Medical Applications of Mass Spectrometry || Mass spectrometry in proteomics

Medical Applications of Mass SpectrometryK. Vékey, A. Telekes and A. Vertes (editors)© 2008 Elsevier B.V. All rights reserved

173

Chapter 8

Mass spectrometry in proteomics

AKOS VERTES*

W. M. Keck Institute for Proteomics Technology and Applications, Department of Chemistry, The George Washington University, Washington, DC, USA

1. Introduction 1732. Methods in proteomics 176

2.1. Peptide mapping 1772.2. Peptide fragmentation 1792.3. Sequence tags 1822.4. De novo sequencing 1832.5. Electron capture and electron transfer dissociations 1842.6. Quantitative proteomics 1862.7. Higher order structures 1872.8. Mapping protein function 189

3. Outlook 190References 191

1. Introduction

In recent decades, rapidly expanding knowledge in molecular biology has providedthe biochemical framework for the functioning of all eukaryotic organisms. Thecore principles of this framework are based on three fundamental classes ofmolecules: nucleic acids, proteins, and metabolites. In a living organism a gene,coded in the DNA, is transcribed into an RNA molecule. Through processing, thenoncoding regions of the RNA are removed and a messenger RNA, mRNA, isspliced. Genes in the DNA are studied by genomics, whereas their expression inthe form of mRNA is explored by transcriptomics. The past 20 years witnessed the

*Tel.: �1-202-994-2717; Fax: �1-202-994-5873. E-mail: [email protected].

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sequencing of the human genome [1,2]; thus, discovering the genetic basis ofcertain diseases became feasible.

According to current estimates, there are approximately 25,000 genes in thehuman genome. Owing to alternative splicing and other mechanisms, the tran-scriptome of an organism is much more complex than the genome. As transcrip-tion and processing are influenced by the condition of the organism, disease statescan be reflected in expression level changes in the transcriptome. Analysis of thetranscribed mRNAs is typically carried out using DNA microarrays.

The second group of molecules, proteins, is produced through the ribosome-mediated translation of the mRNAs. Proteins serve as the general actors incarrying out most cell functions from motility to mitosis. The nature and activityof these functions are regulated by multitudes of posttranslational modifications,e.g., by acetylation, phosphorylation, or ubiquitination, of the proteins. Thesemodifications emerge as the main regulators of protein functions. Proteomics, avigorously developing field, is the systemic study of all proteins produced by anorganism.

Owing to posttranslational modifications, there are many more proteins thanmRNAs. It is estimated that approximately one million different proteins corre-spond to the �25,000 human genes. In addition, protein concentrations varygreatly in space, time, and expression level. Therefore, it is not sufficient to ascer-tain that a particular protein is present in the organism; the spatial and temporaldistributions of its concentration also have to be established. Spatial variations ofprotein expression in an organism are traditionally imaged using quantitativeautoradiography and fluorescent labeling methods, including tagging with greenfluorescent protein. These approaches, however, require the development oflabels for every individual protein. Therefore, their utility for high-throughputsystemic studies is very limited.

Importantly, the proteome can change in response to a disease. The alteredexpression levels can be used in diagnostics or form the basis of treatmentstrategies. Conventional methods of expression profiling were largely based ontwo-dimensional gel electrophoresis (2-DE). However, because of the limitedaccuracy, resolution, and specificity of this method, positive protein identificationhad to rely on additional forms of analysis. As a result of these complicatingfactors, proteomics presents an even greater challenge than genomics.

Some of the common objectives in proteomics include identification of pro-teins in a particular tissue or biological fluid (through peptide mapping, sequencetags, de novo sequencing, etc.), secondary, tertiary, or quaternary structure analy-sis of known proteins, function analysis through epitope mapping, quantitation ofprotein expression levels, and imaging of their distributions. The main methodused for protein identification and quantitation in proteomics is mass spectrometry.An introduction to the established methods of mass spectrometry in proteomics isthe subject of this chapter.

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Mass spectrometry in proteomics 175

Mass spectrometry is uniquely positioned among the large variety of analyticaltechniques to achieve the outlined objectives [3]. Chapter 6 presents a thoroughintroduction to the principles and instrumentation of mass spectrometry. Massspectrometric methods provide a better sensitivity, dynamics range, and selec-tivity than nuclear magnetic resonance (NMR) techniques. Mass spectra aremore specific and less complex than many forms of optical spectroscopy and,given the right ionization technique, they can provide structural information.With the discovery of electrospray ionization [4] (ESI) and matrix-assisted laserdesorption/ionization [5,6] (MALDI) in the late 1980s, the ion sources with thenecessary capabilities (no high mass limit and adjustable amount of fragmenta-tion) became available and the stage was set for the birth of proteomics. For theirrespective role in developing these enabling technologies, John Fenn and KoichiTanaka received the 2002 Nobel Prize in Chemistry [7].

