using these ionization techniques were in the nanomole level, often exceeding the amount of material available, practically, from biological samples. For the last decade mass spectrometers using the two complementary ionization techniques, matrix-assisted laser desorption ionization (MALDI) and electrospray ionization (ESI), have almost ousted all other ionization techniques for phosphoprotein analysis. ESI differs from MALDI in several fundamental aspects. ESI is a very gentle and sensitive ionization technique that permits labile molecules such as phosphorylated species to ionize without fragmentation, whereas MALDI readily promotes prompt and metastable fragmentation during ionization [10]. ESI is directly compatible with a range of liquid chromatography techniques, such as HPLC and capillary electrophoresis (CE). MALDI is advantageous for static experiments as the sample is in solid state. In contrast to MALDI, then ESI readily generates multiply protonated molecular ions that efficiently can be sequenced by low-energy collision-induced dissociation (CID) as well as providing multiply charged ions required for electron capture dissociation (ECD). Most notably, detection efficiencies of phosphopeptides as compared to their unphosphorylated counterparts due to selective suppression of their ionization are significantly more pronounced in MALDI than ESI (Hanno Steen, personal communication; [233]). An exception is element MS by inductively coupled plasma MS (ICP-MS), which was newly introduced as a robust and specific method in phosphoproteomics [250,251]. Profrock and co-workers [197] have recently demonstrated that capillary liquid chromatography interfaced to ICP-MS can determine the degree of phosphorylation in phosphoproteins and phosphopeptides containing cysteine and/or methionine residues by detection and quantification of elemental phosphorus and 287

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sulfur (see Fig. 5.6). The detection limit achieved for the LC-ICP-HR MS runs is approximately 0.1 pmol of phosphopeptide injected [250]. 5.4.1

Preparation of phosphoproteins for MS analysis

Processing of a phosphoprotein sample prior to analysis has to be compatible with subsequent MS techniques for a successful outcome. In many instances, such as tryptic peptide solutions derived from in-gel digestions of phosphoproteins separated by SDS-PAGE, salts, ionic detergents and other contaminants like high-molecular-mass surfactants are found and peptide recovery is frequently poor. Detection sensitivity of MS is not usually limited by the absolute amount of analyte but rather the presence of such contaminants, which increase chemical noise in the mass spectrum and can cause signal suppression [134]. Various chromatography steps are often imperative for efficient sample cleanup [70]. For a comprehensive list of common contaminants and their maximum tolerable concentration see [223]. The initial step for micro-characterization of phosphoproteins separated by SDS-PAGE or 2D-PAGE is the in-situ enzymatic digestion of protein and recovery of the resulting peptides [220]. The recovery of phosphopeptides from gel material is generally incomplete leaving 5– 50% phosphopeptides still in the gel [38,231,260]. At the same time phosphopeptides are notoriously difficult to handle since they have a great tendency to bind to metal or plastic which again reduces their yield. Thus, handling of phosphopeptide samples must be reduced to an absolute minimum. The non-ionic detergent n-octyl glucoside (0.1%) is known to aid protein solubilization and enhances the response of peptides in MALDI-MS [78,117]. Additionally, this detergent reduces adsorption of phosphopeptides to plastic ware that often significantly contributes to losses of phosphorylated peptide species (Nick Morrice, personal communication). The electrostatic surface potential of phosphorylated amino acid residues can interfere with the cleavage efficiency of enzymes, such as trypsin, which are frequently used for protein characterization by MS [22,38,216]. Phosphopeptides are usually produced by cleavage of the protein with a sequence-specific protease or a range of chemical agents, which induces cleavage of the peptide backbone. Trypsin is a favorite protease in many laboratories due its high activity and specificity and because it generates peptides of a size that is appropriate for mass analysis, viz 500–4000 Da. Phosphorylation-generated trypsin-resistant 288

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Fig. 5.6. Phosphorylation profiling in protein analysis using quadrupole-based collision-cell ICP–MS as phosphorus-specific detector. (A) Alignment of the 31 P trace and the ESI-TIC of b casein separated by capLC reveals the singly phosphorylated peptide. (B) Data deconvolution of the ESI-TIC at the retention time of the previously detected peak in the 31P trace. Courtesy of Dr. D. Profrock.

bonds have been generalized as Arg-Xaa-pSer, Lys-pSer, Arg-pSer and Arg-pThr, but not Arg-Xaa-pThr and Lys-pThr [38]. The reduced cleavage efficiency may lead to large phosphopeptides outside the ideal mass range of 5–20 amino acid residues (i.e., 500–2500 Da), which are 289

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not readily detected or sequenced using general mass spectrometric approaches. To reduce the unfavorable phosphopeptide size, several different proteases or dual-enzyme digestion have been investigated [81,215,216,253]. Endoproteinases Lys-C, Glu-C and Asp-N are suitable for generating complementary sequences. Alternatively, Fouriertransform MS (FTMS) employing ECD enables sequencing of large peptides and small proteins with labile modifications [46,221,234]. A targeted chemical strategy applicable for phosphoserine and phosphothreonine residues has been demonstrated that converts these residues into protease-sensitive lysine analogs (aminoethylcysteine and b-methylaminoethylcysteine, respectively) [127,128,209] using b-elimination followed by Michael addition of 2-aminoethanethiol. Creating phosphoamino acid specific cleavage sites for trypsin and Lys-C leads to a facile assignment of phosphorylation sites by MS and MS/MS, where the C-terminal residue would always be a phosphorylated residue, resulting in a unique y1 ion. O-glycosylated peptides are known to undergo the same b-elimination reaction as phosphoserine and phosphothreonine which can be a drawback for this strategy. 5.4.2 Discrimination effects and hindered detection of phosphopeptide by mass spectrometry

Phosphopeptides are often detected with low efficiency or not at all by MS, especially when they are components in complex peptide mixtures. The reduced desorption and ionization efficiency (flyability) of phosphopeptides originate from the inherent properties of the negatively charged phosphoryl group, which affect the physiochemical characteristics of the phosphopeptide. The flyability of phosphopeptides are strongly reduced when compared with their non-phosphorylated counterparts. Two studies have quantitated the differences in ionization efficiencies for MALDI and ESI. For MALDI, the response factor for non-phosphorylated peptide ENDYINASL was 10 times higher than for the tyrosine-phosphorylated peptide [47]. The difference in ionization efficiencies was not as dramatic for ESI, where Miliotis et al. [163] determined an intensity ratio of 1:1.5:5 for the doubly, singly and non-tyrosine-phosphorylated peptide ALGADDSYYTAR. These empirical observations are often referred as ‘‘suppression effects’’ and this term covers, respectively discrimination effects and variations in the desorption and ionization efficiency of equimolar peptides. 290

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The peak intensities of phosphopeptides in mixtures vary due to factors such as the complexity of peptide mixture [33], existence of charged side-chains and aromatic amino acids [53], peptide hydrophobicity and gas-phase basicity [5,113,131,180], proton affinity and mobility [33], size [180], and the existence of secondary structures [246]. Kratzer et al. [131] proposed two important effects responsible for signal suppression of certain types of peptides in MALDI using the a-cyano-4-hydroxycinnamic acid (4-HCCA) matrix [131]. A predesorption effect, where the limited number of useful desorption sites in the matrix lattice is occupied by hydrophobic peptides, leading to a low incorporation rate of hydrophilic peptides in the 4-HCCA matrix lattice. The more hydrophobic peptides the higher the incorporation rate. A postdesorption effect, where the peptides compete for a limited number of ionizing charges. The method of matrix and analyte application onto the target, rate of matrix crystal growth, matrix micro-crystal structure and the pH of the sample and matrix solution have also been determined to play important roles in signal suppression [53,96]. The increased hydrophilicity and acidic character of phosphopeptides due to the chemical properties of the phosphoryl group are generally thought to be responsible for this effect in MALDI. In electrospray, analytes with high affinity for the surface of ESI droplets (surface-active analytes) have higher ESI response. The increased hydrophilicity of phosphopeptides leads to reduced surface-active properties and thus, a decreased ESI response [44,118]. However, recent results indicate that selective suppression of phosphopeptides does not generally occur in ESI. Singly and doubly phosphorylated peptides were found generally not to show lower ionization efficiencies than their unphosphorylated counterparts during micro-scale capillary HPLC/MS [224,225]. However, impurities or additives increase the chemical background that interferes with detection of the analyte where the phosphoryl group have a high propensity for formation of alkali–metal adductions [M
nH+mNa](m
n)+ and [M
nH+mK](m
n)+ [208]. 5.4.2.1 Current approaches for circumventing suppression effects Analytical problems associated with suppression effects reducing the detection of phosphopeptides present in complex peptide mixtures have been sought circumvented by multiple approaches. Addition of volatile ammonium salts to the MALDI matrix spermidine, spermine, ammonium tartrate, ammonium acetate and, in particular, diammonium 291

