High-Resolution Analytical Tools for Quantitative Peptidomics

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High-Resolution Analytical Tools for Quantitative Peptidomics Sayani Dasgupta and Lloyd D. Fricker Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, New York, USA

Chapter Outline 1. Introduction 2. Absolute Quantification 3. Relative Quantification 3.1. Label-Free Quantification

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3.2. Metabolic Labeling 3.3. Chemical Labeling 3.4. Proteolytic Labeling 4. Concluding Remarks References

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INTRODUCTION

Peptidomics is the identification and elucidation of the peptide content or “peptidome” of a living organism, tissue, or cell. Peptides play important roles in regulating biological processes, functioning in cell–cell signaling as peptide hormones and neuropeptides. In addition, peptides have been proposed to function in intracellular regulation of protein–protein interactions. Quantitative peptidomics is a powerful tool to compare peptide profiles between different physiological states. The term “peptidomics” was first used more than 10 years ago (1,2), and the technique has since been applied to the identification and analysis of hundreds of peptides from a variety of tissue samples. Peptidomics has been used to characterize bioactive peptides regulating complex mechanisms such as circadian rhythm, feeding, and behavior in rodents and in lower organisms (3–7). Quantitative peptidomics has been used to measure changes in peptide content of specific brain regions and/or cells (8–11) under disease conditions or in response to drug administration (12,13). Quantitative peptidomic approaches have also been used to identify the substrate specificity of known enzymes as well as pathways regulating the synthesis Comprehensive Analytical Chemistry, Vol. 63. http://dx.doi.org/10.1016/B978-0-444-62651-6.00014-3 Copyright © 2014 Elsevier B.V. All rights reserved.

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and degradation of bioactive peptides (14–17). In addition, peptidomic tools have been applied to the discovery of novel antimicrobial peptides and development of potential biomarkers for various diseases (18–22). Peptidomics and proteomics share similarities in terms of principle and tools required for identification and analysis, although there are some distinct differences between these techniques. Endogenous peptides are often cleavage products of larger precursor proteins, and the goal of peptidomics is to identify the precise form of each peptide. Thus, samples for peptidomics are not enzymatically digested prior to analysis. The absence of enzymatic processing allows the identification of the peptide in its native form, including posttranslational modifications (23,24). In contrast, the goal of most proteomic studies is to identify proteins present in the sample, without concern for the precise molecular form of each protein. To identify the proteins, they are typically digested with trypsin to produce peptides that can then be sequenced (25). Another difference between the overall goal of proteomics and peptidomics analyses has to do with the tendency for different cleavage products of the same peptide precursor to have distinct physiological functions. For example, limited cleavage of prodynorphin generates peptides with highest affinity for kappa opioid receptors while more complete processing of prodynorphin generates smaller peptides with highest affinity for delta opioid receptors. Thus, it is important to identify all peptides produced from a single protein (26–28). This is not a requirement for proteomic studies, since it is possible to ascertain the identity of a protein from a small number of tryptic peptides. Another difference between proteomics and peptidomics is with sample handling and the issue of protein degradation, which needs to be minimized for peptidomic studies. This is usually not a disadvantage for proteomics since the identification of a protein is not hindered by the breakdown of a small fraction of the protein. However, because bioactive peptides are typically present in tissues at low levels, even a small amount of degradation of highly abundant proteins can overwhelm the signal from the endogenous peptides. Therefore, it is essential to eliminate postmortem protein degradation so that the endogenous bioactive peptides can be detected. To counter the problem of protein degradation, the proteases in the tissue need to be heat-inactivated prior to processing of the tissue for peptidomics (10,29–31). In this review, we focus on the various techniques for quantification of peptides/proteins and their advantages and disadvantages with respect to their application in peptidomics.

2 ABSOLUTE QUANTIFICATION Determination of absolute quantities of peptide levels mostly employ the stable isotope dilution (SID) methodology coupled to mass spectrometric analysis. SID involves the use of stable isotope-labeled standards wherein quantification is performed by comparing the mass spectrometric profiles of analytes to their corresponding labeled standards. It is important to choose the internal standard carefully to accurately measure absolute quantities from signal intensities.

