Mapping Posttranslational Modifications of Proteins by MS‐Based Selective Detection: Application to Phosphoproteomics

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[5] Mapping Posttranslational Modifications of Proteins by MS‐Based Selective Detection: Application to Phosphoproteomics By STEVEN A. CARR, ROLAND S. ANNAN ,

AND

MICHAEL J. HUDDLESTON

Abstract

This chapter outlines general principals that apply to the analysis of posttranslational modifications of proteins, with an emphasis on phosphoproteins. Mass spectrometry (MS)‐based approaches for selective detection and site‐specific analysis of posttranslationally modified peptides are described, and an MS‐based method that relies on production and detection of fragment ions specific for the modification(s) of interest and that was developed in the authors’ laboratory is described in detail. The method is applicable to selective detection of N‐ and O‐linked carbohydrates in glycoproteins, O‐linked sulfate, and N‐ and O‐linked lipids. Detailed procedures for application of this strategy to phosphorylation‐site mapping are presented here.

Introduction

Proteins that have covalent modifications are the rule rather than the exception in nature. Nearly 200 structurally distinct covalent modifications have been identified thus far, ranging in size and complexity; these modifications result from a variety of factors, from the conversion of amides to carboxylic acids (delta mass þ0.9840) to the attachment of multiple complex oligosaccharides each with molecular masses up to several thousand daltons (Graves et al., 1994; Krishna and Wold, 1993; Wold, 1981). Most post‐translational modifications are introduced by enzymes. The presence of the modification is often required for normal biological function or tissue disposition of the protein, although, in many cases, the role of the modification is as of yet unknown. Phosphorylation and glycosylation are two of the most biologically relevant and ubiquitous posttranslational modifications of proteins. In both cases, an organism’s commitment to these modifications, as measured by the estimated numbers of genes involved, is substantial. Protein kinases may constitute as much as 3% of the entire eukaryotic genome (Cohen, 1992; Hubbard and Cohen, 1993; Hunter, 1991), while gene products involved in oligosaccharide biosynthesis may represent as much as 1% of METHODS IN ENZYMOLOGY, VOL. 405 Copyright 2005, Elsevier Inc. All rights reserved.

0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)05005-6

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the genome (Varki and Marth, 1995). It is estimated that as many as one‐ third of proteins present in typical mammalian cells are phosphorylated (Hubbard et al., 1993), and up to half of the proteins are glycosylated (Apweiler et al., 1999). Phosphorylation is the most common and physiologically important reversible regulatory modification. An exquisitely complex, integrated network of protein kinases and phosphatases controls many aspects of cell growth, metabolism, division, motility, and differentiation through selective phosphorylation (often at multiple sites) and dephosphorylation of cellular proteins (Cohen, 1992; Hubbard and Cohen, 1993; Hunter, 1991). Phosphorylation of intracellular proteins plays an essential role in signal transduction. This is the process by which extracellular signals are communicated to the cell’s nucleus by the binding of components such as cytokines, neurotransmitters, and hormones to cell‐surface receptors, thereby regulating an array of intracellular physiological processes (Daum et al., 1994; Eck, 1995; Pawson, 1995; Schlessinger, 1994). Posttranslational modifications frequently complicate or even prevent the use of classical tools for protein sequence analysis, such as automated Edman degradation. In addition, the presence of lipid or carbohydrate on proteins can dramatically decrease the accuracy of molecular weight estimates obtained by sedimentation velocity, gel permeation, or SDS– PAGE measurements. Unlike classical biochemical techniques, mass spectrometry (MS) relies on entirely different principles to accomplish structure analysis. Because MS measures the mass of a molecule, it is uniquely suited for detection and structural characterization of covalent posttranslational modifications of proteins that involve either a mass change to an individual amino acid in the sequence or a removal of a portion of the N and/or C terminus of the protein. From the authors’ viewpoint, the potential for MS in the study of posttranslational modifications is virtually unlimited. This chapter outlines general principals that apply to the analysis of posttranslationally modified proteins, with an emphasis on phosphoproteins. The chapter then describes an MS‐based approach for selective detection and site‐specific analysis of posttranslationally modified peptides. It is important to note that the method does not require or depend on incorporation of a tag or label (e.g., a radiolabel like 32P or 33P or an isotopic label like 18O) but rather on production and detection of fragment ions specific for the modification of interest. Detailed procedures for application of this strategy to phosphorylation‐site mapping are presented as well (Annan et al., 1997, 2001; Bean et al., 1995; Carr et al., 1996; Hill et al., 1994; Huddleston et al., 1993a; Hunter et al., 1994; Neubauer et al., 1997; Verma et al., 1997); other related strategies are referenced in Table I.

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TABLE I MARKER‐IONS DERIVED FROM COMMON POSTTRANSLATIONAL MODIFICATIONS m/z

Origin

Indication

PO2 /PO3

63/79

O‐linked phosphate

Phosphopeptides containing pThr, pSer, and pTyr; phosphocarbohydrates

C8H10NO4Pþ

216.043b

pTyr

SO3

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Immonium ion of pTyr O‐linked sulfate

Allen et al., 1997; Annan and Carr, 1997; Annan et al., 2001; Azzam et al., 2004; Bean et al., 1995; Carr et al., 1996; Chen et al., 2002; Crabb, in press; Huddleston et al., 1993a; Hunter and Games, 1994; Jedrzejewski and Lehmann, 1997; Le Blanc et al., 2003; Neubauer and Mann, 1997; Sulivan et al., 2004; Till et al., R 1994; Verma et al., 1997; Wilm et al., 1996; Watty et al., 2000; Zappacosta, 2002 Steen et al., 2001, 2003 Bean et al., 1995

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Sulfopeptides; sulfocarbohydrates

References

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Fragment ionsa

Glycopeptide or carbohydrate containing any N‐ or O‐linked sugar

292

N‐acetyl neuraminic acid

Glycopeptides or carbohydrates containing complex‐type N‐ or O‐linked sugars

Carr et al., 1993; Greis et al., 1996; Hayes and Aebersold, 2000; Huddleston et al., 1993b; Kragten et al., 1995; Mazsaroff et al., 1997; Medzihradszky et al., 1997, 1998; Roberts et al., 1995; Rush et al., 1995; Schindler et al., 1995; Sullivan et al., 2004 Carr et al., 1993; Greis et al., 1996; Hayes and Aebersold, 2000; Huddleston et al., 1993b; Kragten et al., 1995; Mazsaroff et al., 1997; Medzihradszky et al., 1997, 1998; Roberts et al., 1995; Schindler et al., 1995 (continued )

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N‐ or O‐linked HexNAc (e.g., GlcNAc, GalNAc)c

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204

85

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Fragment ionsa

(C15H31CONH2þ and C15H31COþ)

a

m/z

Origin

Indication

366

N‐ or O‐linked Hex‐ HexNAc (e.g. Gal‐GlcNAc, Gal‐GalNAc)c

Glycopeptides or carbohydrates containing any N‐linked and most O‐linked sugars

256 239

N‐linked palmitic acidd

Lipopeptides or lipoproteins

References Carr et al., 1993; Greis et al., 1996; Hayes and Aebersold, 2000; Huddleston et al., 1993b; Kragten et al., 1995; Mazsaroff et al., 1997; Medzihradszky et al., 1997, 1998; Roberts et al., 1995; Schindler et al., 1995 Bean et al., 1995; Gu et al., 1997; Rush et al., 1995; Sullivan et al., 2004