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citrate or solvent additives like phosphoric acid in ESI have been found to enhance the phosphopeptide ion response [13,14,121,148]. The most efficient co-matrix for improving the desorption and ionization efficiency of phosphopeptides using MALDI is the combination of o-phosphoric acid and 2,5-dihydroxybenzoic acid (2,5-DHB), which improves detection of phosphopeptides in the peptide mixtures as well as intact phosphoproteins [124,233]. The practical utility of this matrix additive has been demonstrated by the MALDI MS analysis of bovine PP3 using either formic acid (FA) or o-phosphoric acid (PA) in combination with 2,5-DHB matrix (Fig. 5.7) [233]. The relative intensity enhancement is most notably for multi-phosphorylated peptides (indicated by asterisks), whereas singly phosphorylated peptides show minor improvements in desorption and ionization efficiency resulting in an improved signal-to-noise ratio. Phosphorylated peptides exhibit an increased signal intensity in negative-ion mode when compared to non-phosphorylated analogs whereby a comparative use of positive-ion mode and negative-ion mode in MALDI MS and ESI MS can reveal phosphopeptide candidates [19,57,106,167]. However, as results in Fig. 5.8 indicate, the improved relative signal-to-noise ratio obtained in negative-ion mode MALDI is, however, penalized by an overall lower sensitivity. The comparison of positive-and negative-ion spectra (relative intensity ([M
H]
)/relative intensity ([M+H]+)) can instead be used to identify phosphopeptides in the peptide mass maps which then can be separated and analyzed further. Phosphopeptide enrichment techniques such as immobilized metal affinity chromatography (IMAC) prior to MS analysis have been very successful for studies of individual phosphoproteins as well as for phosphoproteomic investigations. IMAC reduces the proportion of easily ionizable peptide components that supposedly cause suppression of phosphopeptides [8,169,171,194,232]. Chemical derivatization strategies to eliminate and convert phosphoamino acid residues into more stable and tractable species have also been used extensively (reviewed by MacLaclin and Chait [34,104,156,162,176,205,230,244]). The chemical modification of serine and threonine phosphorylation sites in phosphoproteins has been facilitated by b-elimination in alkaline conditions followed by Michael addition reaction with nucleophilic agents such as ethanethiol [103,105,205], ethanedithiol [74,177] and dithiothreitol [7]. The advantages of this method include increased hydrophobicity and hence retention time of the modified peptides, a 292

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Fig. 5.7. Increased phosphopeptide response in MALDI MS analysis using phosphoric acid as matrix additive (phosphopeptides are indicated with asterisks). (A) Tryptic MALDI peptide mass map of bovine PP3 prepared by the dried droplet method using 2,5-DHB in 50% acetonitrile/2.5% formic acid. (B) By including o-phosphoric acid (2,5-DHB in 50% acetonitrile/1% o-phosphoric acid (PA)) as matrix additive increased the response of phosphopeptides.

facilitation of positive-ion production, and an increased susceptibility to tryptic digestion as a result of conversion of negatively charged phosphorylated residues to neutral residues [104]. This approach has proven especially successful for analysis of multi-phosphorylated 293

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Intensity

x 12.0

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*

*

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* *

*

* 1000

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2500 m/z

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Fig. 5.8. Comparison of positive- and negative-ion mode MALDI MS for phosphopeptide detection by dried droplet preparation of trypic peptide digest (phosphopeptides are indicated by asterisks). An improved relative signal-tonoise ratio in MALDI negative-ion mode (top panel) is observed, but also an overall lower sensitivity.

peptide, where Resing et al. as well as Jaffe and co-workers have published studies in which large numbers of phosphorylations sites were identified [103,105,205]. The pitfalls of this approach is that O-linked sugars undergo the same b-elimination reaction potentially leading to O-glycosylation sites being incorrectly assigned as phosphorylation sites and the chemistry do requires significant effort to perform properly [156,245]. In situ Liquid/Liquid Extraction (LLE)-MALDI-MS has recently been demonstrated for enhanced separation and structural analysis of posttranslationally modified peptides [123]. Recently, nanoscale graphite columns in a multi-tiered approach have proved very useful for the recovery of small and hydrophilic phosphopeptides prior to MS analysis [49,135–137]. 5.4.3

Detection of phosphorylated peptides in peptide mass maps

Phosphopeptide mass mapping is the simplest way to identify phosphopeptides by MS [19,57,106,167]. Phosphopeptides can be identified in ESI and reflectron-MALDI mass spectra by either the specific phosphoamino 294

The phosphoproteome story

acid masses or the detection of prompt and/or metastable decay product ions diagnostic for phosphopeptides [10,165,248]. Phosphopeptide candidates may be identified comparing the list of experimentally determined peptides with the theoretical list of proteolytic fragments allowing phosphoamino acid residues having a mass increase of 80 Da corresponding to the mass of H3PO4. Serine and threonine phosphorylated peptides display the two fragment ions [MH
H3PO4]+ (98 Da) and [MH
HPO3]+ (80 Da) due to gas-phase b-elimination of phosphoric acid from phosphoserine and phosphothreonine and prompt fragmentation of phosphate in the source region of MALDI-TOF instruments, respectively. An apparent mass difference (86Th) between the precursor ion and the metastable ion is observed in the mass spectra in Fig. 5.9, actually corresponds to a 98 Da difference (loss of phosphoric acid). This discrepancy is due to the characteristics of the reflector TOF analyzer. Metastable ions that have a minor loss of kinetic energy to the neutral fragment during decomposition are not optimally focused onto the second detector and therefore, they do not follow the same calibration curve as the intact precursor ion which is calibrated for ions with the full accelerating energy. The decay products can be observed to have a decreased resolution. In addition to serine and threonine phosphorylated-amino acid residues, post-acceleration PTM-specific metastable molecular ions allow specific detection of carboxamidomethylated cysteines, oxidized methionines and glycosylated amino acid residues [252]. The intensity of phosphospecific satellite ions are partly due to MALDI matrix used. The two MALDI matrices 2,5-DHB and 4-HCCA are commonly used due to the high sensitivity for peptide analysis [10]. The ‘‘cold’’ 2,5-DHB matrix imparts only little internal energy in the precursor molecular ions when using threshold laser for the generation of gas-phase ions in MALDI, whereas the ‘‘hot’’ 4-HCCA MALDI matrix shows an increased level of metastable fragmentation due to the lower initial velocity of gas-phase ions generated by this matrix. In the linear mode, a loss of kinetic energy of the precursor ion will not alter the time of flight, thereby the metastable ions strikes the detector at the same time as the precursor ion, thus all detected at the same apparent mass. If the suspected metastable fragment ion [MH-98]+ disappears when the mass spectrum is recorded in the linear mode (Fig. 5.9, right panel), one can be reasonably certain that the peptide is phosphorylated. The disappearance of the metastable fragment ion occurs because the fragment ion only has a fraction of the full accelerating voltage applied. 295

A. Stensballe and R. J. Simpson Matrix: a-4HCCA (hot matrix)

83.037 Da

0 10

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1131.70 0 167.032 Da

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1120

1171.69

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Fig. 5.9. Identification of phosphopeptides by peptide mass mapping using reflectron-MALDI TOF MS and linear-MALDI TOF MS. (Left) Structures and monoisotopic masses of phosphoserine and phosphothreonine. The b-elimination of phosphoric acid converts from dehydroalanine and dehydroamino-2butyric acid, respectively. Phosphotyrosine cannot undergo b-elimination due to an aromatic ring. (Right) Comparison of MALDI mass spectra obtained in linear TOF mode and reflector TOF mode. Metastable decomposition products diagnostic of gas-phase b-elimination of phosphoric acid from the two phosphopeptides can be observed (indicated with asterisks). Phosphopeptide candidates are labeled with numbers and metastable decomposition products. The apparent mass difference (86Th) between the precursor ion and the metastable ion observed in the mass spectra, actually corresponds to a 98 Da difference (loss of phosphoric acid). Adapted from Stensballe et al. [232].