The thirdclassofmolecules,metabolites, is adiversecollectionof typically smallerspecies (�1500 Da) that participate in cellular energy production and in the synthesisand degradation of macromolecules. The systematic study of the human metabolomehas started only recently. By early 2007 already over 2000 endogenous metaboliteshave been identified, quantitated, and catalogued [8]. There is clearly a large diversityfor this class of molecules; for example, the number of different metabolites in theplant kingdom is estimated to be �200,000. In addition to the endogenous metabo-lites, molecules introduced from the environment through nutrition or as drugs andtheir degradation products are also present in living organisms. A simplified view ofthe three major molecular classes, their hierarchy and interactions, and the corre-sponding disciplines devoted to their study are presented in Fig. 1.

Most biomedical samples contain thousands of biochemical components andthus are too complex even for mass spectrometry. Separation methods are needed toreduce this complexity by selecting smaller groups of components from the origi-nal specimens (see Chapter 5). The most commonly used separation methods in pro-teomics are affinity chromatography with its high selectivity, multidimensionaltechniques, such as 2-DE, and the combination of ion exchange (IEX) and high-performance liquid chromatography (HPLC). More recently, ion mobility spec-trometry was used to separate the polypeptide components before mass analysis.

These separation methods and especially their combinations with mass spectrom-etry are capable of producing data in large volumes. Curated archiving and interpre-tation of these data require sophisticated computational resources. Bioinformaticsaims to manage and mine the rapidly growing information from genomic, prote-omic, and metabolomic investigations including the discovery of reaction networks(see Chapter 10). There are numerous bioinformatics databases and tools availableon the Internet (e.g., http://www.ncbi.nlm.nih.gov/, http://www.expasy.ch/, andhttp://prospector.ucsf.edu/) and from commercial sources. The leading mass spec-trometer manufacturers integrate their data acquisition systems with these tools toprovide comprehensive solutions for proteomics research.

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2. Methods in proteomics

A common task in proteomic analysis is to identify a subset of proteins in a bio-medical sample. In principle, this can be accomplished through two differentroutes. The first, and most common, approach is to break down the proteins intopeptide segments of manageable size through enzymatic digestion and analyzethese building blocks using mass spectrometry. This is the so-called bottom-upapproach. The other, less common, method that relies on the analysis of intact pro-teins is the top-down approach. The top-down strategy requires high-performancemass spectrometers (e.g., ion cyclotron resonance, ICR, or orbitrap systems; seeChapter 6) with exceptional mass resolution and accuracy in combination withpowerful fragmentation techniques (such as electron capture dissociation, ECD) to

Fig. 1. Fundamental molecular classes in eukaryotic organisms and their interactions. Subdisciplinesdevoted to studying particular classes are shown on the perimeter. Genomics gives an unprecedentedglimpse into the DNA-based molecular design of life. Proteomics studies the translated and modifiedproteins, the main actors of cellular processes, and metabolomics tracks the dynamic changes in themakeup of small molecules brought about by inherent and environmental conditions. Ultimately, allthe molecular constituents, their interactions, and the knowledge of the entire reaction network areneeded to understand the basic processes in physiology.

DN A

RN A /mR N A

PR OTE INS

Tr an s c r ip t io n

Tran s lat io n

M etab o lit es

RE A CTI ON N ETW OR K S

ge n es

M o di fi ed pr o tein s

METABOLOMICS

PR OTE O M ICS

GENOMICS

Pro c essi n g

Mo d i f ic at io n s

TR A N SCRI PTO M ICS

DNA genes

RN A /mR N ARNA/mRNA

PROTEINS

Transcription

Translation

Metabolites

REACTION NETWORKS

M o di fi ed pr o tein sModified proteins

M E T A B O L O M I C S

PROTEOMICS

G E N O M I C S

Processing

Modifications

TRANSCRIPTOMICS

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enable sequence readout. In the following sections we briefly review the methodsused in the bottom-up and top-down approaches.

2.1. Peptide mapping

Peptide mapping takes advantage of the accurate mass measurement of uniqueprotein fragments produced by highly specific enzymatic digestion. Typically,trypsin is used due to its high fidelity in producing peptides in the size rangemost efficient for protein identification (400 � m/z � 5000). This range corre-sponds to �4–45 amino acid residues; thus, the corresponding peptides exhibitsufficient specificity. It also coincides with the m/z range where some commonmass analyzers (e.g., quadrupoles or ion traps) show their best performance.

Accurate mass measurement of the resulting peptides produces a set of m/zvalues that can be compared against a database of protein fragment masses[9,10]. These fragment databases are produced by the in silico digestion of all theentries in large protein databases. Several fragment databases are availableonline with the necessary searching tools. For example, as of January 9, 2007, theSwissProt protein database contained 252,616 entries. Their in silico digestionusing trypsin with a single missed cleavage allowed the production of 10,225,094peptides [11]. The search algorithm finds the proteins with enzymatic fragmentsin this database that match the measured peptide masses within a predefined tol-erance. Usually there are multiple possible matches and a review is required tofurther narrow the set and ultimately identify the unknown protein.

The efficiency of identification greatly depends on the performance of the massspectrometer. Most notably, the mass accuracy of the instrument, usually deter-mined by studying standards, has a dramatic effect. Clearly, the more accurate themeasured masses are the narrower is the set of proteins that produce fragmentswith masses within the tolerance. The number of peptides identified is correlatedwith the amino acid residue coverage of the original protein.