Charge on fragment indicates required analysis mode (positive or negative ion). Numerous potential interferences exist at nominal mass 216, which requires use of instruments capable of accurate mass and high resolution. c Fragment ions shown will form regardless of sequence position of indicated sugar in the carbohydrate or if it is directly linked to the peptide; ions due to consecutive losses of water (18 Da) are also commonly observed. d Other lipids (saturated and unsaturated) will produce analogous marker‐ions at the expected masses. b

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TABLE I (continued)

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Similar methods for the analysis of N‐ and O‐linked carbohydrates in glycoproteins (Carr et al., 1993b; Greis et al., 1996; Gu et al., 1997; Hayes et al., 2000; Huddleston et al., 1993b; Kragten et al., 1995; Mazsaroff et al., 1997; Medzihradszky et al., 1997; Roberts et al., 1995; Schindler et al., 1995), O‐linked sulfate (Bean et al., 1995), and N‐ and O‐linked lipids (Bean et al., 1995; Gu et al., 1997) have also been developed (see Table I). Detailed procedures for analysis of these posttranslational modifications may be found in the references cited at the end of this chapter. Since this chapter was written in 2000, there has been a dramatic increase in the phosphoproteomics literature. The expansion in phosphoproteomics has been aided in large part by significant improvements in the ability of mass spectrometers to carry out data‐dependent experiments, such as the ability to automatically select and further fragment ions that have a mass‐to‐charge ratio (m/z) corresponding to neutral loss of phosphoric acid (Bateman et al., 2002; Covey et al., 1991; Schroeder et al., 2004). In addition, selective enrichment strategies for phosphoproteins and phosphopeptides based on the use of immunoprecipitation‐capable antibodies (Rush et al., 2005) and immobilized metal affinity chromatography (Ficarro et al., 2005) have seen great improvement through 2005. For a recent review of the phosphoproteomics literature, which emphasizes strategies that have been proven useful for identification of previously unknown phosphorylation sites, we recommend the review by Loyet et al., (2005). Special Issues with Respect to the Analysis of Posttranslationally Modified Proteins: Focus on Phosphorylation‐Site Mapping

The goals of any posttranslational modification analysis are to provide as complete a map as possible of the modification sites in a protein and to define the structure(s) of the modifications at each specific attachment site. This requires detection and analysis of peptides covering all of the potential modification sites. It is important to note that this is a much more stringent analytical requirement than for protein identification by MS and database searching (a subset of ‘‘proteomics’’), which can often be accomplished with molecular weight and partial sequence for any peptide (preferably more than one) derived from the protein (Larsen et al., 2000). A reasonable starting strategy is to obtain coverage of the known consensus sites for a given modification. However, it must always be kept in mind that consensus sites serve only as a guide. An exclusive focus on predicted sites will work against finding sites of attachment that violate the ‘‘known’’ consensus rules (a fairly common occurrence in the case of

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phosphorylation). The goal in analyzing posttranslationally modified proteins should be to obtain as complete a sequence coverage as possible. This goal is often difficult to achieve in practice for single phosphoproteins, and it is presently unachievable for the analysis of large numbers of phosphoproteins as complex mixtures (phosphoproteomics). Site‐specific analysis of posttranslational modifications by either conventional approaches or MS requires that the modified protein first be cleaved enzymatically or chemically into peptides of a size suitable for sequence analysis. In the case of MS, this size is ideally between 500 Da and 3000 Da. Trypsin is usually the first choice because the rules for fragmentation of peptides having Lys or Arg at their C termini are well understood and predictable. It is possible to directly analyze unfractionated protein digests by techniques such as nanoelectrospray mass spectrometry (nanoESMS) and matrix‐assisted laser desorption/ionization (MALDI). Methods employing direct analysis of unfractionated peptide digests minimize losses that invariably occur in sample handling at low levels. However, direct analysis of complex mixtures suffers from well‐ known problems of suppression effects (particularly for highly charged peptides like phosphopeptides), charge‐state overlap, limited dynamic range for peptide signal detection, and limited sampling frequency of the mass spectrometer. To minimize these problems and to maximize sequence coverage of posttranslationally modified proteins, it is necessary to employ chromatographic or electrophoretic separation to fractionate the mixture prior to or during MS analysis. Several studies of more highly phosphorylated proteins illustrate the advantages of employing a separation stage prior to analysis (Watty et al., 2000; Wu et al., 2000). Use of high‐ performance liquid chromatography (HPLC) during on‐line liquid chromatography (LC)–ESMS (as employed in the strategy detailed below) is particularly effective as it accomplishes both desalting and peptide separation in one step. A separation step is also required after digestion and prior to analysis by MS if the protein has been reduced and alkylated (to maximize sequence coverage) or if the protein was derived from digestion of a gel band. In both instances, the sample will contain excess reagents, by‐ products, or impurities that would severely compromise or defeat analysis by either MALDI–MS or ES–MS if not removed. In addition to the general concerns previously noted for analysis of posttranslationally modified proteins, phosphoproteins present some unique challenges. Many phosphoproteins of interest are present in cells at only very low concentrations, and often only femtomole to low picomole amounts of phosphopeptides may be recovered for analysis following enzymatic digestion and chromatographic isolation (Bean et al., 1995; Boyle

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et al., 1991; Luo et al., 2005). Phosphoproteins are also usually phosphorylated on a number of different sites throughout the protein, with individual sites being phosphorylated to varying degrees (e.g., 1 to 100%). Upon digestion with trypsin (or another enzyme), it is very common to generate peptides containing more than one potential phosphorylation site (Ser, Thr, and Tyr). Furthermore, an observed molecular mass for a phosphopeptide can sometimes be assigned to more than one reasonable peptide sequence from the protein. Thus, it is necessary to sequence phosphopeptides by MS/MS to assign the part of the protein sequence from which that the peptide derives and to establish which residues in that peptide are phosphorylated. To estimate the stoichiometry of modification at each site, both the modified and unmodified forms of each phosphopeptide must be detected so that the ratio of the ion abundances can be determined. The serious constraints of low phosphopeptide yield and the mixture complexity noted earlier make it desirable, if not essential, to have analytical methods capable of preferentially detecting and analyzing phosphopeptides. By far, the most commonly employed techniques for the isolation of phosphopeptides from phosphoproteins have employed cells metabolically labeled in vivo or in vitro with [32P]‐phosphate or proteins that are labeled in vitro with [32P]‐phosphate by reaction with purified kinase (Boyle et al., 1991; Kuiper et al., 1995; Luo et al., 2005; Wettenhall et al., 1995; Winz et al., 1994). Efforts to identify and/or isolate phosphopeptides that circumvent the requirement for radiolabeling have centered mainly on two approaches: (i) use of immobilized metal‐ion affinity techniques both off‐ line (Posewitz and Tempst, 1999) and coupled on‐line with an ES mass spectrometer (Posewitz and Tempst, 1999; Watts et al., 1994) and (ii) selective detection in MS‐based methods on the unique fragmentation behavior of phosphopeptides (Carr et al., 1996; Huddleston et al., 1993a). The latter MS‐based methods are described in detail in the upcoming paragraphs. As already noted, since this chapter was written in 2000, a number of advances have been introduced for phosphopeptide detection and sequencing. A recent survey of the literature that is current to 2004 can be found in Loyet et al. (2005). Selective Detection of Posttranslational Modifications by MS