Validation of candidate phosphopeptide assignments can be attained by enzymatic dephosphorylation of a phosphopeptide sample either in solution [141,232,254,258,260] or in situ on a MALDI probe by protein phosphatases will decrease the mass by 80 Da per phosphate group removed [138]. Detection of phosphorylated species can be revealed by comparing mass spectra or base peak chromatograms obtained from LC MS recorded before and after phosphatase treatment, and looking for peaks that disappear from the treated sample as well as peaks that appear or increase in intensity, it is possible to identify candidate phosphopeptides. 5.4.4

Phosphorylation-specific precursor ion discovery by MS

The detailed knowledge of gas-phase fragmentation processes during mass spectrometric analysis has been used to detect modified peptides 296

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through specific phosphospecific fragment ions. Despite that ESI is a gentle ionization technique, phosphopeptides can easily be induced to loose the phospho moiety during MS experiments. This is especially the case for serine- and threonine-phosphorylated peptides, which readily undergo b-elimination upon low-energy CID [42]. A range of MS-based approaches for studying PTMs take advantage of characteristic fragment ions or neutral losses for a given type of modification. Thus, labile phosphate side chains of phosphopeptides or formation of immonium ions can be used for a specific detection in the neutral loss scan mode or parent ion scan mode (Fig. 5.10). These MS scan modes have previously been demonstrated to selectively detect not only phosphorylation [11,42,167,168,226], but also modifications such as glycosylation [42], acetylation [29], acylation [18], sulfation [18], bromotryptophan [229], hydroxyproline [229] and arginine dimethylation [202]. Strategies for detection of phosphopeptides include monitoring the characteristic losses of the phosphate moiety in phosphoserine and phosphothreonine and the phosphotyrosine immonium ion. In negativeion mode phosphorylation-specific marker ions can be utilized in combination with online or off-line chromatographic by parent ion scanning (m/z 79 Da; PO
3 ) and skimmer collision-induced dissociation (sCID) (m/z
63 Da; PO
2 or m/z ¼ 97 Da; H2PO4 ) under alkaline conditions. Annan et al. [11] have described an approach that utilizes two orthogonal MS scanning techniques, both of which are based on the production of

Fig. 5.10. (A) Triple–quadrupole in parent ion scanning; (B) Triple–quadrupole in neutral loss scan mode, where mass X-98 corresponds to loss of phosphoric acid by gas-phase b-elimination from pS and pT. 297

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phosphopeptide-specific marker ions at m/z
63 and/or
79 in the negative-ion mode. Detection of phosphopeptide candidates by precursor-ion scan methods allowed selective detection and identification of phosphopeptides in complex proteolytic digests by LC-MS/MS and nanoelectrospray MS/MS. Beck et al. [19] have investigated the use of LC-MS under alkaline conditions together with selective detection of phosphopeptide ions by phosphorylation-specific marker ions (m/z ¼ 79 and 97) generated by sCID in the negative-ion mode. This approach later allowed the identification of novel phosphorylation sites in IRS1 [20]. Flora and Muddiman [65] have introduced a method for the selective detection of phosphopeptides in Fourier Transform Ion Cyclotron Resonance Mass Spectrometry (FTICR MS), which is based on the loss of the phospho moiety. Performing ESI FTICR experiments in negative-ion mode mass spectra of the peptide mixture was before and after lowintensity infrared irradiation. Although this level of irradiation did not induce cleavage of the peptide backbone, it was sufficient to induce the loss of the phospho moiety. Comparing the two spectra and searching for peak pairs differing by 98 Da allowed the identification of phosphorylated species in the mixture. Under positive-ion conditions both serine- and threonine-phosphorylated peptides undergo loss of phosphoric acid by b-elimination (m/z 98 Da; H3PO4) and 80 Da (HPO3) by prompt fragmentation from the molecular ion, whereas tyrosine-phosphorylated peptides preferentially lose 80 Da. Phosphotyrosine-containing peptides can also decompose during MS analysis to lose HPO3 to generate a fragment ion [MH-80]+. Since phosphotyrosine is much more stable than either phosphoserine or phosphothreonine, this ion is normally not very abundant. The metastable neutral loss of phosphoric acid, typical of phosphoserine and phosphothreonine, is not favored in the case of a phosphotyrosine because it would require cleaving a bond adjacent to an aromatic ring, leaving a radical on an aromatic ring. However, if [MH-98]+ ions are observed in mass spectra of phosphotyrosinecontaining peptides, the loss is likely due to the result of sequential losses of HPO3 and H2O. It is therefore generally possible to distinguish tyrosine phosphorylation from serine or threonine phosphorylation, at least for singly phosphorylated peptides, based on the type of fragment ions present. The quadrupole mass analyzer, especially when used in a triple–quadrupole mass spectrometer, can efficiently be employed for detection and analysis of phosphopeptides. First, in the precursor ion 298

The phosphoproteome story

scanning mode the Q3 mass filter is set to detect a single mass only. The deprotonated PO3
ion (m/z ¼ 79) is the best diagnostic ion for specifically detecting phosphopeptides, since it is rarely observed as a product of fragmentation of non-phosphopeptides (Fig. 5.10A). The precursor ion scan technique is well established and very useful for the characterization of unseparated peptide mixtures at the femtomole level [167]. The off-line approach allows detection of phosphopeptides in the negative-ion mode at alkaline pH to obtain maximum signal for the m/z ¼ 79 ion, and subsequently performing sequencing in positiveion mode after acidification of the sample in the nanoelectrospray needle. In the neutral-scan mode the Q1 and Q3 mass filters are scanning in parallel but with a specific offset by the difference in mass between the phosphopeptides and de-phosphopeptides due to collisions in the Q2 collision cell (Fig. 5.10B). Upon neutral loss of phosphoric acid by b-elimination, the observed mass difference in m/z between Q1 and Q3 will be 98 and 49 for the singly and doubly charged precursor ions, respectively. The tyrosine-phosphorylated peptides must instead be detected by the neutral loss of a phosphate group HPO3 (i.e., 80 for 1+ and 40 for 2+). The limitations for neutral loss scanning have a lower sensitivity than the precursor ion scanning method and significant ‘‘cross-reactivity’’ with signals generated by other side chains (e.g., b1 ions of Pro and Val with the masses of 97 and 99 amu, respectively). The phosphotyrosine immonium ion (Im[pY]; m/z ¼ 216.043), being mass-deficient, has enabled precursor ion scanning with high mass accuracy as well as mass resolution for specific detection using highperformance QTOF-based mass spectrometers [226,228]. The use of traditional triple-quadrupole instruments for precursor ion scanning is limited due to the low mass resolution of this type of instrument [228]. In practice, for daughter ion scanning of complex mixtures, i.e. a tryptic digest of a large phosphoprotein or protein complex, triple–quadrupole mass spectrometers is very often unsuccessful due to overlapping isotope clusters prohibiting sequencing of phosphorylated peptides (B. Ku ¨ ster, personal communication). Instead, the high resolving power of hybrid-quadrupole TOF mass spectrometers enables the selective detection of phosphotyrosine immonium ions without interference from other peptide fragments of the same nominal mass (see Fig. 5.11). For serine –– and threonine-phosphorylated peptide, a similar strategy has been devised that uses b-elimination/Michael addition of a nucleophile agent. Based on this chemistry, a new functional group is introduced at the original site of phosphorylation, which gives rise to a 299

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MS/MS of TNLSEQ (pY) ADVYR: QqQ

QqTOF

214.0

216.0

218.0 m/z

QqTOF

216.05 216.10

215.7

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Fig. 5.11. Mass resolution of triple–quadrupole (QqQ) and quadrupole-timeof-flight (QqTOF) type mass spectrometer for detection of Im[pY]. Courtesy of H. Steen [226].

labile dimethylamine-containing sulfenic acid derivative. An abundant immonium ion with a unique m/z value then enables the detection of the former phosphorylated species within peptide mixtures by precursor ion scanning in positive-ion mode [230]. Although precursor scanning enables highly specific detection of modified phosphotyrosine-modified peptides, the duration of MS experiments (many seconds) render this approach impossible in combination with online liquid chromatography, e.g., LC-MS/MS (H. Steen, personal communication). Instead, software based Parent Ion Discovery (PID) of phosphopeptides in combination with online liquid chromatography experiments have been developed for QTOF mass spectrometers [17]. Ideally, online PID during nanoflow LC-MS/MS of complex peptide mixtures allows selection of the subset of modified peptides for fragmentation. This would, in principle, allow selection of phosphorylated precursor ions among co-eluting precursor ions of higher abundance, thus allowing improvement of the ion statistics for 300

The phosphoproteome story

the selected precursor ion by prolonged MS/MS acquiring. Figure 5.12 shows the analysis of a tryptic peptide mixture by LC-MS/MS precursor ion discovery. Real-time investigation of consecutive wideband MS scans indeed allowed specific detection of low-intensity phosphorylated peptides in preference of far more abundant unmodified co-eluting peptides (A. Stensballe, unpublished data). 5.4.5

Phosphopeptide sequencing by mass spectrometry

Phosphopeptide sequencing can be performed by both MALDI- and ESI-based instruments using PSD, tandem MS or MS3 approaches. Sequencing of phosphopeptides has been performed by ESI triplequadrupole instruments [248,249], ion-trap instruments [104] or ESI hybrid-quadrupole/TOF instruments [232]. Currently, the primary tool for gas-phase sequencing of phosphopeptides is low-energy CID MS/MS due to the efficient fragmentation processes of multiply protonated ions. Fragmentation of singly charged peptide ions produced by MALDI is also achievable using MALDI tandem mass spectrometers, such as QTOF, TOF–TOF and ion trap instruments [133,143,157]. The complexity of the peptide fragmentation patterns due to a significant sidechain losses of HPO3 and H3PO4, the low yield of fragment ions and the limited mass accuracy and sensitivity have made PSD-based protein identification by MALDI-reTOF MS and localization of PTMs a challenge [76,93]. Protein phosphorylation, N- and O-glycosylation, sulfation and g-carboxylation are rapidly lost upon vibrational excitation by lowenergy CID [122,234]. Especially characterization of phosphopeptides with multiple phosphates and many potential phosphorylation sites also remains troublesome even by current MS-based methods. The site-specific identification of phosphorylated residues is often limited by the fact that multiply phosphorylated peptides are likely to produce very complex CID spectra, or ionize poorly in the process of ionization. Figure 5.13 illustrates the differences in fragmentation by low energy CID spectra of small serine/threonine phosphorylated peptides using ESI. These phosphopeptides generate ion series (mainly b-/y-) that reflects the loss of phosphoric acid (M-98 Da) in addition to cleavage of the backbone peptide linkages and intensive loss of water (M-18 Da). MALDI-MS/MS can be used to identify a phosphoprotein either by peptide mass mapping of the protein digest or from tandem mass spectra acquired by low-energy CID of individual singly charged phosphopeptide precursor ions 301