We demonstrate the mechanics of peptide mapping using the example ofthe �-chain of human hemoglobin. This protein is composed of 141 residues:VLSPADKTNVKAAWGKVGAHAGEYGAEALERMFLSFPTTKTYFPHFDLSHGSAQVKGHGKKVADALTNAVAHVDDMPNALSALSDLHAHKLRVDPVNFKLLSHCLLVTLAAHLPAEFTPAVHASLDKFLASVSTVLTSKYR.Unfragmented, it appearsin the MALDI mass spectrum as a protonated ion with a molecular weight of15,126.5 Da. This single number is clearly not specific enough to identify theprotein. There are many other proteins with the same m/z, e.g., the ones with anypermutation of the residues. Tryptic digestion with no missed cleavages producescharacteristic fragments in the 400 � m/z � 5000 range. Table 1 shows thesefragments, their location in the original protein molecule, and the correspondingcalculated monoisotopic and average masses.

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Table 1

Monoisotopic (mi) and average (av) peptide masses from tryptic digestion of human hemoglobin � chain [11]

m/z (mi) m/z (av) Start Sequence End

461.2718 461.5416 8 TNVK 11532.2878 532.6235 12 AAWGK 16729.4141 729.8564 1 VLSPADK 7818.4407 818.9537 93 VDPVNFK 991071.5543 1072.3195 32 MFLSFPTTK 401252.7147 1253.4903 128 FLASVSTVLTSK 1391529.7343 1530.6470 17 VGAHAGEYGAEALER 311833.8919 1835.0415 41 TYFPHFDLSHGSAQVK 562996.4894 2998.3651 62 VADALTNAVAHVDDMPNALSALSDLHAHK 903038.6496 3040.6206 100 LLSHCLLVTLAAHLPAEFTPAVHASLDK 127

In our first example we use a low-performance mass spectrometer. Assumingthat five peptides (m/z 729.86, 818.95, 1072.32, 1253.49, and 1530.65) appear inthe mass spectrum (e.g., as commonly observed, due to the ion-suppressioneffect we do not detect all tryptic peptides), the average masses are determinedwith 2000 ppm mass accuracy, and the search in the SwissProt database isrestricted to the proteins of Homo sapiens, the MS-Fit searching tool of ProteinProspector [11] finds 114 entries that are more or less consistent with this data.The relevant section of the mass spectrum is shown in Fig. 2. Note that in thisexample no impurities complicate the spectrum.

Fig. 2. Five fragment masses determined from the mass spectrum of the mock unknown protein(human hemoglobin � chain) tryptic digest form the basis of peptide mapping. The mass-to-chargeratio is labeled m/z on the horizontal axis.

500 1000 1500 20000

20

40

60

80

1001530.65

1253.49

818.95

729.86

Inte

nsity

m/z

1072.32

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The top-ranked hit is human hemoglobin � subunit with all five massesmatched, but with only 35.5% coverage (in light gray below).

The other proteins on the list showed fewer number of matching peptides or lowerdegree of coverage.

There are several ways to increase the fidelity of protein identification. Chiefamong them are to use better performing instrumentation [12] (nowadays a typicalhigh-performance mass spectrometer can achieve �5–10 ppm mass accuracy) andto identify more peptides. Improving the mass accuracy to 50 ppm for the sameset of m/z values does not increase the coverage, but it reduces the number of hitsfrom 114 to a single one, human hemoglobin � subunit.

Increasing the number of peptides used in the search to 10 (m/z 461.54, 532.62,729.86, 818.95, 1072.32, 1253.49, 1530.65, 1835.04, 2998.37, and 3040.62) with-out improving mass accuracy (keeping it at 2000 ppm) actually increases thenumber of hits in the search to 1151, but the coverage of the top scoring humanhemoglobin � subunit increases to 93.6%.

Enhanced mass accuracy (50 ppm) for this set of peptides reduces the numberof hits to one, the human hemoglobin � subunit, with 93.6% coverage. Thus, theright sample preparation and ionization method in combination with speciesinformation and reasonable instrument performance enabled us to identify a singleprotein in a database of over 250,000 entries.

2.2. Peptide fragmentation

Peptide mapping does not require any knowledge about the primary structure of theprotein or of its fragments. Owing to peptide fragmentation, however, parts of theprimary structure might become known from the mass spectra. The spontaneousfragmentation of peptides is relatively slow; it mostly takes place in the postsourceregion of the mass spectrometer. In time-of-flight instruments equipped with anion reflector, the ions produced by postsource decay (PSD) become observablein the mass spectrum at appropriate reflector voltage settings. This gives rise topeptide-sequencing capabilities [13].