Post‐translational modifications on proteins are often more susceptible to cleavage by collision‐induced fragmentation in the mass spectrometer than the peptide backbone. The methods the authors of this chapter and others have developed for selective detection of such modifications take advantage of this fact using three approaches: (i) detection of the loss of the

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modification from the peptide precursor, (ii) detection of the low mass ‘‘signature’’ or ‘‘marker’’ ions from the modification itself, or (iii) selective detection of the precursor ions that give rise to the low mass signature peaks (referred to as precursor‐ion scanning). Neutral loss monitoring has long been used in ESMS for detection of phosphorylation (Covey et al., 1991). This approach has been automated on orthogonal time‐of‐flight mass spectrometers (Bateman et al., 2002) and on ion‐trap instruments (Schroeder et al., 2004). The approach relies on the detection of the loss of 98, 98/2, 98/3, and so on from singly, doubly, triply, and other multiply charged precursors of phosphopeptides. Detection of the loss can be used to trigger acquisition of MS/MS spectra for sequencing of the peptide. Neutral loss of phosphoric acid from pSer and pThr is a facile process on any mass spectrometer, but it is especially dominant on ion trap instruments that induce fragmentation by resonance excitation. To compensate for the lack of sequence informative backbone cleavage in the ion trap MS/ MS spectra, automated procedures have been developed that carried out MS3 on the neutral loss peak(s). The strategies developed in the authors’ laboratory utilize marker‐ ion production and precursor‐ion scanning in specific combination to selectively identify peptides containing a particular covalent modification and to selectively fragment modified peptides to define the attachment site(s) of the modification (Annan et al., 2001; Zappacosta et al., 2002). Briefly, the experiments involve first a modification‐specific marker‐ion scan that enables selective detection and fractionation of modified peptides in a protein digest during LC–MS analysis. The marker‐ions are produced by low energy collisions with gas molecules in the premass analysis region of the mass spectrometer (see Table I). Second, the experiments use a precursor‐ion scan during LC–MS analysis or of fractions collected during a marker‐ion analysis. This experiment identifies the molecular weights of only those peptides carrying the modification of interest. Third, the experiments perform sequence and structure analyses of the identified precursor ions by MS/MS. The first two steps just described detect the presence of a modification and establish the molecular masses of modified peptides in the presence of an excess of unmodified peptides, even in cases where the signal from the modified peptide is indistinguishable from background in the normal MS scan. Knowledge of the molecular masses of the modified peptides enables targeted use of MS/ MS for sequencing. These experiments and how they interrelate are described in greater detail in the following paragraphs for phosphoprotein analysis.

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Selective Detection and Preparative Fractionation of Modified Peptides Using Marker Ions Modified peptides may be selectively detected in mixtures of unmodified peptides by formation and detection of low‐mass fragment ions that serve as specific markers for the modifications of interest. Abundant low‐ mass marker ions, however, are not usually observed in ESMS spectra under normal operating conditions. All mass spectrometers, by design or default, have one or more defined regions of reduced but still relatively high pressure between the atmospheric pressure ion source and the true high vacuum regions where the mass separation and detection elements of the mass spectrometer reside (Fig. 1A). In conventional LC–ESMS analysis, the velocity of the ions in these relatively high‐pressure zones is kept low to minimize fragmentation that occurs as a result of collision with gas molecules and maximize transmission of intact parent ions. To produce abundant marker ions, the velocity of the ions in this region is increased to induce fragmentation via collision‐induced dissociation (CID) of all ions prior to mass separation. The highest sensitivity for marker‐ion detection is obtained by selected ion monitoring (SIM) of just the marker ions of interest under conditions of continuous, high‐level CID in the ion‐sampling/collision region. Alternatively, CID may be ‘‘turned on’’ while scanning the lower mass range and then ‘‘turned off’’ during the remainder of each scan. In this stepped collision‐energy scanning mode, peptide molecular weight information and modification‐selective, low‐mass marker ions may be detected in the same scan, albeit at lower sensitivity for detection of the marker ions. While these experiments can be carried out on any type of mass spectrometer, they are ideally suited to a single quadrupole mass spectrometer (or one mass analyzer of a triple quadrupole) because of the ability to accumulate ion current from one or a few masses/markers so as to increase sensitivity of detection. The SIM traces for the marker ions indicate which regions of the chromatogram contain the modified peptides (Fig. 1B). Marker‐ion traces are readily compared with the total‐ion current (TIC) and/or the UV chromatogram traces to give an indication of peak complexity. In the case of phosphopeptides, the marker ion profile is analogous to the output from an HPLC radioactivity detector or the autoradiogram from a two‐dimensional phosphopeptide map but without the requirement for 32P or 33P labeling of the protein. This trace also serves as a fingerprint for the phosphorylation profile of a protein. Changes in the phosphorylation state of a protein would be reflected by a change in the phosphorylation profile. Only those components of the profile that change would need further analysis. By splitting

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FIG. 1. Selective detection and preparative fractionation of modified peptides using marker ions during LC–ESMS. (A) Typical experimental arrangement for formation and detection of modification‐specific marker ions during LC–MS analysis of proteins. (B) Representative data showing how fractions containing modified peptides are identified and collected.

the column effluent, greater than or equal to 80% of the modified peptides are fraction‐collected during the on‐line LC–ESMS analysis (Fig. 1A). Columns with internal dimensions of 180
m or larger may be effectively used in this first dimension step (see Zappacosta et al., 2002 and detailed procedures described later in this chapter).

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Table I contains a list of useful marker ions. The general approach used for identification of suitable marker ions for modifications of interest is to analyze model compounds, such as suitably modified peptides, proteins, and carbohydrates, in product‐ion scanning, MS/MS mode to ascertain if there are appropriate diagnostic fragments produced and to determine the optimum collision energy for their formation. The model compounds are then added to defined mixtures of peptides or into a digest of a model protein or mixtures of proteins to test for specificity and sensitivity of detection. The m/z 63 and 79 marker ions appear to be highly specific for phosphopeptides. Other markers, such as m/z 204 used for carbohydrate analysis (see Table I), may occasionally be formed from CID of unmodified peptides. However, nonspecific fragment‐ion formation is usually readily distinguished from modification‐specific responses by the observation of synchronous changes of several (or all) of the marker ions for a given modification. For example, in the case of glycopeptides or carbohydrates, one would commonly monitor m/z 204, 292, and 366 (see Table I) in a single LC–ESMS experiment. Coincident increases in response for two out of three of these ion traces would indicate a specific rather than a nonspecific response. Ions due to loss of water from the ions just listed can also be monitored for an added degree of assurance. The use of marker‐ion formation and detection is illustrated for phosphorylation analysis of bovine s‐casein in Fig. 2. Phosphopeptide content was assessed by SIM of m/z 63 and 79 during negative‐ion LC–ESMS. Superposition of the SIM trace on the UV trace identified eight HPLC fractions eluting between 13 to 22 min as containing phosphopeptides. These collected fractions were further analyzed in the second and third dimension steps of the analysis described in the following paragraphs. Defining the Molecular Masses of Modified Peptides by Precursor‐Ion Scanning and Sequence Analysis by MS/MS The fractions containing the modified peptides that were isolated in the marker‐ion experiment almost always contain additional, unmodified peptides that can make recognition of the masses of modified peptides difficult or impossible. Furthermore, the modified peptides may represent only a few percent of the total peptide in the fraction and may not even be discernable in a conventional MS scan of the fraction. To selectively identify the masses of the modified peptides, a precursor‐ion scan is used that only produces signals for those ions that fragment to yield the marker‐ion monitored. Figure 3 illustrates the precursor scan experiment in a triple quadrupole mass spectrometer for detection of phosphopeptide precursor ions. Triple