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A 100

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The phosphoproteome story

Fig. 5.13. Phosphopeptide sequencing by QTOF tandem mass spectrometry. (Upper panel) Electrospray QTOF MS/MS spectrum of the unmodified peptide (IPFDGESAVSIALK, m/z 723.9). The spectrum exhibits a near-complete y-ion fragment ion series. (Lower panel) Electrospray Q-TOF MS/MS spectrum of the doubly protonated phosphopeptide ion (IPFDGE[pS]AVSIALK, m/z 763.9) from B subtilis PrkC protein. C-terminal peptide ion series is indicated (y and y* ions). This phosphopeptide generates a y8* – y13* ion series that reflects the loss of phosphoric acid from the y-ions and intensive loss of water [MH
18 Da]+ (compare with upper panel). Fig. 5.12. Analysis of phosphopeptide mixtures by LC-MS/MS precursor ion discovery triggered by neutral loss of phosphoric acid. (A) Base peak intensity (BPI) chromatograms for the lower and higher collision energy wideband MS survey scans and MS/MS switching events. (B) An expanded view of the range m/z 650–710 illustrating the neutral loss of H3PO4 (48.989 Da) from a doubly charged phosphopeptide molecular ion. (C) Tandem MS identifies the phosphopeptide and site of phosphorylation. 303

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[21,204,219]. The sequencing capability makes protein identification more specific than MALDI peptide mass mapping and enable strategies that takes advantage of the complementary nature of MALDI-MS/MS and ESI-MS/MS thereby increasing proteome coverage [1,27]. Often small peptides, especially non-tryptic, end up singly charged using ESI which is less favorable for low-energy CID and often not sequenced by data-dependent acquisition. MALDI tandem mass spectra of phosphoserine and phosphothreonine containing peptides often display significant fragment ion signals which originate from partial or complete neutral loss of phosphoric acid as well as loss of water (-18 Da; H2O) and ammonia (-17 Da; NH3) molecules [21]. The overall complexity of MS/MS spectra increases dramatically with increasing peptide length and number of phosphorylated residues [21,147,171]. One significant advantage of using MALDI Q-TOF MS/MS for amino acid sequencing of multiply phosphorylated, singly protonated peptide ions is that the resulting tandem mass spectra are relatively simple as compared to ESI MS/MS data obtained from the corresponding multiply charged phosphopeptide ions as presented in Fig. 5.14. The ESI tandem mass spectrum of a quadruply phosphorylated peptide was exceedingly complex due to the presence of multiple charge states (z ¼ 1 to 4) of y, y-NH3 and y-H2O ion series as well as incomplete neutral loss of phosphoric acid. In contrast, MALDI Q-TOF MS/MS sequencing of the corresponding singly charged phosphopeptide ion produced a much simpler and easy to interpret tandem mass spectrum to allow exact localization of four closely spaced phosphoamino acid residues. To compensate for the less efficient CID process of singly charged precursor ions, often higher amounts of analyte must be used in order to generate sufficient ion signals by MALDI MS for subsequent acquisition of high-quality MS/MS spectra of

Fig. 5.14. Phosphopeptide sequencing of large multiphosphorylated peptide by ESI and MALDI tandem mass spectrometry. (Upper panel) MALDI-QTOF MS/MS spectra of singly protonated phosphopeptide ion 150 VTDFGIATALSS[pT][pT]I[pT]H[pT]NSVLGSVHYLSPEQAR
183 derived from PrkC protein from B. subtilis. (Lower panel) Nanoelectrospray MS/MS spectrum of quadruply protonated phosphopeptide ion. The quadruply phosphorylated peptide displayed a near complete series of y-ions as well as extensive ion signals due losses of H2O (
18 Da). Gas-phase b-elimination of phosphoric acid from phosphothreonine (181 Da) to dehydroamino-2-butryric acid (83 Da) were evident in the y-ion series confirming all four modified residues.

304

The phosphoproteome story

phosphorylated peptides for unambiguous assignment of phosphoamino acid residues. To overcome the challenges of interpreting phosphopeptide mass spectra other fragmentation mechanisms more suited for sequencing 100

VTDFGIATALSSbbIbHbNSVLGSVHYLSPEQAR b = 2-aminodehydrobutyric acid MOWSE score 170

∆

y15 y9

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labile modifications have been demonstrated. FTMS employing ECD enabled efficient fragmentation of large peptides and small proteins with labile modifications [45,46,122,221,234,264]. In this non-ergodic fragmentation mechanism, polypeptide backbone cleavage is favored, leaving the more labile PTMs intact. ECD leads to extensive fragmentation of the polypeptide backbone generating c- and z type ions providing a good amino acid sequence coverage for the fragmented peptides (Fig. 5.15) [119]. In particular multi-phosphorylated peptides are otherwise difficult to sequence by MS/MS using low- or high-energy CID because of the lability of the phosphate groups resulting in complex fragmentation behavior. Recently, Syka and co-workers [236,237] have introduced a related fragmentation mechanism termed electron transfer dissociation (ETD). In contrast to ECD, automated acquisition of single-scan ETD tandem mass spectra from phosphopeptides separated by nanoflow HPLC (nHPLC) in a linear ion trap has been demonstrated. 5.4.6 Nanoscale chromatography for affinity enrichment and separation of phosphopeptide mixtures

Integration of nanoscale multidimensional chromatographic techniques interfaced to MS has shown to improve the sensitivity and selectivity for comprehensive phosphoprotein analysis [11,54,63,171]. Indeed, the use of any form of upfront chromatographic separation that either simplifies peptide mixtures or isolates phosphopeptides prior to MS analysis can be advantageous [33]. 5.4.6.1 Nanoscale affinity purification of phosphopeptides Hitherto, one of the most successful approaches for enrichment of phosphopeptides has been IMAC. Affinity purification of phosphoproteins and phosphopeptides by IMAC (also called metal-chelate affinity chromatography) was originally established by the observation of a very high affinity of phosphate groups toward a range of multivalent transition metal ions [8,88,193]. In contrast to multiple other phosphopeptide enrichment strategies, IMAC have now proven to efficiently enrich phosphothreonine, phosphoserine and phosphotyrosine containing peptides and do not necessarily require chemical derivation [54,171,176,210,262]. Enrichment by IMAC has primarily been performed off-line in nanocolumns or capillary columns [63,232] or by batch incubation 306

The phosphoproteome story a

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Fig. 5.15. ECD analysis of the quadruply phosphorylated peptide RELEELNVPGEIVE(pS)L(pS)(pS)(pS)EESITRINK (Mr ¼ 3476.48) isolated by Fe(III)-IMAC. The quadruply protonated phosphopeptide at m/z 870 was isolated and fragmented by ECD producing mainly c- and z type ions. Adapted from Stensballe et al. [234].

[54,171]. The selectivity and recovery of phosphopeptides by IMAC depend on several factors, including the choice of chelating resin or metal ion used, the sample loading conditions, the washing conditions and the elution conditions for recovery of phosphopeptides. Metalchelating stationary phases such as imino-diacetic acid (IDA) or nitrilotriacetic acid (NTA) coupled to a support like Sepharose, agarose or macroporous silica are the most commonly utilized resins, however, the latter have shown the lowest propensity for unspecific binding of nonphosphorylated peptides [63,84,171,194,232]. The trivalent metal ions Fe3+ (Fe(III)) and Ga3+ (Ga(III)) have so far shown the highest specificity for phosphopeptides [9,84,171,194]. Loading conditions for all types of IMAC resin were acidic conditions (2.5opHo3.5), resulting in the protonation of the majority of glutamic and aspartic acid. Also a 307