More energetic ionization methods (in-source decay, ISD) or collisions withinert (collision-activated dissociation, CAD, also known as collision-induced

1 VLSPADKTNVKAAWGKVGAHAGEYGAEALERMFLSFPTTKTYFPHFDLSHGSAQVKGHGKKVADALTNAVAHVDDMPNAL81SALSDLHAHKLRVDPVNFKLLSHCLLVTLAAHLPAEFTPAVHASLDKFLASVSTVLTSKYR

1 VLSPADKTNVKAAWGKVGAHAGEYGAEALERMFLSFPTTKTYFPHFDLSHGSAQVKGHGKKVADALTNAVAHVDDMPNAL81SALSDLHAHKLRVDPVNFKLLSHCLLVTLAAHLPAEFTPAVHASLDKFLASVSTVLTSKYR

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dissociation, CID) or reacting species (ECD and electron transfer dissociation,ETD) also produce structural data. In these techniques, fragmentation of thepeptide backbone is induced through increasing the internal energy of the ions(ISD and CAD) or through ion chemistry (ECD and ETD). Thus, the presence ofparticular fragments in the spectrum is the function of the different ionizationmethods, e.g., MALDI and ESI, and more recently desorption/ionization on sili-con [14] (DIOS) and laser-induced silicon microcolumn arrays [15] (LISMA) aswell as instrument types (TOF, ion trap, ICR, etc.). There is more control overfragmentation patterns in tandem mass spectrometers (e.g., MS/MS and MSn),where the primary ion internal energy can be adjusted by, for example, CAD.

Depending on the actual bond that breaks in the peptide backbone (C–C, C–N,or N–C) and on the partitioning of the charge on the resulting fragments (amino orcarboxyl side fragment), there are six major fragment types. Their nomenclaturefor a pentapeptide is shown below.

Other less common fragmentation pathways, e.g., resulting in internal fragmentsor neutral loss ions, are not discussed here. As an example we can look at the neu-ropeptide leucine enkephalin, which has a sequence of YGGFL and a protonatedmonoisotopic mass of m/z 556.28. The fragmentation of this ion in a collision cellthrough CAD might produce a tandem mass spectrum similar to the one in Fig. 3.

H2NNH

HN

O

O

O

NH

HN

O

O

OHR1

R2

R3

R4

R5

a1 b1 c1 a2 b2 c2 a3 b3 c3 a4 b4 c4

x4 y4 z4 x3 y3 z3 x2 y2 z2 x1 y1 z1

Fig. 3. Fragmentation of the protonated leucine enkephalin molecular ion via CAD in a tandem massspectrometer.

0 100 200 300 400 500 6000

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LFGGG

a1

136.09

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221.10

425.17556.29

336.20

278.12

Inte

nsity

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279.16

[M+H]+b4

y3

y2

b3

Y

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In this simplified case the identity of amino acid residues in the peptide canbe inferred from the mass differences of successive peaks by comparing themwith the known masses of the amino acids. In real-world samples, the presence ofother ions and the absence of certain fragments make this task fairly com-plex. Comparison of the measured m/z values in the spectrum with the calculatedfragment masses in Table 2 enables the assignment of the peaks.

In addition to a1 and the molecular ion, parts of the bn and yn series are presentin Fig. 3. The sequence can be read as YGGFL. Note that the y series reads thesequence from right to left, whereas the b series reports it from left to right.Coincidentally, the mass difference between b2 and b3 and between y2 and y3 iden-tify the same residue.

Changing the internal energy of the ions through CAD can reveal more aboutthe primary structure. This can be induced by changing the collision energy ofthe primary ions or by changing the collision gas pressure in the tandem massspectrometer [16]. With the emergence of new laser desorption/ionizationplatforms based on nanostructured silicon, simpler instrumentation can alsoyield similar data. Fig. 4 shows the spectrum of a vasodilator peptide,bradykinin (RPPGFSPFR), as a function of relative laser intensity. At low laserpower the molecular ion dominates the spectrum. This can be advantageous incomplex mixtures, where the molecular weights of the different components canbe identified. Increasing the laser power from 95 to 145 relative value resultedin enhanced structure-specific fragmentation. Although the entire primary struc-ture cannot be inferred from this spectrum, the identity of the N-terminalresidues is revealed.

Even in the case of unmodified residues, entire peptide sequences are rarelyrevealed by fragment spectra induced by CAD. The task is even more complexwhen posttranslational modifications are present. Phosphorylation, for example, isprevalent due to its role in signal transduction and in the regulation of proteinfunction. In eukaryotic cells, as much as 30% of the proteins can be phosphory-lated. Histone protein functions are believed to be regulated by acetylation,phosphorylation, methylation, and ubiquitination. These modifications play animportant part in fundamental biological functions, e.g., gene silencing. Identifying

Table 2

Monoisotopic masses of major fragment ions for leucine enkephalin

N-terminal ions C-terminal ions

N 1 2 3 4 N 4 3 2 1

an 136.08 193.10 250.12 397.19 xn 419.19 362.17 305.15 158.08bn – 221.09 278.11 425.18 yn 393.21 336.19 279.17 132.10cn – 238.12 295.14 442.21 zn 377.19 320.17 263.15 116.08

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the modified residues by mass spectrometry requires comprehensive fragmentationof the protein domains of interest [17].

2.3. Sequence tags

Although comprehensive sequence information on protein domains is not avail-able on most instruments, shorter segments are often revealed by PSD, CAD, orother techniques. The concept of a sequence tag is based on using the partialsequence of a peptide digestion product, usually composed of a few residues, incombination with the masses of the adjoining N- and C-terminal fragments to effi-ciently search protein databases for the identity of unknown proteins [18,19].