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FIG. 2. Selective detection of phosphopeptides of bovine s‐casein. (A) LC–ESMS SIM trace for m/z 63 and 79. Phosphopeptide content was assessed by selected ion monitoring (SIM) of the marker ions m/z 63 and 79 during negative‐ion LC–ESMS. (B) LC–UV trace for tryptic digest of bovine ‐casein. Superposition of the SIM trace on the UV trace identified eight HPLC fractions eluting between 13 and 22 min as containing phosphopeptides. These collected fractions were further analyzed in the second and third dimension steps of the analysis as described in the text.

quadrupole mass analyzers equipped with ion‐counting detectors provide the highest sensitivity of any mass analyzer for precursor‐ion scanning due to their capability to signal‐time average very weak data and reject noise. In this simplified scheme, three precursor ions (M1, M2, and M3) are emerging from the electrospray needle at the start of a 2s‐scan of the mass

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FIG. 3. Scheme illustrating precursor‐ion scanning to determine the molecular masses of phosphopeptides for subsequent MS/MS analysis. (A) Q1: Normal scan (2s scan). Observed are (M – H) of all peptides. (B) Q2: Collison‐induced decomposition of all ions. All peptides fragment; only phosphopeptides yield m/z 79 marker ion, which is observed only when peptide 3 fragments. (C) Q3: Selective detection of m/z 79. Observed intensity of m/z 79 was recorded at the m/z of the precursor that produced it in Q1. See text for detailed discussion.

spectrometer’s first mass analyzer (Fig. 3A). As the m/z scan of Q1 progresses, the three precursor ions are transmitted sequentially and individually into the true collision cell, Q2, where each fragments (Fig. 3B). The fragment patterns for all three are shown superimposed, color coded to the parent from which they derived. The second mass analyzer (Q3) is set to pass one marker ion (e.g., m/z 79 for phosphopeptides, Fig. 3C). Only precursor M3 fragments to produce a negative ion of m/z 79. The precursor‐ion spectrum is obtained by recording the observed ion abundance of

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m/z 79 at the m/z of the precursor that produced it, which in this case is M3. For simplicity, the authors show only a single peak for each precursor in Fig. 3A. However, ESMS spectra often exhibit multiple charge states for any given parent/precursor. In practice, each member of a charge series for a given phosphopeptide precursor will fragment to produce m/z 79, and so each will be recorded in the resulting precursor‐ion scan. Precursor‐ion scans may also be used to selectively detect modified forms of an intact protein analyzed by ESMS (Neubauer et al., 1997). Precursor‐ion scanning can be accomplished in an on‐line LCMS mode as was first demonstrated for glycopeptide analysis (Carr et al., 1993) or in an off‐line mode following collection of fractions identified to contain the modified peptide by marker‐ion scanning. At the time, we, the authors, had developed this strategy, data‐dependent control of the triple quadrupole MS instrument used was very limited. This necessitated the use of an off‐line approach in which phosphopeptide‐containing fractions were subsequently analyzed off‐line by precursor‐ion scanning using nanoelectrospray (Carr et al., 1996; Wilm et al., 1996). Improvements in control of scan functions and data‐dependent experiments have enabled MS systems, such as the AB 4000 triple quadrupole, to automatically switch from negative‐ion precursor‐ ion mode to positive‐ion mode full‐scan MS/MS mode following detection of the m/z 79 precursor ion, which permits the analysis to be carried out in the course of a single LC–MS experiment (Le Blanc et al., 2003). The selectivity of this step for phosphopeptide detection is illustrated in Fig. 4 for analysis of phosphopeptide‐containing fraction 18 collected during the LC–ESMS analysis of bovine s‐casein (see Fig. 2). Panels A and B compare the full‐scan, negative‐ion nanoelectrospray mass spectrum with the precursor ion spectrum for m/z 79. These data were acquired from circa 0.5
L of the collected fraction after adjusting the pH to greater than 10 to improve the sensitivity for detection of the phosphopeptides (see following subsection entitled Procedures). The phosphopeptide of determined Mr ¼ 1832.8 is clearly a very minor component and, in the absence of the precursor‐ion data, may otherwise have gone unnoticed in this fraction. In contrast, only this phosphopeptide is evident in the precursor‐ion spectrum (Fig. 2B). Once the molecular weights of the phosphopeptide ‘‘needles in the haystack’’ are determined, it is straightforward to switch analysis modes and select the masses for (M þ 2H)2þ, (M þ 3H)3þ, and additional forms of the phosphopeptides in the positive ion data for sequencing by nanoESMS/ MS (Fig. 4C). Generally, no additional sample is required because most of the 0.5–1
l aliquot of sample that provided the precursor‐ion scan data is usually remaining. The nanoESMS/MS product‐ion spectrum of the (M þ 2H)2þ parent ion of m/z 917.4 is shown in Fig. 5. The series of ions marked

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FIG. 4. Selective phosphopeptide analysis of fraction 18 collected during the LC–ESMS analysis of bovine s‐casein (see Fig. 2). (A) Full scan negative‐ion nanoelectrospray mass spectrum. (B) Precursor‐ion mass spectrum for m/z 79. (C) Full scan positive‐ion mass spectrum. The precursor‐ion scan detects a single phosphopeptide with an apparent average mass of 1833.6. Arrows point to the (M 2H)2 and (M þ 2H)2þ ions in the respective full‐ scan, negative‐ion data (shown in A) and positive‐ion data (shown in C) that are dominated by signals from nonphosphorylated peptides coeluting in this fraction.

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FIG. 5. Sequence localization of modification by MS/MS. (A) NanoES (þ) ion CID product ion spectrum of m/z 917.2, the doubly charged ion for the phosphopeptide found in fraction 18. The partial sequence XEX (where X ¼ Leu or Ile) is readily determined by inspection. This partial sequence matches part of a known phosphorylated sequence from s1‐casein, but the mass does not fit this phosphopeptide suggesting further modification. (B) Expansion of the low mass region of the spectrum shown in panel A. Internal fragment ions produced by cleavage of two amide bonds are represented by the single letter amino acid code for the residues included in the internal fragment. (C) Final sequence determined from the product ion spectrum. The amino acid sequence coverage provided by bn and yn fragment ions is shown. Sequence corresponds to a previously unknown variant of s1‐casein.