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range of washing steps and equilibration steps as well as elution solvents have been investigated to reduce the level of unspecific binding peptides to the IMAC resin. A low level of non-phosphopeptides have been observed to be present in the IMAC eluate using both MALDI-MS and ESI-MS for phosphopeptide mapping [24,194,263]. The retained non-phosphopeptides have previously been shown to be rich in acidic and/or hydrophobic residues [84,166,169,194]. However, recent investigations by Nuhse et al. [171] on the properties determined that mainly high abundance peptides are the primary source of non-phosphorylated contaminants in IMAC preparations. In order to reduce binding of acidic peptides, both loading and washing steps were performed under acidic conditions (0.1 M acetic acid), thereby preventing these peptides from binding to the immobilized metal. Two factors are reported in the literature to have significant influence upon the level of non-phosphopeptides in the IMAC eluate. First, a high degree of nonspecific binding to the column support material was generally observed [171]. This binding have, to a great extent, been reduced by either including a percentage of acetonitrile and/or sodium chloride during the loading of the sample or by including a washing step containing a percentage of acetonitrile (e.g. 25%) after loading of the sample. Acetonitrile is known to break up interactions between hydrophobic peptides and the polymeric IMAC resin. Alternatively, peptides mixtures have been carboxy-methylated prior to loading thereby eliminating non-specific binding by acidic peptides; however, recent data do indicate that the carboxy-methylation process also results in incomplete methylation of the aspartic, glutamic and C-termini of tryptic peptides, thereby increasing the complexity of the digest and complicating the identification of phosphopeptides and specific phosphorylated residues [56,63,84]. Observations by Cao and Stults [39] additionally suggest, that sodium adduction to phosphopeptides influences negatively on the binding to the Fe(III)-IMAC column. Also ammonia, acetate and bicarbonate have been reported to be potential metal-binding ligands that may reduce the efficiency of phosphopeptide recovery [194]. Multiple approaches have been used for elution and sample preparation of the retained phosphopeptides from the IMAC resin. (I) Alkaline buffers including 0.5–2% ammonium hydroxide and 0.1% ammonium acetate adjusted to pH 8.7–9.5 by ammonium hydroxide [39,40,66,140,173,241]. (II) Competitive displacement by salts like 50–250 mM sodium phosphate (Na2HPO4, pH 8.4/9.0), followed by a desalting step or LC-MS/MS analysis; 0.1%/50–100 mM ammonium 308

The phosphoproteome story

dihydrogen phosphate (NH4H2PO4, pH 4.2–9.6); 50–200 mM K2HPO4 or 50 mM triethylammonium hydrogen carbonate (pH 8.0) [63,67,83,142,166,169,194,196,207,210,212,215]. (III) Removal of the chelated-metal ion by 100 mM EDTA [63]. (IV) Phosphoric acid (0.2%) [129,130]. (V) ‘‘Direct MALDI MS analysis’’ of phosphopeptides affinity bound to immobilized metal ion agarose and direct elution by MALDI matrices, in particular 2,5-DHB [35,83,199,204]. In our experience, the most efficient eluents remains high pH elution or MALDI matrix combined with phosphoric acid. The latter eluent enables immediate sample preparation for MALDI MS/MS analysis without further sample handling and potential risk of phosphopeptide loss by desalting step otherwise necessary [90,233]. A significantly enhanced detection of singly and multiply phosphorylated peptides analyzed by MALDI MS can be accomplished using a one-step elution of nanoscale IMAC (Fig. 5.16). Recently, titanium oxide (TiO2) has been introduced as a new affinity media with high potential in the field of phosphoproteomics [101,192]. Pinkse and co-workers demonstrated the selective enrichment and characterization of phosphopeptides from both simple and complex mixtures. 5.4.6.2 Multidimensional chromatographic techniques for analysis of phosphopeptide mixtures The integration of single- or multidimensional chromatography to MS provides powerful methodologies for analysis of simple as well as very complex phosphopeptide mixtures. Multiple analysis strategies have investigated combinations of chromatographic approaches prior to offline or online analysis by MS for reduction of suppression effects or/and to obtain a higher protein sequence coverage (Fig. 5.17). Desalting and concentration of simple protein digests by nanoscale reversed-phase chromatography is a common analytical step prior to phosphopeptide analysis by MS [59,70]. Although sample desalting and up-concentration by nanoscale reverse-phase columns using off-line nanoscale systems show excellent performance, there can be loss of phosphopeptides in the desalting step due to the hydrophilic nature of the phosphopeptides. Previous reports have suggested that short and/or hydrophilic peptides are poorly retained by reversed-phase columns [38,173]. To recover most peptides and thereby improving protein sequence coverage, a multi-tiered approach using increasingly more hydrophobic chromatographic materials can be applied [135]. The use of graphite columns 309

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4000

Fig. 5.16. Improved detection of phosphopeptides by combined IMAC and optimized MALDI matrix. The application of DHB and o-phosphoric acid as eluant in IMAC-enabled recovery of both singly and multiply phosphorylated species in a tryptic hydrolysate of PrkC from B. subtilis. (A) MALDI-QTOF MS analysis of 0.5 pmol trypic digest of PrkC phosphoprotein. The analyte/matrix deposit was prepared by the dried droplet method using 2,5-DHB in 50% acetonitrile/ 1% o-phosphoric acid. (B) MALDI MS analysis of IMAC-enriched phosphopeptide fraction from 0.5 pmol trypic digest of PrkC eluted directly on the MALDI probe by 2,5-DHB in 50% acetonitrile/1% o-phosphoric acid.

for retaining small hydrophilic peptides can successfully capture otherwise non-retained phosphopeptides [135,137]. Splitting up the initial sample into multiple fractions will reduce signal-suppression effects on phosphopeptides caused by the presence of non-phosphopeptides during 310

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Fig. 5.17. Approaches for phosphopeptide analysis of peptide mixtures of increasing complexity using combinations of chromatographic separations as a frontend for mass spectrometry.

the MALDI process. Further subfractionation can be accomplished by a multi-tiered scheme for phosphopeptide analysis using affinity enrichment by IMAC, optimized MALDI matrix for phosphopeptides and increasingly more hydrophobic chromatographic materials as illustrated in Fig. 5.18 (A. Stensballe, unpublished). First, the fraction of phosphopeptides having affinity for IMAC are recovered. The recovery phosphopeptide species by IMAC in both small- and large-scale experiments can be incomplete, especially for non-acidic phosphopeptides [63,171]. Therefore, the flow from the IMAC is collected by reversed-phase media (C18; Poros R2/Oligo R3) in a nanocolumn. In the case of both known or novel proteins, this peptide fraction (i.e. non-retained peptides) can be used for protein identification, determination of sequence coverage or subjected to MS/MS analysis and used for searching in both protein and DNA sequence databases as well as subjected for further investigation of posttranslationally modified peptides. Especially, small hydrophilic peptides may not be adsorbed even by the highly hydrophobic Poros Oligo R3 chromatographic material, but however, will be trapped by a subsequent graphite column. All fractions are eluted directly onto the MALDI probe using 2,5-DHB in 50% acetonitrile/1% O-phosphoric acid for MALDI-MS/ MS analysis. The mass spectrometric data may be evaluated by a hypothesis-driven approach after initial MS analysis and phosphopeptide 311

312 A. Stensballe and R. J. Simpson

Fig. 5.18. Analysis of human RFX5 [P48382] by a multi-tiered scheme for phosphopeptide analysis using the optimized affinity enrichment and increasingly more hydrophobic chromatographic materials thereby increasing phosphopeptide recovery and protein sequence coverage. Following in-gel trypsination an aliquot of the digest (5%) was separated into three fractions according to the scheme, 1, Fe(III)-IMAC; 2, OligoR3; and 3, graphite, where all fractions were eluted with MALDI matrix. Incorporation of radioactive phosphate enabled visualization by autoradiography and localization of phosphopeptide-containing fractions. Analysis of each fraction by MALDI-MS allows off-line investigation of the peptide mass maps for the presence of phosphopeptide candidates. Phosphopeptides can be sequenced by MALDI tandem MS to identify sites of phosphorylation.

The phosphoproteome story

candidates subsequently sequenced by tandem MS or verified by phosphatase assay [92]. The ionization method of the mass spectrometer determines the interfacing to the mass spectrometer. ESI-based mass spectrometers are ideally suited for direct coupling to liquid chromatographs due to the analyte being on liquid phase. In contrast to ESI, MALDI necessitates the analyte to be imbedded in microcrystals (matrix) prior to ionization. Hence, MALDI MS is decoupled from the actual data acquisition. Offline acquision allows, in principle, for analysis of each phosphopeptide sample preparation where 2,5-DHB matrix remains stable for weeks without significant reduction in performance unlimited time. Recent investigations have demonstrated that an improved proteome coverage can be obtained using a combination of ESI and MALDI for LC-MS/MS studies [27]. Improved sequence coverage of the analyzed samples has accomplished by iterative reanalysis using permanent exclusion lists or retention-time-dependent exclusion lists, and multi-enzyme digestion for generation of overlapping sequences [54,215]. Nanoscale capillary liquid chromatography (capLC) coupled to ESI or MALDI MS/MS is a powerful approach for automatic analysis of complex phosphopeptide mixtures. The reduction in column volume and flow rate in the low nanorange effectively results in a high gain in sensitivity [48,159]. The hydrophilic character of phosphopeptides affects their chromatographic properties. Short and/or hydrophilic phosphopeptides are observed to elute very early or in the void volume when using reversed-phase chromatography for phosphopeptide separation [38,135,137]. To lower the loss of phosphopeptides in the void volume during reversed-phase chromatography the addition of the hydrophobic ion-paring agent heptafluorobutyric acid (HFBA) as counterion in the mobile phases delay the elution time of phosphopeptides [178]. Incorporation of radioactive phosphate enable visualization by autoradiography and allow localization of phosphopeptide-containing fractions from off-line chromatographic separation (Fig. 5.19). An optimized sample preparation using the combination of phosphoric acid and 2,5-DHB for sample preparation has been shown to improve the performance of off-line LC-MALDI MS/MS of phosphopeptides [124]. The relative acidity of phosphopeptides has been exploited by Gigy and co-workers for partial enrichment of phosphopeptides by ion-exchange chromatography. They successfully applied strong cation exchange (SCX) for the identification of 500 phosphorylation sites in the mammalian brain 313