For example, let us assume that we find three b series fragment ions, m/z 908.4,1021.5, and 1108.5 in the CAD spectrum from the tryptic digest of the humanhemoglobin � subunit that belong to the peptide parent ion with m/z 1833.9 (seeFig. 5). This is the peptide between residues 41 and 56 in Table 1.

The mass differences in the b series reveal the presence of L/I followed by S inthe sequence. This information is sufficient to attempt a sequence tag search.Searching the SwissProt database for H. sapiens proteins by entering m/z 1833.9for the parent ion and 1108.5, 1021.5, and 908.4 for the b series fragments in theMS-Seq searching tool of Protein Prospector [11] turns up a single protein, humanhemoglobin � subunit with primary accession number P69905.

Fig. 4. Laser desorption/ionization of 1 pmol of bradykinin from a LISMA surface produces increas-ing amount of structure-specific fragmentation as the relative laser power increases.

0

100

80

60

40

20

a5 [M+H]+

c6

-NH

3

-NH

3

a6

y8

y7

y6 Laser Power145

135

120

95

Mass/Charge600 800 1000 1200 1400

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The initial 252,616 entries in the database are reduced to 1328 by the parentmass filter. Using the three fragment masses the number of matching proteins forall species is 80. At this point, all the hits are related to the hemoglobin � subunit.Introducing the information on the species produces a single hit. Note, however,that the sequence coverage of the protein is only 11.3%. This limitation curtailsthe value of sequence tag identifications in the presence of multiple posttransla-tional modifications.

2.4. De novo sequencing

We have seen powerful methods to identify proteins in a sample based on massspectra and information from large protein databases. These strategies require thatthe protein of interest exists in the database. Protein databases contain informationthat was originally produced by traditional Edman sequencing or by meticulousmass spectrometric methods commonly known as de novo sequencing. Theseapproaches are necessary if the protein of interest is undescribed or substantiallymodified. Although both Edman degradation and tandem mass spectrometry canprovide sequences with acceptable accuracy, recently mass spectrometry seems tohave come out on top due to its dramatically higher throughput and better sensitivity.

There are two major approaches to de novo sequencing by mass spectrometry.The first one is based on a number of empirical rules obtained by observing typi-cal peptide fragmentation schemes [20]. Current versions of this approach rely oncomputerized expert systems that are built on the dozens of empirical rules andfactors. These include general observations on the prevalence of certain fragmentsin spectra produced by the used fragmentation methods and in typical instruments.For example, CAD is known to produce predominantly y- and b-type ions. There

Fig. 5. MS/MS mass spectrum reveals the partial sequence of a tryptic peptide from the humanhemoglobin � subunit. This information is sufficient to successfully perform a sequence tag searchand identify the protein. L�I stands for leucine or isoleucine, whereas Xm and Xn denote unknownsequences.

600 800 1000 1200 1400 1600 1800 20000

20

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100

XnSL/I

Xm

b7

908.4

1108.5

1833.9

1021.5

Inte

nsity

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[M+H]+

b9

b8

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are other rules related to neutral losses and relative intensities of spectral features,and on determining the presence of certain amino acid residues based on immoniumions formed by the combination of a- and y-type cleavages.

Furthermore, it is imperative to recognize the ambiguities resulting from iden-tical or indistinguishable masses (isobars). Common examples are leucine andisoleucine or lysine and glutamine with only 0.0364 Da mass difference for thelatter. Similar problems arise when dipeptide masses are isobaric with singleamino acids or with other dipeptides. These challenges can only be resolved byusing instrumentation of sufficiently high mass accuracy or by residue-specificchemical derivatization. The expert systems can successfully call sequences ofover 10 residues, including posttranslational modifications.

The other approach to de novo sequencing is based on a systematic treatmentof tandem mass spectrometric data and database search. An excellent descriptionof these methods is available in Chapter 9; thus, we refrain from the detailed dis-cussion here.

As the exploration of the human proteome advances from better known proteinsto more and more obscure ones, the significance of de novo sequencing as the pri-mary source of information is likely to grow. Similarly, the identification of splicevariants, mutations, and modifications calls for increasing number of de novoinvestigations.

2.5. Electron capture and electron transfer dissociations

As we pointed out in Section 2.2, the y and b series ions induced by CAD, or othermethods of gradually producing elevated internal energy, rarely reveal even themajority of the residues. For example, only �25% of the 76-residue ubiquitinsequence can be identified through CAD. This incomplete information leaves theprimary structure unresolved.

The problem with gradually energizing these polypeptide ions seems to be therapid redistribution of internal energy, which leads to the preferential breakage ofa low number of the weakest bonds. After several years of searching for a methodto produce more complete fragmentation, great improvement was achieved byreacting low-energy electrons and the multiply charged peptide ions, [M � nH]n�,produced by ESI [21]. This method, termed electron capture dissociation (ECD),produced a radical cation, [M � nH](n�1)�•, that in turn rapidly dissociated intoc and z series ions with the degree of fragmentation approaching 80% and withoutpreference to bond strength [22]. An alternative fragmentation pathway can alsoproduce a- and y-type ions. Not only the fragments in ECD provide higher cover-age than CAD but also the information in the two methods is complementary.Thus, a mass spectrometric method to sequence large peptides and small proteins intheir entirety became feasible. This also presented a realistic approach to top-down

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proteomics, i.e., to the analysis of intact protein components without enzymaticcleavage.