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y8 – y11 in the spectrum clearly indicate the partial sequence XEX (X ¼ Leu or Ile), which matches residues 109–111 from the sl‐casein tryptic peptide, VPQLEIVPNSAEER (residues 106–119, calc. Mr avg. 1660.8), which was identified in an earlier eluting fraction. Interpretation of the lower m/z region of the spectrum allowed us, the authors of this chapter, to extend the determined sequence toward the C‐terminus using yn ions, yielding the partial sequence XEXVPNpSAEER. Analysis of the low m/z region (Fig. 5B) led to the identification of a series of b‐ions (and some a‐ions) that overlapped with the sequence already defined and extended toward the N terminus of the peptide to yield (E,F or Y,X) GYLEIVPNpSAEER. The choice and order of the first two residues are defined by a series of internal fragment ions that included XGYL and XGYLE. The full sequence of the peptide is therefore YXGYLEIVPNpSAEER, which corresponds to a previously unreported variant of bovine s1 casein. There are several reasons to include sequencing in any modification mapping strategy. The most obvious is the case in which a given peptide contains more than one potential modification site. It is also relatively common to observe peptide signals for which no reasonable sequence assignment can be made based on the determined molecular weight or where the assignment is ambiguous. The quality of the MS/MS data required to confirm peptide identity from a protein of known sequence and to localize the modifications need not be as high as for de novo sequencing. Even weak, incomplete MS sequence data is often sufficient to answer the specific questions noted earlier. Nanoelectrospray provides long data acquisition times and very good sensitivity of analyte detection from very small sample volumes (Wilm and Mann, 1994a,b). This permits optimization of all experimental parameters and sensitivity improvement by accumulating ion counts for weak signals. This is especially important for large peptides and whenever a more complete sequence is required (e.g., in cases where the protein sequence is not known and is not in a database). Furthermore, several peptides can be sequenced without resorting to the use of another aliquot of sample. By making the pH of the spray solution basic, we demonstrated selective detection and sequencing of phosphopeptides in complex mixtures at the low (<10) femtomole range (Zappcosta et al., 2002). Targeted on‐line LC–MS/MS may also be used for sequencing of phosphopeptides with high sensitivity whose molecular masses have been established. Stoichiometry of Modification: Phosphosite Occupancy

The extent to which each modification site is utilized often relates to its physiological relevance. For example, phosphoacceptor sites that are more heavily utilized are the first to be targeted for mutational analysis; this is a

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common way to assess the functional relevance of a phosphorylation site. A semiquantitative estimate of the extent to which each modification site is utilized can be obtained from the ratio of the ion abundances for the modified and unmodified forms of the peptides. For example, the ratio of MS response for phosphorylated to nonphosphorylated peptides in the full‐ scan, positive‐ion ESMS data obtained at acidic pH may be used to determine phosphorylation stoichiometry with an accuracy of circa 30% (Carr et al., 1996). The ability to provide site‐specific information about the extent of phosphate incorporation is a key advantage of the present approach relative to techniques that employ affinity‐based methods like immobilized metal‐ion affinity chromatography for selective fractionation of phosphopeptides. Finally, if low picomole amounts of intact phosphoprotein can be obtained in solution, the molecular weight of the intact protein is obtained by ESMS (or MALDI, especially in cases where ESMS is unsuccessful) so as to provide an overall check on the total number of moles of phosphate added to the protein and the relative distribution of protein molecules with different numbers of phosphate (Annan et al., 2001; Verma et al., 1997). Sensitivity

As noted earlier, the sensitivity for precursor‐ion scanning for selective detection and MS/MS for sequencing phosphopeptides by nanoESMS and nanoESMS/MS is in the low femtomole range (Carr et al., 1996; Zappacosta et al., 2002). However, the overall sensitivity of the method is currently limited by the first‐dimension LC–ESMS analysis. ESMS (at flow rates above circa 50 nl/min) behaves as if it were a concentration‐sensitive detector. Thus, smaller internal diameter HPLC columns will provide higher sensitivity analyses as the effective concentration for a given amount injected will be higher. These factors militate for use of as small an internal diameter (I.D.) column as possible. However, the off‐line precursor‐ion scanning method requires collection of fractions during LC–ESMS. Fraction collection, even by hand, requires a minimum of 1 to 2
l per minute of collection flow in practice. This collection flow rate, together with the flow rate required by the electrospray source to provide a stable spray, sets the minimum flow rate necessary through the HPLC column. This flow rate, in turn, limits the I.D. of the HPLC column that can be practically employed. Microionspray sources (the part downstream of splitter and fraction collection in Fig. 1) are now available and produce stable sprays at flow rates of 100 to 200 nl per minute. These flow rates permit use of either 180
m I. D. or 320
m I.D. HPLC columns, with column flows of 2 to 4
l per

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minute, which leaves sufficient flow for both MS detection and fraction collection. Using such a microionspray source flowing at circa 200 nl per minute and a 180‐
m I.D. HPLC column flowing at rates of 3 to 4
l per minute, the practical limit for phosphopeptide analysis is greater or equal to 50 fmoles on a column (Zappacosta et al., 2002). Conditions are described in the next section named Procedures. Procedures This section presents the experimental procedures involved in optimizing the chromatographic conditions, electrospray source, and mass analyzer for each step of the multidimensional analysis of phosphopeptides derived from phosphopeptides. Wherever possible, we, the authors of this chapter, have provided benchmark data using readily available standards so that interested readers can work through optimization of the methods on their specific instruments. Reagents. Phosphopeptides and their nonphosphorylated analogs (Table II) were obtained from the Protein and Carbohydrate Structure Facility, University of Michigan Medical School (Ann Arbor, MI, www. brcf.med.umich.edu). Other useful phosphopeptide standards are available from Bachem Bioscience (Philadelphia, PA, www.bachem.com). The phosphopeptide standards are dissolved in water, and 500 fmoles aliquots are dispensed into polypropylene microvials (Eppendorf, 0.5 ml size) and used fresh. Table II shows a test mixture of peptides and phosphopeptides that is used routinely to test system readiness for ‘‘real’’ samples. Peptides 9–13, Table II, were obtained as a mixture (Peptide Retention Standard, S1–S5) from Pierce Biotechnology (Rockford, IL). All other peptides were obtained from Peninsula Laboratories (Belmont, CA, www.penlabs.com) or Bachem Bioscience. Evaluation of Mass Spectrometer Performance in Negative‐Ion Mode. Conditions described here are specific to the acquisition of ES mass spectra on Sciex/Applied Biosystems quadrupole mass spectrometers (Concord, Ontario, Canada, www.appliedbiosystems.com). The tuning and calibration solution consists of a mixture of polypropylene glycol (PPG) 425, 1000, and 2000 (3  10–5 M, 1  10–4 M, and 2  10–4 M, respectively) in 50/50/0.1 water/methanol/formic acid (v/v/v), 1 mM NH4OAc. This mixture is used for tuning and calibrating the instrument in both positive‐ and negative‐ion modes. The m/z range 10 to 2400 is calibrated in the negative‐ion mode by multiple‐ion monitoring of the isotope clusters of six PPG ion signals and two trifluoroacetic acid (TFA)‐related ions, m/z 69 (CF3) and m/z 113 (CF3CO2). The TFA‐derived ions are present as background in any mass

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TABLE II PHOSPHORYLATED AND NON‐PHOSPHORYLATED PEPTIDES USED FOR OPTIMIZATION AND EVALUATION OF THE MULTIDIMENSIONAL PHOSPHOPEPTIDE ANALYSIS STRATEGY Peptide number 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Sequence

Mr (monoisotopic)

KRT(PO3H2)IRR (UOM 11) KRTIRR (UOM 10) KRPS(PO3H2)QRHGSKY (UOM 9) KRPSQRHGSKY (UOM 8) Ac‐RRLIEDAEY(PO3H2)AARG‐NH2 (UOM 7) Ac‐RRLIEDAEYAARG‐NH2 (UOM 6) TYSK LGG RGAGGLGLGK‐NH2 Ac‐RGGGGLGLGK‐NH2 Ac‐RGAGGLGLGK‐NH2 Ac‐RGVGGLGLGK‐NH2 Ac‐RGVVGLGLGK‐NH2 ISRPPGFSPFR MLF DRVYIHPFHLLVYS