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Fig. 5.19. Off-line chromatographic separation with MALDI-MS and tandemMS can be a powerful approach to improve peptide mass mapping of phosphoproteins. Analysis of four PKA kinase substrates labelled with [g-32P]ATP in the presence or absence of the kinase were accomplished by nanoflow LCMALDI-MS and MS/MS. Following in-gel trypsination the protein digests were separated using nanoflow HPLC coupled to a liquid sampling robot and the eluate spotted directly in 30 s fractions onto a 384-well MALDI probe. Visualization by autoradiography enabled localization of phosphopeptide-containing fractions. These fractions were analyzed by MALDI-MS, analyzed in silico for the presence of phosphopeptide candidates, which were sequenced by MALDI tandem MS (A. Stensballe, unpublished results;[92]).

[15]. Highly complex peptide mixtures can be separated by strong anion exchange (SAX) chromatography into multiple fractions. A multi-dimensional fractionation scheme for efficient separation and isolation of phosphopeptides from very complex peptide mixtures was conceived based on the technical difficulties of analyzing complex phosphopeptide mixtures (see Fig. 5.20) [171]. Special loading conditions including organic solvent in SAX fractionation makes this approach less suitable for an online approach [188]. SAX fraction was individually incubated in a batch format with IMAC resin to isolate phosphopeptides followed by analysis by LC-MALDI MS or LC-ESI-MS/MS [171,223]. 314

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Fig. 5.20. Multidimensional SAX/IMAC/RP-LC-MS/MS analysis of complex protein mixtures.

5.4.6.3 Scalable approaches for automated phosphopeptide analysis Several dedicated systems have been developed for the automated enrichment and analysis of phosphopeptides combining affinity chromatography, nanoflow liquid chromatography and MS. A commercial alternative for automatic sample cleanup and IMAC enrichment of phosphopeptides in an integrated, highly parallel and disposable format has been developed by Gyros AB (Uppsala, Sweden) [89,90,115]. The Gyrolab MALDI IMAC enables analysis of 48 protein digests prepared in duplicate (2  48) in a micro fluidic Lab-on-chip microlaboratory. The phosphorylated peptides are concentrated, purified and crystallized directly onto MALDI target areas on the CD using micro fluidic structures for solvent pathways and low-nanoliter chromatographic columns. Each duplicate sample is subjected to the same process, but with the addition of an enzymatic step that removes any phosphate groups before crystallization. The CD is transferred to MALDI MS/MS for analysis. Phosphorylated peptides are detected by comparing mass spectra from phosphorylated and dephosphorylated samples. A complementary integrated system for the automated enrichment and analysis of phosphopeptides by IMAC/nano-LC/ESI-MS has been introduced by Ficarro and co-workers [64]. This system utilizes two independently controlled HPLC pumps, an autosampler and microvalves to perform sequential analysis of IMAC-retained fraction and non-phosphorylated peptide fraction into an electrospray mass spectrometer. Their use of robust IMAC and reversed-phase HPLC 315

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columns with integrated ESI emitter tips enabled the reproducible detection and identification of low-femtomole quantities of phosphopeptides. Pinkse et al. [192] introduced a similar automated method for the selective enrichment of phosphopeptides from complex mixtures using a two-dimensional column switching setup, with titanium oxide-based solid-phase material as the first dimension and reversed-phase material as the second dimension.

5.5

MASS SPECTROMETRY-DRIVEN STRATEGIES FOR PHOSPHOPROTEOMICS

As significant improvements in MS and bioinformatics have facilitated analysis of post translationally modified proteins, large-scale identification and characterization of phosphoproteins have become increasingly more common. Phosphoproteome studies aim to identify the constituents of the proteome that become phosphorylated, determine the exact localization of the modified residues and ideally quantify the regulatory changes in protein phosphorylation between cellular states. More than 10 successful large-scale phosphoprotein studies aiming at mapping phosphorylation sites in large scale have been published so far. These studies include the analysis of yeast whole-cell protein lysate [63]; human T-cell phosphotyrosome [210]; outer cellular membrane subproteome from Arabidopsis thaliana [171,172]; shotgun identification of protein modifications from protein complexes and membranes [145,253], Pheromone signaling pathway in yeast [78]; the mouse synapse phosphoproteome [54]; the global, time-dependent analysis of the EGFR phosphotyrosine proteome [25]; MFC7 phosphoproteins [198] and the developing mouse brain [15]. The low stoichiometry, heterogeneity and low abundance of phosphoproteins led most investigators to include one or several enrichment steps for phosphoproteins and phosphopeptides prior to MS analysis, thereby efficiently increasing the relative abundance of phosphopeptides prior to phosphorylation site determination by MS and MS/MS. 5.5.1 Selective purification of phosphoproteins by immunoprecipitation and affinity enrichment

Phosphospecific antibodies can efficiently enrich phosphoproteins by immunoprecipitation prior to subsequent separation techniques 316

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enabling a powerful targeted phosphoproteomics approach to investigate signaling pathways [183,184]. Antibodies specific to phosphotyrosine residues are used most commonly because a variety of high-affinity, high-specificity antibodies are commercially available. To a lesser extent, antibodies to pSer and pThr have been used. Antiphosphotyrosine antibodies can efficiently be utilized in order to detect, visualize or purify tyrosine-phosphorylated proteins. Cross-reaction with phosphoserine and phosphothreonine is not generally a problem with these immunoreagents. However, many of the commercially available monoclonal anti-phosphotyrosine Ig can also recognize nucleotides and phosphohistidine, which necessitates control experiments [126]. Screening for N-phosphates with antibodies to phosphohistidine is currently not possible due to unsuccessful attempts to prepare antibodies for phosphohistidine [126]. The combination of MS and elegant biochemical approaches based on high-affinity antiphosphotyrosine antibodies has become a very powerful tool for analysis of phosphotyrosine-regulated signaling pathways [139,181,183,210]. Most recently, Blagoev and Mann studied the global dynamics of phosphotyrosine-based signaling events in early growth factor stimulation, leading to the identification of virtually all known epidermal growth factor receptor substrates and aided resolving the time course of their activation upon epidermal growth factor stimulation [25]. Also, phosphotyrosine antibodies have been used in a ‘reactor’ column prior to LC-MS/MS by Aebersold and co-workers [6]. Several commercial anti-pSer/pThr antibodies can facilitate enrichment of phosphoproteins by immunoprecipitation as established by Grønborg and Pandey [77]. This study explored a range of commercial antibodies that could recognize phosphoserine/ phosphothreonine-containing proteins by Western blotting and investigated if any of these antibodies could be used to enrich for proteins phosphorylated on serine/threonine residues by immunoprecipitation. MS-based analysis of bands from one-dimensional gels, which were specifically observed in calyculin A-treated samples, resulted in identification of several known serine/threonine-phosphorylated proteins as well as novel phosphoproteins. The potentials of phosphoprotein enrichment using IMAC have been investigated in phosphoproteome studies [54,77]. Collins et al. [54], were more successful who integrated multiple IMAC fractionation steps on proteins and peptides level using optimized separation conditions for phosphoprotein enrichment using Ga(III)-NTA agarose. 317

A. Stensballe and R. J. Simpson 5.5.2 Multidimensional protein identification technology (MudPIT) strategy for ‘‘shotgun’’ phosphoprotein identification