A comparison of the fragments produced by CAD and ECD shows the advan-tages of the latter in phosphopeptide analysis. Quadruply charged molecular ionsof a 28-mer phosphopeptide, atrial natriuretic peptide substrate (ANPS),SLRRSpSCFGGRIDRIGAQSGLGCNSFRY, were fragmented by the two methods[23]. The resulting patterns showed incomplete fragmentation (20 of the 27 pep-tide bonds) for CAD with significant loss of the phosphorylation site information.The corresponding ECD spectrum showed complete sequence coverage and thelocation of the phosphorylation site (see Fig. 6).

ETD takes the concept of ECD to the next level [24]. Owing to the conditionsrequired to trap the thermalized electrons that produce ECD, it can only be per-formed in ICR mass spectrometers. These systems are large and expensive;thus, this technical requirement limits the availability of ECD to a relativelysmall number of laboratories. To make the benefits of ECD available on morecommon instrumentation (e.g., ion traps), heavier electron-donating agents, i.e.,low electron affinity anions are needed that can be trapped together with thepeptide ions. Anthracene [24] and fluoranthene [17] radical anions as ETDagents were shown to generate primarily c- and z-type ions from multiplycharged large peptide, phosphopeptide, and small protein species. Like ECD,ETD produces close to complete fragmentation and thus enables the elucidationof primary structures.

Fig. 6. Comparison of fragmentation patterns for a 28-mer phosphopeptide. In the top pattern pro-duced by CAD, incomplete backbone fragmentation and extensive phosphate loss (denoted by �P)can be observed. Numbers indicate the charge carried by a particular fragment. Complete sequencereadout and identification of the phosphorylation site are straightforward for ECD (bottom pattern).(Reprinted with permission from: Shi, S.D.H., Hemling, M.E., Carr, S.A., Horn, D.M., Lindh, I. andMcLafferty, F.W., Anal. Chem., 73, 19–22 (2001). Copyright 2001. American Chemical Society.)

S

Collision activated dissociation

b-P

b-P

b

1,2

1,2 1,21

L R R RS S C F G G2,3

I D R I G A Q S G GL C N S F R Y

S L R R RS S C F G G I D R I G A Q S G GL C N S F R Y

2

2

1,2

4

4

111111111222

22 2

2

*

*

y

44y-P 3

2

2222

2 2

22222 2 2

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222

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1111

11 1

1 1,2

34

3 32 2

22 2

22

22

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222 2

2,31

11

11

11

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abc

yz

Electron capture dissociation

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A fascinating application of ETD is the analysis of posttranslational modifica-tions on the H3.1 histone tail [17]. Histones are proteins found in chromatin andserve as the core for DNA coils. It is hypothesized that particular combinations ofposttranslational modifications, e.g., acetylation, methylation, and phosphoryla-tion, on the histone tail at the amino terminus, form a code that is directly involvedin gene regulation [25]. A 50-mer peptide from the amino terminus of H3.1 wasisolated from human cells and subjected to ETD by fluoranthene anions in an ion-trap mass spectrometer. To reduce the charge state of the produced fragments,proton-transfer reactions were performed by benzoic acid anions. The resultingmass spectra showed a unique pattern of methylation sites that showed systematicvariations during chromatographic separation. Correlating these modificationswith gene expression data is instrumental in understanding the role of histonemodifications in gene regulation.

2.6. Quantitative proteomics

Unlike nucleic acids, proteins in an organism are present at very different concen-tration levels. Thus, it is not sufficient to demonstrate that a particular protein ispresent; we also need to know its concentration. From the high-concentrationglobulins in blood to the low-copy-number proteins that are represented by only afew molecules per cell, there is an enormous dynamic range. This presents achallenge to the utilized analytical methods because of the potential interferences,especially when quantitating the proteins at low concentration. For example, thehigh-abundance proteins can compete in the ionization process and suppressthe ion formation from the low-level species. This ion suppression effect is quitecommon in MALDI and ESI ion sources.

Common approaches to minimize these problems include extensive separationbefore mass spectrometric analysis. Typical separation protocols consist of anorthogonal combination of affinity chromatography, 2-DE, IEX, HPLC, and ionmobility techniques. If these steps can reduce the sample complexity to a singlecomponent, the signal from the separation method (e.g., chromatographic peakarea) can be used for quantitation. Frequently this is not achievable or verifiable.Relative quantitation in these instances can be performed by stable isotope labelingmethods.