908.5 828.5 1422.7 1342.7 1639.8 1559.8 497.2 245.1 883.5 911.5 925.5 953.6 995.6 1259.7 409.2 1757.9

spectrometer that is regularly exposed to TFA‐containing HPLC mobile phases (see following paragraphs). Mass spectra are recorded at instrument conditions sufficient to resolve the first two isotopes of anion m/z 991.7 (PPG þ HCO2) so that the valley between them is 55% of the height of the second isotope for the singly charged ion. At this resolution, singly charged ions can be distinguished from ions having two or more charges provided there are good ion statistics. In positive‐ion mode, resolution is adjusted such that the first two isotopes of cation m/z 906.7 (PPG/NHþ 4 ) are resolved with a valley between them of 40% of the height of the second isotope for the singly charged ion. At this resolution, it is possible to assign charge states of 1þ or 2þ and to distinguish these from more highly charged ions. Mass‐to‐charge ratio assignments for the measured peak tops can be closer either to the monoisotopic or to the average Mr depending on the charge state and the isotopic distribution. Calibration and instrument resolution is checked prior to LC–ESMS using the background TFA anions at m/z 69 and 113. Alternatively, if these ions are weak or absent, one can use TFA‐containing mobile phases and a high enough declustering potential to keep the m/z 69 and 113 ions of TFA from saturating the detector.

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Prior to LC–ESMS, the declustering potential (source and collision region prior to first mass analyzer, Fig. 1) used to produce the marker ions is determined by infusion of a 5 pmol/
l solution of phosphopeptide standard 3 (Table I) in 30% HPLC mobile phase B containing 0.02% TFA (see following paragraphs). These test conditions are also used to routinely check for best performance by adjusting spray position, sample flow rates, gas flows, voltage settings, and other such factors. The S/N for m/z 63 and 79 are simultaneously monitored in real time by SIM (200 ms dwell per ion) and the signal for each maximized by adjusting the declustering potential for each. On the Sciex API‐III, the optimal declustering voltages are circa 350 V for m/z 63 and 300 V for m/z 79 using TFA‐containing mobile phases; these voltages are 250 V and 200 V, respectively, when 0.2% formic acid without TFA is used. These numbers may exceed the highest voltage provided on the particular MS system used. In this case, an external programmable power supply is added as previously described (Huddleston et al., 1997). The appearance of the low‐mass region containing the marker ions is shown in comparison to the solvent background in Fig. 6. The signal‐to‐background ratio for m/z 63 and 79 is typically circa 20:1 under these conditions. HPLC Systems for Separation and Analysis of Phosphopeptides The overall sensitivity of the method is currently limited by the flow rate and sample concentration requirements of the first dimension LC– ESMS analysis. HPLC columns with internal diameters of 0.5 mm work well for phosphopeptides in the range of 5 pmoles and higher. Columns with internal dimensions of 180‐
m I.D. work well with phosphopeptide amounts in the 50 fmole to 5 pmole range. Procedures for these two levels of chromatography are given in the upcoming sections. While conventional 0.1% TFA‐containing mobile phases may be employed (for examples, see Fig. 7), sensitivity for phosphopeptide detection is increased approximately 2.5‐fold (as measured by signal‐to‐noise ratio) using a combination of 0.02% TFA and 0.1% formic acid. The degree to which the quality of the chromatographic separation is compromised can range from minimal to significant depending on the specific type of C18 employed. Therefore, columns should be tested with a mixture of standard phosphorylated and nonphosphorylated peptides (for example, see Table II and following text) prior to committing real samples for analysis. For maximum sensitivity of peptide analysis by the LC–ESMS positive ion, TFA may be eliminated altogether and replaced with 0.2% formic acid. However, no apparent benefit is seen for phosphate marker‐ion sensitivity when doing LC–ESMS SIM with 0.2% formic acid. Some chromatographic peak

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FIG. 6. (A) Typical appearance of the marker ion region for infusion of a 5‐pmol/
l solution of a phosphopeptide standard (peptide 3, Table I). (B) TFA‐containing mobile phases alone at a declustering potential of 350 V. On other instruments, one or more voltages may need to be adjusted to affect the extent of ‘‘in‐source’’ CID so as to obtain a similar spectrum. The S/N for m/z 63 and 79 are simultaneously monitored in real time by SIM (200 ms dwell per ion), and the signal for each is maximized. On the Applied Biosystems Sciex API‐III, the optimal declustering voltages are circa 350 V for m/z 63 and 300 V for m/z 79 using TFA‐containing mobile phases; the voltages become 250 V and 200 V, respectively, when 0.2% formic acid without TFA is used. The signal‐to‐background ratios for m/z 63 and 79 are typically circa 20:1 under these conditions.

integrity is sacrificed under these conditions although certain column packings (such as the PepMap C18 phase, LC‐Packings) can perform very well. LC–ESMS Using 0.5 mm I.D. HPLC Columns. Any gradient HPLC system capable of forming reproducible gradients at flow rates of 20
l per minute (either directly from the pumps or using an appropriate preinjector split flow system) with TFA‐containing mobile phases may be employed. Desalting and preconcentration of dilute samples are accomplished as part of the sample injection step using a microprecolumn of C18 (PepMap phase, 1 mm  5 mm LC‐Packings, San Francisco, CA) as the sample loop of the injector. The HPLC column is a 0.5 mm  150 mm C18 (‘‘Magic’’ ˚ pore‐size particles, Michrom BioResources, Auburn, phase, 5
m, 200‐A

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FIG. 7. Evaluation of overall system performance in negative‐ion LC–ESMS mode. (A) UV chromatographic detection at 214 nm. (B) Selected‐ion monitoring for phosphopeptide marker ions. A mixture of model peptides (for peptide identities, see Table II) at 20 pmoles was injected onto a 0.5‐mm I.D. HPLC column using 0.1% TFA in the mobile phases (i.e., the least sensitive conditions; see text). Only the two phosphopeptides (Table II, lines 1 and 3) in the mixture are observed in the marker‐ion trace (as shown in B). Marker ions at m/z 63 and 79 were monitored using a 0.2‐Da window, 0.04‐Da step, and a 100‐msec dwell, making a 1.1 sec scan rate. The ion‐spray capillary voltage was operated at 3.7 kV with the interface plate at 700 V. The nebulization gas consisted of zero‐grade compressed air at 48 psi. Nitrogen (99.999%) at a flow rate of 0.7 L/min was used as the ‘‘curtain’’ gas. Peptides 5 and 6 (Table II) were not included in the mixture analyzed.