Multidimensional protein identification technology (MudPIT) has been investigated for large-scale identification of phosphorylated proteins without prior affinity-based enrichment techniques. An integration of online multi-dimensional LC to data-dependent MS/MS and automated database searching constitute the MudPIT strategy for ‘‘shotgun’’ identification of protein modifications; here under protein phosphorylation [80,145,253]. The non-gel MudPIT strategy, popularized by Yates and co-workers, provides an alternative to gel-based proteomics strategies [154,242,243]. MudPIT is essentially a pre-fractionation of complex peptide mixtures by SCX capLC and subsequent separation by reversed-phase capillary LC coupled online to an ion-trap mass spectrometer for peptide sequencing. Macross et al. [145] applied the ‘‘shotgun’’ strategy for discovering co- and posttranslational modifications in simple and very complex protein mixtures and facilitated proteomic analysis of posttranslationally modified membrane proteins and protein topology in membranes [253]. The use of nonspecific enzymes results in the production of a large heterogeneous group of peptides and phosphopeptides. MudPIT analysis of the highly complex peptide mixture allowed each amino acid to be sequenced multiple number of times as it is part of several different peptides of varying length. Database search algorithms that are capable of identifying peptides without cleavage specificity allowed reassembly of protein sequence with an improved sequence coverage permitting localizing of membrane spanning regions of proteins and the polarity of their integration into the membrane. However, the overall effectiveness of the ‘‘shotgun’’ strategy for phosphoproteome analysis remains poor due to the fact that low-abundance phosphorylated peptides were still highly suppressed and to a large extent evaded detection due to the complexity of the peptide mixture. Large efforts are currently carried out to validate the authenticity of protein and PTM assignments by statistical methodology due to typically poor MudPIT data quality of generated by ion trap-MS rather than high-quality MS data enabling data mining with high constraints to reduce levels of false positives and false negatives [78,179]. 5.5.3 Ion-exchange-based separation of complex mixtures in phosphoproteome studies

In contrast to affinity enrichment of phosphopeptides in phosphoproteome studies, several reports highlight the usage of ion-exchange 318

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chromatography to increase detection of phosphorylated peptides in complex peptide mixtures. First, Ballif et al. [15] established that SCX chromatography optimized at pH 2.7 can enrich phosphopeptides into a far less complex mixture. Under these conditions, phosphopeptides have a lower solution charge state than the majority of tryptic peptides and elute in the first fractions during SCX. The caveats of this approach employing the relative lower solution charge state of phosphopeptides are that the enrichment only works with tryptic peptides and co-fractionation with histidine-containing, N-acetylated and c-terminal peptides in which the latter two have lower charge compared to internal unmodified tryptic peptides [78]. Second, Nuhse et al. [171] successfully reduced very complex peptides mixtures prior to IMAC enrichment by SAX thereby improving phosphopeptide coverage. A bias has been observed by several investigators [63,160] toward multi-phosphorylated peptides in large-scale IMAC-based experiments. Subfractionation of complex peptide mixture by SAX prior to IMAC enrichment significantly increased the total number of identified phosphopeptides, hereof a large number of singly phosphorylated peptides with very little overlap in identified phosphorylation sites. 5.5.4 Affinity-based enrichment of phosphopeptides in phosphoproteome studies

One of the first pioneering large-scale phosphoproteome analysis was carried out by Ficarro and co-workers investigating the phosphoproteome yeast Saccharomyces cerevisiae [63]. Enrichment of phosphopeptides from a whole-cell lysate was performed in a single experiment by combinations of IMAC purification of phosphorylated peptides, followed by automated nanoflow LC-MS/MS on an ion-trap for identification and mapping of phosphorylated peptides by bioinformatics tools. In order to enrich phosphopeptides with sufficient purity from wholecell lysate, the unfractionated peptide mixture was carboxy-methylated prior to IMAC in order to block the binding of unphosphorylated aspartate- and glutamate-containing peptides minimizing interference from non-specific binding of acidic peptides during IMAC enrichment of phosphopeptides otherwise reported. In a single experiment, more than 1,000 putative phosphopeptides were detected when the methodology was applied to the analysis of a whole-cell lysate readily defining 383 sites of phosphorylation in a total of 216 phosphopeptide sequences. 319

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The combination of IMAC and MS has now proven very efficient for large-scale phosphoproteome analysis in several other organisms. The specificity of the IMAC capture of phosphopeptides is not absolute, but high enough for large-scale identification of phosphorylation sites. Contaminant peptides in IMAC preparations have determined to mainly originate from highly abundant proteins and not acidic peptides [84,171]. Further refinement of the strategy by Ficarro et al. (2002) was used to investigate tyrosine phosphorylation in whole protein digests from capacitated human sperm including differential isotopic labeling to quantify phosphorylation changes occurring during capacitation [63]. Same strategy was employed by Salomon et al. including an additional phosphotyrosine protein enrichment step using antibodies for investigating the changes in tyrosine phosphorylation patterns occurring over time during either the activation of human T cells or the inhibition of the oncogenic BCR-ABL fusion product in chronic myelogenous leukemia cells in response to treatment with the drug ‘‘Gleevec’’ [210]. In an elegant ‘‘gel-free’’ strategy, Nuhse et al. [171,172] investigated the plasma membrane subphosphoproteome of Arabidopsis Thaliana. This strategy comprised of nanoflow LC-MS/MS (1D-LC), IMAC-enriched nLC-MS/MS (2D-LC) and SAX sub-fractionated and IMAC-enriched nLC-MSMS (3D-LC) (Fig. 5.20). The perhaps most comprehensive phosphoproteome study to date is the investigation of the pheromone signal transduction pathway in Saccharomyces cereviciae [78]. Gruhler and co-workers [78] combined SCX chromatography with IMAC and LC MS in a quantitative phosphoproteomic study of the yeast pheromone response resulting in the identification of more than 700 phosphopeptides of which 139 were differentially regulated at least two-fold and at least 20 belonged to proteins with explicit functions in pheromone signaling and mating. 5.5.5 Chemical derivatization strategies in phosphoproteome studies

Several strategies involve chemical modification of phosphopeptides or phosphoproteins within complex mixtures thereby making them amenable for specific enrichment. A method reported by Oda et al. [176] involves the replacement of Ser/Thr phosphate groups by a biotinylated affinity tag via band Michael addition. Modified peptides with an affinity tag can be enriched by high-affinity avidin–biotin coupling to immobilized avidin prior to MS and MS/MS analysis. At the same time, 320

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Aebersold and co-workers [262] introduced an alternative derivatization strategy that included conversion of pSer, pThr and pTyr phosphoamino acids into a phosphoramidate derivative. Both these methods have not yet matured for assessment of global protein phosphorylation, however, by derivatization with isotope-coded affinity tags these approaches enable quantitative comparison of phosphopeptide levels in different extracts [4]. A targeted chemical strategy applicable for phosphoserine and phosphothreonine residues has been demonstrated that converts these residues into protease-sensitive lysine analogs (aminoethylcysteine and beta-methylaminoethylcysteine, respectively) [127,128]. Adaptation of the aminoethylcysteine reaction to solid-phase support facilitated a catch-and-release strategy providing a one-step modification and enrichment of phosphopeptides.

5.6

BIOINFORMATICS TOOLS FOR PHOSPHOPROTEIN CHARACTERIZATION, HYPOTHESIS-DRIVEN PHOSPHOPROTEIN ANALYSIS AND DATA MINING

Protein phosphorylation is easily revealed by mass spectrometric analysis of proteins and peptide due to the mass increase of 80 Da per phospho moiety, allowing mass measurements by MALDI MS or by partial peptide sequence information obtained by ESI MS/MS to query protein or DNA sequence databanks (see Table 5.1) [111,185,256,257]. Database search engines such as Mascot, Sequest, Protein Lynx Global Server, SpectrumMill and VEMS2 are able to operate with partial covalent modifications, including phosphorylation of serine, threonine and tyrosine residues [153,190,203]. Prediction algorithms for phosphorylation site localization are available that will use prior knowledge of well-characterized phosphoproteins to predict probable phosphorylation sites in a query protein sequence (see Table 5.1) [26,132]. These algorithms enable rational design of experimental approaches aimed at determination of the actual utilized phosphorylation sites. Prediction methods may work very well for some phosphorylation motifs/domains in phosphoproteins, but generally tend to generate large number of candidate phosphorylation sites. However, kinase-specific neural network-based prediction algorithms now allow much more accurate predictions [92]. Knowledge of candidate phosphopeptides have shown to be useful for ‘‘inclusion 321

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lists’’ containing theoretical masses of putative phosphopeptides for selection of an increased number of precursor ions during iterative data analysis and for preferential selection of modified precursor ions during LC-MS/MS [54]. Carefully planning the choice of enzyme for proteolytic cleavage prior to separation and MS analysis will increase the likelihood of generating phosphopeptides in a molecular weight range optimal for MS and MS/MS. For this purpose in silico digestion of proteins by software programs (e.g. GPMAW) is recommended [189]. A range of publically available databases contains repositories of experimentally verified phosphorlation sites as well as prediction of protein domains (see Table 5.1).