A common example of relative quantitation is used in comparative pro-teomics. For example, to uncover the differences in protein makeup and concen-tration levels between the healthy state and a particular disease (e.g., proteinexpression in normal vs. HIV-infected cells [26]), stable isotope labeling can beapplied to one or the other. A frequently used variant of this approach is the iso-tope-coded affinity tag (ICAT) method [27]. Fig. 7 shows how an ICAT reagentis used to tag the cysteine residues of a peptide, human insulin chain B in thisexample. First, the reactive end of the ICAT reagent covalently attaches to the

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cysteine residues through thiol chemistry. One form of the reagent, d0-ICAT withno deuterium atoms, can be used to label the sample from the healthy source,whereas the other, d8-ICAT with eight hydrogens in the linker replaced by deu-terium, can designate the diseased sample. As a result the tagged peptides in thehealthy and the diseased samples will exhibit a mass difference of 8 or its multi-ples depending on the number Cys residues. In the next step the two samples arecombined and the biotin end of the ICAT reagent is used to separate the taggedpeptides through affinity capture with avidin. This results in significantlyreduced sample complexity.

The mass spectrum of the captured mixture exhibits the peptide peaks as dou-blets with a mass shift of 8 or, in case of multiple cysteine residues, its multiplesbetween the normal and the diseased sample. The abundance ratios of these dou-blets characterize the relative quantity of a particular protein in the two samples.As both the d0- and the d8-tagged components are in the same matrix and differonly in isotope composition, the relative peak intensities are a true reflection of theprotein level changes in disease. The ICAT method is limited to cysteine-containingproteins, but other tagging protocols (e.g., through proteolytic 18O labeling) arebeing developed to eliminate this restriction [28].

2.7. Higher order structures

The efficiency of mass spectrometric methods in determining primary proteinstructure naturally leads to the question of their utility to characterize secondary,tertiary, and quaternary structures as well as the formation of noncovalent com-plexes. The success of mass spectrometry in approaching these problems is morelimited. For example, there are some legitimate questions about the correspondence

Fig. 7. Cysteine residues of human hemoglobin chain B are tagged with ICAT reagent. The codingof the linker, X, can be hydrogen (d0-ICAT) for the normal sample and deuterium (d8-ICAT) for thediseased sample.

NH NH

S

O

H N

O

O

X

XX

X

OO

X

X H N

X

XI

O

Phe Val Gln His Leu Cys Gly Ser His Leu Val Ala Leu Tyr Leu Val Cys Gly Glu ArgGlyPhePheTyrThrProLysThr

ICAT ICAT

BIOTIN ISOTOPE CODED LINKER REACTION WITH Cys

GluAsn

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of these structures between the native solution state and their ionized form in thegas phase. Are there significant structure changes as the molecule is ionized? Howdoes the structure change when the molecule loses its solvation shell duringvolatilization?

It was noticed in the early 1990s that conformation changes, for example, dueto pH changes, resulted in altered charge-state distributions in ESI spectra [29].Although there are literature reports on successful deconvolution of these chargedistributions to assess the relative weight of coexisting conformations (bothsecondary and tertiary structures) [30], the method is far from being routinelyapplicable. This approach hinges on the differences in the available protonationsites in a multiply charged ion in its folded and stretched conformations. When themolecule is folded, only the protonation sites exposed on its surface are accessi-ble, whereas in its stretched conformation, at least in principle, all amenable sitesshould be ionized. Thus, unfolding of the molecule is reflected in a charge statedistribution shifted to lower m/z values. Under limited conditions, folding andunfolding kinetics can also be followed measuring the time dependence of chargestate distributions following a chemical perturbation (e.g., pH change) of thesystem.

Another method to study higher order protein structure is hydrogen–deuteriumexchange [31]. When a protein molecule is dissolved in deuterium oxide, D2O(“heavy water”), deuterium atoms start to exchange their accessible hydrogens. Theresulting mass difference in the mass spectrum of the protein and its digestion prod-ucts can reveal which part of the folded protein is accessible for the D2O molecules.

Carbon-bound hydrogens do not exchange, whereas the exchange on the sidechains of certain residues (e.g., Arg, Asn, Cys, and Trp) is very fast, essentiallyimmediate on the timescale of the experiment. The exchange rate of amide hydro-gens on the peptide backbone is between the two extremes and can be used toexplore protein structure. The exchange rates of these amide hydrogens alsodepend on the pH and the temperature, so adjusting these parameters gives addi-tional control. A typical experiment starts with exchanging the solvent to D2O atpH 7.0 and at room temperature. This initiates the exchange of accessible amidehydrogens at the surface of the protein to deuterium. Changing the pH to 2.5 andthe temperature to 0ºC arrests the exchange process and gives enough time to per-form enzymatic digestion (typically with pepsin) followed by HPLC separationand mass spectrometry. A complicating factor is back exchange that can replacethe deuterium already in the peptide fragments with hydrogen. This effect can beestimated and the results corrected for it.

The hydrogen–deuterium exchange method can be used to study secondary, ter-tiary, and even quaternary structures. Amide hydrogens in the hydrophobic core ofthe protein or at the interface of attached subunits are less accessible for theexchange reaction. Studying the kinetics of the exchange can reveal unfolding

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dynamics and the association of partners in noncovalent complexes. The advantagesof mass spectrometry over competing techniques used in combination with hydrogen–deuterium exchange (e.g., NMR) are the very low amount of protein required(�1 nmol) and the ability to tackle very large proteins including an entire 2.5 MDaribosome and its subunits [32]. In addition, protein mixtures can also be studied withmass spectrometry.