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CA, www.michrom.com). The flow rate to the MS is reduced to 4 to 5
l per minute after the column and UV detector (see Fig. 1) by splitting the flow using a Valco (Houston, TX) tee with an appropriate length of fused silica acting as a restrictor on the fraction collection leg. A length of 50‐
m‐ I.D.‐fused silica goes from the tee to the tip of the ionspray capillary. The mobile phases used for gradient elution consist of (A) water/acetonitrile 98:2 (v/v) and (B) acetonitrile/water 90:10 (v/v), with both A and B containing either 0.1% TFA or a combination of 0.02% TFA and 0.1% formic acid by volume (see earlier note). Typical gradient conditions are 5% B to 50% B linearly in 30 min, then linearly to 95% B in 5 min, and then a hold for 10 min. Prior to injecting samples, the guard column is conditioned using water/acetonitrile 98:2 (v/v) containing 0.1% TFA regardless of acid type or concentration used in mobile phases. The samples are made acidic with solvent A and then loaded onto the guard column, which is washed with solvent A. On this HPLC system, it is necessary to ground the metal tee to prevent charge flow to the UV electronics that would cause unacceptable UV baseline noise during LCMS. LC–ESMS Using 180‐mm I.D. HPLC Columns. Peptide mixtures are preconcentrated onto a PepMap C18 trap cartridge (300
m  5 mm; LC Packings) used in place of the injector loop. The trap is conditioned with 0.1% TFA, and the sample is then loaded. Samples are dissolved in or diluted with the 0.1% TFA (with or without 2% CH3CN) prior to loading on the trap. Mobile phases are as noted previously, except that the concentration of TFA is reduced to 0.02%, and 0.1% formic acid is added to increase the sensitivity and improve electrospray stability without sacrificing chromatographic resolution. The sample is back‐flushed off the cartridge onto the column at 4
l per minute with a gradient from 0% to 50% B in 30 min and then from 50% to 95% B in 5 min with a 10‐min hold. The flow from HPLC pumps (530
l per minute) is reduced to 4
l per minute using a microflow splitter (Accurate Splitter, LC‐Packings). The 180‐
m I.D. HPLC column (LC‐Packings PepMap C18 capillary column, 15 cm long, 3
m particles) is fitted directly into the injector. The column outlet, a 25‐
m I.D. and 280‐
m‐O.D.‐fused silica transfer line, is connected to the nanoflow electrospray block/splitter using a Teflon sleeve connector and a short piece (15 cm) of fused silica with the same dimensions. A UV detector can be plumbed in‐line prior to MS if desired. Flow is split for simultaneous fraction collection and MS using the microvolume Valco tee insert of the block (0.15‐mm I.D. through‐hole, Valco Instruments, Houston, TX, www.vici‐store.com) to direct 0.6
l per minute to the 20‐
m‐I.D.‐fused silica (also tapered) ES tip (New Objective, Waltham, MA) and 3.4
l per minute to the fraction collection line for manual collection into polypropylene microtubes. The fraction collection line is composed of

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two pieces of tubing connected in series through a stainless steel union (63‐
m I.D. PEEK tubing from splitter and 50‐
m I.D.‐fused silica from union to fraction collector). The union is electrically grounded to prevent electrospray conditions at the fraction collector. Evaluating the Overall Performance of the LC–ESMS System. The overall performance of the LC–ESMS system is evaluated using both UV detection at 214 nm and SIM of the respective marker ions. For a 0.5 mm scale chromatography, a mixture of model peptides (Table II) at 20 pmoles, each is injected. Representative UV and selected‐ion current traces for this mixture using 0.1% TFA in the mobile phases (i.e., the least sensitive conditions) are shown Fig. 7 (mixture of all peptides except 5 and 6, Table II). All peptides in the mixture (with the frequent exception of Table II lines 7 and 8, which have weak UV response and elute at or near the injection void volume) produce peaks in the UV trace (Fig. 7A), whereas only the two phosphopeptides (Table II, lines 1 and 2) in the mixture are observed in the marker‐ion trace (Fig. 7B). Optimizing Nanoliter Flow Electrospray for Precursor‐Ion Scanning and Peptide Sequencing by MS/MS NanoES mass spectra are obtained using an articulated nanoelectrospray interface (Wilm and Mann, 1994a) and the ‘‘medium’’‐sized metal‐ coated capillary spray tips available commercially (Protana, Inc., Odense M, Denmark). Samples are loaded from the back of the electrospray capillary using an electrophoresis gel‐loading pipette tip (Eppendorf Geloader tip, Brinkmann Instruments, Westbury, NY, www.brinkmann.com). Prior to sample loading, the gel‐loading tips are washed with the solution in which the samples are dissolved. The dissolution phase (50
l) is loaded from the top of the pipette and then pushed out through the tip using pressure from a 1‐mL syringe. All plastic surfaces are washed, and freshly prepared solutions are used to remove compounds that may cause chemical noise in the mass spectra. The nanoelectrospray capillary tip is positioned in the ion source with the aid of the two cameras supplied with the setup from Protana. The optimal position in terms of S/N on the Sciex electrospray sources is approximately 1 to 2 mm directly in front of the orifice. To generate a stable signal and maintain a minimum flow rate (20–40 nl per minute), a positive air pressure on the sample in the capillary, of slightly more than atmospheric pressure, is usually required. The capillaries that work the best for the authors’ research require touching the capillary to the gate valve plate (voltages off and applying air pressure) to initiate sample flow. With sample flow, the electronics are then turned on before the final

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adjustment of moving the capillary toward the orifice. This last adjustment is made using the cameras and by monitoring the changes in the mass spectra from scan to scan in real time. Tuning and calibration for full‐scan data acquisition in both positive and negative ion nanoelectrospray modes are carried out as described earlier. A phosphopeptide standard is used to tune and optimize production and detection of m/z 79 (PO3 ) marker ion in precursor‐ion scanning mode. The resolution of Q3 is reduced to pass a 4 to 5 Da window around the precursor ion of interest to enhance precursor scan sensitivity. A further small gain in sensitivity is obtained by decreasing the resolution of Q1 to yield circa a 75% valley between singly charged ions. Depending on the instrument, all other important tuning parameters are checked, such as the quadrupole offsets (related to collision energy), lens settings, curtain gas (generally need to be reduced), and the collision gas pressure to maximize transmission of m/z 79. An argon:nitrogen (85:15) gas mix is used as the collision gas in Q2. Tuning and calibration for peptide sequencing by MS/MS is carried out by nanoelectrospray of glu‐fibrinopeptide B (Bachem Bioscience) at a concentration of 1 pmole/
l. On the Sciex triple quadrupole mass spectrometers, spectra are acquired over the desired mass range using a mass step of 1 Da with a dwell of 20 ms per mass step and a mass defect of 50 mmu per 100 Da. This approach produces analytically useful MS/MS data rapidly, allowing many product‐ion spectra to be acquired using a single 1‐
l loading of sample. The resolution of Q3 is adjusted to unresolved fragment‐ion isotope clusters, scattered across the mass range, so as to maximize signal. The instrument is calibrated so that for the product ions, the most abundant peak in a cluster consisting of three 1‐Da steps is the ‘‘monoisotopic peak.’’ This method is used routinely when acquiring product‐ion data at the femtomole level on precursor ions with 105 counts per second per scan or less. When sufficient amounts of peptide are available, or when isotope resolution is required, tuning and calibration in the product‐ion scan mode is performed using a mass step of 0.2 Da with isotopes clearly resolved. Optimizing Phosphopeptide Selective Detection by Precursor‐Ion Scanning and Sequencing by Nanoelectrospray MS/MS. Phosphopeptide selective detection by precursor‐ion scanning and sequencing by

FIG. 8. Optimizing phosphopeptide selective detection by precursor‐ion scanning. (A) Negative‐ion nanoelectrospray MS of a 100 fmole/
L solution of phosphopeptides 1, 3, and 5 (Table II) in basic water/methanol. (B) Precursor‐ion mass spectrum of m/z 79. (C) Full scan positive‐ion mass spectrum. See full text for details.