5.7

QUANTITATIVE STRATEGIES FOR PHOSPHOPROTEOME ANALYSIS

Quantification of expressed phosphoproteins is a very important aspect of phosphoproteomics. Signal-transduction pathways and metabolic systems are highly dynamic processes in which successive phosphorylation and dephosphorylation events are responsible for biological activity of participating phosphoproteins. Hence, differential display of phosphoproteins or protein complexes (e.g. in response to stimulation) can provide insights into how phosphoproteins are regulated under certain physiological conditions and provide insights into cellular dynamics. MS is in principle a non-quantitative technique; however, semi-quantitative and relatively accurate quantitative estimates may be deduced directly from mass spectrometric data. MS-based strategies for protein quantification mainly differ in whether degree of phosphorylation of particular phosphorylation site changes over time (i.e. relative quantification) and what degree of occupation of each phosphorylation site is to be determined (i.e. absolute quantification). MS-driven methods for relative quantitative determination of protein phosphorylation can widely be accomplished by MS-based detection of relative isotopologue abundances using various means of stable isotope labeling (SIL) [16,112,151,182]. Differentially tagging of phosphoprotein/ phosphopeptide samples with either naturally occurring most abundant isotope ‘‘light’’ or isotopically enriched isotopes ‘‘heavy’’ allows detection of the isotopologue by MS after mixing the two samples. Each massencoded phosphopeptide thus appears as duplet of molecular ions 322

The phosphoproteome story TABEL 5.1 Publicly available Internet-based MS tools and protein sequence analysis services Peptide Mass Mapping (MS) ProFound Mascot PeptIdent FindMod

http://prowl.rockefeller.edu/ http://www.matrixscience.com http://www.expasy.org http://www.expasy.org

MS/MS data interpretation (peptide sequencing) Mascot http://www.matrixscience.com VEMS http://www.yass.sdu.dk ProteinProspector http://prospector.ucsf.edu/ Phosphorylation site prediction ScanSite NetPhos Prosite

http://scansite.mit.edu/ http://www.cbs.dtu.dk/services/ http://www.expasy.org

Protein phosphorylation site databases PhosphoELM http://phospho.elm.eu.org/ Phosphosite http://www.phosphosite.org/ Login.jsp Human Protein Reference http://www.hprd.org/ Database The Protein Kinase Resource http://pkr.sdsc.edu/html/index.shtml PlantsP http://plantsp.sdsc.edu Prediction of protein domains, function and gene ontology SMART http://smart.embl-heidelberg.de/ ProtFun http://www.cbs.dtu.dk/services/ PSORT http://www.psort.org/ Proteome Analyst http://www.cs.ualberta.ca/bioinfo/ PA/ SubLoc http://www.bioinfo.tsinghua.edu.cn/ SubLoc/ InterPro Scan http://www.ebi.ac.uk/interpro/ scan.html Geneontology http://www.geneontology.org/

323

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separated by a mass tag corresponding to the absolute molecular mass between the ‘‘light’’ and ‘‘heavy’’ forms, and the relative abundances of the peaks reflect the abundance of phosphopeptide in each sample. Incorporation of stable isotopes can be accomplished by a number of techniques including metabolic, enzymatic and chemical labeling. SIL strategies for quantification of protein phosphorylation can be grouped into three major categories (Fig. 5.21). First, metabolic labeling has been investigated in prokaryote cells, eukaryote cell culture systems and animal models. Differential conditions are obtained by growing cell cultures for days in ‘‘heavy’’ isotope-substituted culture media that contains isotopically labeled precursor molecules like 15N amino acids, 2 H-leucine or 13C-leucine/lysine/arginine. Whole-cell SIL using 15N labeling method was used for MS-based relative quantitation of yeast protein phosphorylation under various growth conditions [175]. Ibarrola and Pandey [100] investigated stable isotope-containing amino acids in cell culture (SILAC) for the relative quantitation of phosphorylation by double-isotope labeling with [13C6]lysine and [13C6]arginine. In this study, SILAC enabled quantitation of the extent of known phosphorylation sites as well as identification and quantitation of novel phosphorylation sites. Second, post-biosynthetic labeling of intact proteins and peptides was performed by chemical derivatization in vitro. Quantification of the relative levels of expressed dephospho:phosphoproteins was investigated by b-elimination of phosphate from phospho-Ser/Thr followed by Michael addition of ethanethiol and/or ethane-d(5)-thiol SIL and LC-MS analysis [69,244]. Goshe and co-workers [74] also utilized the b-elimination/Michael addition chemistry for incorporation of an isotope-coded affinity tag (PhIAT) in phosphopeptides. Differential derivation of model phosphopeptides with either light or heavy PhIAT reagent-enabled identification identified and quantification by LC-MS/MS. Stover et al. [235] successfully investigated differential phosphoprofiles using labeling of acidic residues with d0 and d3 methyl esters, previously introduced by Ficarro and co-workers [62]. Third, incorporation of stable isotope labels can be during or after proteolysis by enzymatic digestion 18 in H16 2 O or isotopically enriched H2 O [16,206]. The proteases trypsin, Lys-C and Glu-C are capable of incorporating two 18O molecules during proteolysis (4 Da mass tag) or by post-digest singly and doubly exchanging of 18O [16]. Bonenfant and co-workers devised a strategy combining 18 affinity selection of H16 2 O/H2 O-labeled phosphopeptides from the combined digests by IMAC followed by dephosphorylation with alkaline 324

The phosphoproteome story B Pre-digestion (in vitro)

A In vivo labeling Sample stage A 12C; 14 N

Sample stage B 13C; 15N; 3D

Sample stage A

Sample stage B

Sample stage A

Sample stage B

(SILAC)

Extract/Fractionate Combine

Extract/Fractionate

Digest

C Post-digestion (in vitro)

Label (light/heavy)

Combine Digest

Extract/Fractionate Digest/Label Label (light/heavy)

Combine

Identify & Quantify

Fig. 5.21. Schematic representation of MS-based quantification methods for the study of protein phosphorylation by stable isotope labeling. (A) Labels can be introduced in vivo by SILAC; (B) in vitro labeling of proteins before digestion or (C) in vitro labeling of proteins after digestion of phosphoproteins.

phosphatase to allow for quantitation of changes of phosphorylation by MALDI MS [28]. Absolute quantitation of protein phosphorylation levels can be determined using several analytical strategies. First, synthetic heavy isotope-labeled phosphopeptides can be used as internal peptide standards for stable isotope dilution in the AQUA strategy [68]. Here the expected unphosphorylated or phosphorylated proteolytic peptides are spiked into the samples in known quantities, such that the degree of phosphorylation can be inferred by monitoring the signal intensities. A major advantage of this method is that it does not require living cells and can therefore be used to study in vivo phosphorylated proteins such as tissue samples. To ensure high accuracy of quantitation, the synthetic isotope-labeled phosphopeptide can be prepared in a prolonged peptide construct incorporating an amino acid specific to the chosen protease within the chemically synthesized peptide, thereby minimizing errors resulting from alterations in digestion efficiency [112]. Second, site-specific phosphorylation stoichiometries of singly phosphorylated species have been derived from the comparison of chemically identical but isotopically distinct peptide species by SIL, 325

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dephosphorylation of one fraction, pooling of the two fractions followed by MS analysis [87,261]. The degree of phosphorylation was then derived by comparing the signal intensity of the two differentially labeled unphosphorylated species, assuming that the increase in signal intensity of the species is due to the dephosphorylation of the singly phosphorylated species. Hegeman et al. [87] also estimated the extent of phosphorylation from the mass-spectral-peak areas for the phosphorylated and unmodified peptides, and these estimates, when compared with stoichiometries determined using the isotope-coded technique, differed only marginally (within approximately 20%). Recently, Steen and Kirschner [224] have described means for a stable isotope-free strategy for both relative and absolute phosphorylation stoichiometry using careful statistical analysis of LC-MS/MS data sets of phosphorylated peptides. A robust normalization routine accounts for run-to-run variations and variations in starting material thereby allowing relative quantitation based on determination of variations in normalized ion currents of a phosphopeptide and its unmodified cognate. Absolute phosphorylation stoichiometry is determined by measuring the ion currents of a particular phosphopeptide and its unmodified cognate, because the changes in the signal intensities of the phosphorylated and unphosphorylated form of a peptide are correlated.

5.8

SUMMARY

Despite the progress in performance of MS-driven analytical methods during the last decade with regard to sensitivity and selectivity, the identification of phosphorylation sites is still not a trivial task. Thus, today no single method can reliably detect and characterize all modified residues in a phosphoprotein and far most successful analysis strategies for phosphoprotein and phosphoproteome, including multiple levels of enrichment and separation methods as well as biological follow-up analysis. However, recent improvements in MS have spawned improved and far more robust analytical strategies. An improved effiency of enrichment and separation techniqes on both peptide and protein level, the improved data quality by ECD or ETD peptide fragmentation and the improved confidence in phosphopeptide detection by MS3 phosphopeptide sequencing using high mass accuracy FTICR-based mass spectrometers have enabled multiple comprehensive studies of protein 326

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phsophorylation. Last, but not least, multiple complementary MS-driven strategies for relative and absolute quantitation of protein phosphorylation will ease rapid investigations of signal transduction systems and, thus, provide the basis for great advances.

ACKNOWLEDGMENTS I greatly thank Professor Ole Nørregaard Jensen, Protein Research Group at the University of Southern Denmark for his supervision during method development for mass spectrometry-based phosphoprotein analysis. This work is supported by a grant from the Danish Natural Sciences Research Council. Dr. Nick Morrice and Dr. Daniel Profrock are acknowledged for contributing to figures in this work.

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