Molecular recognition and noncovalent complexes are at the core of reactionnetworks in biology. Molecular complexes are often associated with the prolif-eration of disease (see, for example, the Tax-associated complexes in humanT-cell leukemia type 1, HTLV-1 [33]). Along with other competing techniques(e.g., surface plasmon resonance), mass spectrometry can be successfully usedto detect noncovalent complex formation. The corresponding ions can be pres-ent in both MALDI [34] and ESI [35] spectra, although the latter is used moreoften. A wide variety of protein–protein interactions as well as protein interac-tions with other species (nucleotides, carbohydrates, etc.) have been studied.The spectra can reveal the components of the complex and in some cases theassociation constant.

2.8. Mapping protein function

From the biomedical perspective, structural and kinetic studies are incompletewithout determining the function of the protein. In the discussion of posttransla-tional modifications and noncovalent complexes, we have already indicated theirimportant role in regulating the role a protein plays. In addition to biological func-tion, protein-based drug and vaccine design also requires the elucidation of theirmechanism of action. From heart disease to cancer, there are many examples inthis volume showing the variety of implicated proteins [36]. Conversely, structuraldiscrepancies in proteins are shown to result in disease states.

An interesting example of using mass spectrometry to unravel protein functionis epitope mapping. In broad terms, an epitope is the binding site on the surface ofa protein that attaches to another molecule; for example, to a monoclonal antibody.There are two general strategies to identify the epitope. In the first one the proteinis attached to the antibody. Then, proteolytic digestion is performed that removesthe nonattached parts of the protein. Mass spectrometric analysis of the removedfragments and the segment retained on the antibody can reveal the epitope. In thesecond strategy, the studied protein is digested first and the resulting mixture isaffinity separated by the monoclonal antibody. The protein fragment that containsthe epitope is preferentially captured [37].

Even if the participating protein segments are discontinuous, epitopes can alsobe identified by hydrogen–deuterium exchange. The components of the noncova-lent complex are deuterated in D2O environment and allowed to react. When the

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solvent is changed to water, the amide deuterium atoms on the exposed surface ofthe formed complex are exchanged with hydrogen. The epitope region, however, isnot affected because it is not exposed. Displacing the protein from the complex fol-lowed by pepsin digestion produces peptides that are deuterated at the epitope. Theresulting mass differences can be detected by MALDI mass spectrometry [38].

Although epitope mapping can contribute an important piece of the puzzle,identifying protein function requires a more complex approach. The available sub-set of genetic, X-ray diffraction, NMR, and mass spectrometric data has to be con-sidered in its entirety to shed light on the function of newly discovered proteins [39].Often similarity searches in genomic and proteomic databases can provide an ini-tial hypothesis based on homology with proteins of known function. For example,proteomic analysis of the Torpedo californica electric organ, a large-scale modelfor the neuromuscular junction, identified 11 human open reading frames codingfor proteins of unknown function [40]. When similarity is not found, high-resolution structures (X-ray and NMR data) as well as mass spectrometric study ofnoncovalent complexes can be used to identify active sites and infer the possiblefunctions of the protein.

3. Outlook

In the past few years we have witnessed the explosive growth in the field ofproteomics. During this period, proteomics has captured the attention of academia,government, and industry alike. At the universities, new courses are being intro-duced to teach the related technologies and applications for the emerging genera-tion of biomedical professionals. Government funding in developed countries isincreasingly available in the proteomics field. The landscape of mass spectrometermanufacturing has been reordered by the technological demands of proteomics;reagent, diagnostic, and pharmaceutical vendors gear up to take advantage of thenew market opportunities.

This dramatic new focus was already clearly discernable from the presentationsat the 2002 symposium organized by the U.S. National Academies, Defining theMandate of Proteomics in the Post-Genomics Era as well as from the launchingof three dedicated journals, Journal of Proteome Research, Molecular andCellular Proteomics, and Proteomics. Learning from the lessons of the HumanGenome Project, it was clear from the outset that international efforts had to becoordinated. In 2001 an international consortium, the Human ProteomeOrganization (HUPO), was launched to facilitate several initiatives, includingprojects related to the proteomes of the liver, brain, and plasma, to the develop-ment of proteomics standards, and to mouse models of human disease [41].

Despite its short history, the field of proteomics has already started to differen-tiate. Beyond the basic distinction between methods, including instrumentation and

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bioinformatics, and applications to biomedical problems of interest, more or lesscoherent subfields are beginning to appear. Among them are proteomics within thesubdisciplines of biology (e.g., proteomics in cell biology and microbiology, plantproteomics, and animal proteomics) as well as proteomics in the medical fields(e.g., the proteomics of a certain organ or disease). As the discovery of disease-related protein biomarkers continues, proteomics is poised to become an everydaytool in clinical diagnostics and serve as a basis for new therapies.

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Medical Applications of Mass Spectrometry || Mass spectrometry in proteomics