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FIG. 9. (A) Representative MS/MS data for phosphopeptide 5 (Table II and panel B) obtained by switching to positive‐ion mode during nanoelectrospray of the phosphopeptide‐ containing fraction in Fig. 8. The (M þ 3H)3þ of peptide 5 at m/z 547.8 was mass‐selected for sequencing by CID MS/MS. Spectra were acquired over the mass range 50–1850 using a mass step of 1 Da, with a dwell of 20 ms per mass step and a mass defect of 50 mmu per 100 Da. (B) Sequence of peptide 5 (Table II) indicating y‐ and b‐ions observed in MS/MS spectrum.

nanoelectrospray MS/MS are optimized using freshly prepared stock solutions of standard phosphopeptides (Figs. 8 and 9). The conditions are used to test detection limits and mass accuracy as well as to simulate conditions used to prepare a real sample for the same analyses. A dried aliquot containing 500 fmoles each of phosphopeptides 1, 3, and 5 (Table II) is diluted just prior to analysis to a final concentration of 100 fmole/
l using 5
l of 50:40:10 methanol/water/ammonium hydroxide (v/v/v). The basic solution is made up fresh by diluting a 30% ammonium hydroxide solution (Instra‐Analyzed reagent grade, J. T. Baker, Phillipsburg, NJ) with H2O to make a 20% NH4OH solution, which is then mixed 1:1 with methanol in a 1.5‐ml Eppendorf tube. Then 1.5
l of the standard solution is loaded into the nanoelectrospray capillary as described previously using prewashed gel‐loader pipette tips. The experimental sequence is as follows:

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1. Acquire full‐scan, negative‐ion data in a multichannel analyzer (MCA) mode (Fig. 8A). Monitor the data visually in real time, and terminate the acquisition when the overall ion statistics of the spectrum are satisfactory. To help prevent plugging of the capillary and, hence, deterioration of spray stability (which can happen during negative ion nanoelectrospray of basic solutions), use a slightly higher spray air pressure compared to that needed for positive ions. If plugging occurs, remove by increasing the air pressure to the capillary and by touching the tip to the entrance plate forcefully (with voltages off). When a droplet is observed, reduce the air pressure to normal operating levels. 2. Acquire m/z 79 precursor‐ion spectra in the negative‐ion, MCA mode to determine the molecular weights of the phosphopeptides (Fig. 8B). As previously noted, terminate the acquisition when the overall ion statistics of the spectrum are satisfactory (usually 3 to 6 min of data). 3. Acquire full‐scan, positive‐ion spectra in the MCA mode to identify species to select for CID tandem MS (Fig. 8C). Identify the ions corresponding to the (M þ 2H)2þ, (M þ 3H)3þ, and additional ions of the phosphopeptides of interest, bearing in mind that when comparing positive‐ion to negative‐ion data, there can be significant differences in signal intensities and in charge state distributions. If the parent ions of the phosphopeptides are not apparent above the background in the full‐scan, positive‐ion data, select for positive‐ion MS/MS the most abundant charge state observed in the m/z 79 precursor‐ion scan for each phosphopeptide as well as the next higher negative ion charge state (e.g., M 2H2 observed, select (M þ 2H)2þ and (M þ 2H)3þ for MS/MS). 4. Acquire product‐ion mass spectra of selected phosphopeptides and/ or nonphosphorylated peptides. If at anytime the overall signal appears unstable or weak, optimize the system before going on to the next experiment or before acquiring MCA data. ‘‘Real’’ samples or HPLC fractions containing less than or equal to 200 fmoles of peptide are split, and the half that is dried down is taken through the latter protocol. The sample is redissolved in 2–3
l basic solution, vortexed, and centrifuged; then 1–1.5
l is loaded into the nanoelectrospray capillary. Sample dissolved in basic solution should be used the same day. If the positive‐ion precursor(s) needed to acquire MS/MS data are weak but give a good response in negative‐ion mode, an aliquot of the sample may be taken to dryness and brought up in 35:65:5 water/methanol/formic acid (v/v/v) and reanalyzed. When acquiring data on samples that approach the limit of detection (less than 50 fmoles total), there may be little or no obvious peptide signals

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by negative ion. This makes it difficult to know if the flow rate and spray conditions are optimal. In such cases, phosphopeptide standard 3 can be added (as shown in Table II) as an internal standard to the sample that is dried down (50 fmole/
l when redissolved). A narrow m/z window around m/z 710.4, the (M – 2H)2 for this peptide, can be used to rapidly assess signal‐to‐noise and spray stability while optimizing air pressure (flow rate) and sprayer position. In precursor scan mode, it is also helpful to observe a phosphopeptide‐related ion that can be monitored over time to detect a no‐flow condition. References Allen, M., Anacleto, J., Bonner, R., Bonnici, P., Shushan, B., and Nuwaysir, L. (1997). Characterization of protein digest using novel mixed‐mode scanning with a single quadrupole instrument. Rapid Commun. Mass Spectrom. 11, 325–329. Annan, R. S., and Carr, S. A. (1997). The essential role of mass spectrometry in characterizing protein structure: Mapping post‐translational modifications. J. Protein Chem. 16, 391. Annan, R. S., Huddleston, M. J., Verma, R., Deshaies, R. J., and Carr, S. A. (2001). A multidimensional electrospray MS‐based approach to phosphopeptide mapping. Anal. Chem. 73, 393–404. Apweiler, R., Hermjakob, H., and Sharon, N. (1999). On the frequency of protein glycosylation as deduced from analysis of the Swiss–Prot Database. BBA 1473, 4–8. Azzam, R., Chen, S. L., Shou, W., Mah, A. S., Alexandru, G., Nasmyth, K., Annan, R. S., Carr, S. A., and Deshaies, R. J. (2004). Phosphorylation by cyclin B‐Cdk underlines releases of mitotic exit activator Cdc14 from the nucleolus. Science 305, 516–519. Bateman, R. H., Carruthers, R., Hoyes, J. B., Jones, C., Langridge, J. I., Millar, A., and Vissers, J. P. (2002). A novel precursor in discovery method on a hybrid quadrupole othrogonal acceleration time of flight (Q‐TOF) mass spectrometer for studying protein phosphorylation. J. Am. Soc. Mass Spectrom. 13, 792–803. Bean, M. F., Annan, R. S., Hemling, M. E., Mentzer, M., Huddleston, M. J., and Carr, S. A. (1995). ‘‘Techniques in Protein Chemistry’’ (J. W. Crabb, ed.) Vol. VI. Academic Press, San Diego, CA. Boyle, W. J., van der Geer, P., and Hunter, T. (1991). Phosphopeptide mapping and phosphoamino acid analysis by two‐dimensional separation on thin‐layer cellulose plates. Meth. Enzym. 201, 110. Carr, S. A., Huddleston, M. J., and Annan, R. S. (1996). Selective detection and sequencing of phosphopeptides at the femtomole level by mass spectrometry. Anal. Biochem. 239, 180–192. Carr, S. A., Huddleston, M. J., and Bean, M. F. (1993). Selective identification and differentiation of N‐and O‐linked oligosaccharides in glycoproteins by liquid chromatography–mass spectrometry. Prot. Sci. 2, 183–196. Chen, S. L., Huddleston, M. J., Wenying, S., Deshaies, R. J., Annan, R. S., and Carr, S. A. (2002). Mass spectrometry‐based methods for phosphorylation site mapping of hyperphosphorylated proteins applied to Net1, a regulator of exit from mitosis in yeast. Mol. Cell. Proteomics 1, 186–196.

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