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Different compounds have variable ionization intensities, which are further affected by factors such as chromatographic retention times and suppression by compounds present in the analyte fluid and sample matrix, so it is essential to use isotope-labeled internal standards, which are physicochemically identical to the molecule of interest rather than structural analogs, to normalize for effects that can lead to erroneous quantification. Isotope-labeled standards also account for any differences arising out of sample preparation and absorptive loss due to selective binding on surfaces during chromatography (32). Precision and sensitivity of quantitation has been greatly advanced by the use of selected reaction monitoring (SRM). SRM allows for selective monitoring of specific molecular ions with known fragmentation properties. SRM is performed on triple-quadrupole mass spectrometers, where the first mass analyzer selects ions with m/z corresponding to that of the peptide of interest for collision-induced dissociation. Out of the various fragmentation ions produced, only specific ions uniquely derived from the peptide precursor of interest are further selected for detection and measurement in the second analyzer. SRM enables the detection of several precursor-fragment ion pairs within a set of specific retention times. This facilitates the measurement of many target peptides simultaneously, which is known as multiple reaction monitoring (MRM) (33). A combination of SID and SRM has been applied to the absolute quantification of proteins and posttranslational modifications directly from cell lysates through measuring their tryptic peptide levels. A tryptic digest of a protein mixture is supplemented with a known amount of isotopic-labeled peptide derived from the target protein precursor and subjected to SRM mass spectrometry. Similar retention times and ionization properties enable matching of native peptides to their appropriate labeled standard and quantification from comparing their mass spectral intensities, leading to absolute measurement of endogenous protein levels (34,35). Absolute quantification is limited to the measurement of selected peptides/proteins for which internal standards are available, unlike the relative quantitative peptidomic approaches that can detect unknowns as well as known peptides.

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RELATIVE QUANTIFICATION

Relative quantification methods provide information on the identity of peptides/proteins in a sample as well as their levels expressed in amounts relative to each other. Some of these methods rely on label-free strategies while others incorporate stable isotopes into one or more of the samples, allowing them to be combined and analyzed together. In both cases, the strength of the signal for each peptide is a reflection of the amount of peptide present in the sample, providing quantitative information. In the following section, both label-free and isotopic label approaches are described in detail. In both approaches, the point of the experiment is to compare the relative levels of peptides in two or more distinct groups;

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Figure 1 shows an example in which an experimental group is compared to a control group. The group can consist of animals, tissues, cells, or other sources of proteins/peptides. Due to normal biological variation, it is important to include replicates of the members of each group. Figure 1 shows four biological replicates for a total of eight samples that need to be compared. In both label-free and isotopic label approaches, the peptides are first extracted from the tissues or cells. In the label-free approach, the samples are directly analyzed by liquid chromatography/mass spectrometry (LC/MS). While this would seem to save time (due to the simpler sample preparation), each biological replicate is usually analyzed in at least three technical replicates, requiring much more time for LC/MS and subsequent analysis than the isotopic label approaches. The technical replicates are first averaged to determine the levels of peptides in each of the biological samples. Then, the individual values for each biological replicate are averaged together and standard error of the mean (or standard deviation) is calculated. Finally, the average of the experimental group is compared to the average of the control group and statistical testing performed to determine if there is a significant difference between any of the peptides in the two groups. The isotopic approach involves the introduction of stable isotopes so that the peptides in each group can be distinguished by mass. The isotopes chosen are usually 15N, 13C, 18O, or 2H (or a combination of these). In all cases, the isotopes are stable (i.e., nonradioactive) so that they are safe to work with and will not contaminate the mass spectrometer. In some cases, the stable isotope is incorporated into the protein/peptide in cell culture, or even in living animals. In other cases, the isotopic label is added in a chemical reaction once the peptides are extracted from tissue/cells. The example shown in Figure 1 is for a multiplex label, in which eight different isotopic forms can be used. In this example, all eight biological samples can be included in a single LC/MS run, thus saving considerable instrument time compared to the label-free approach. Data analysis for the isotopic label approaches involve averaging levels among the biological replicates and then comparing the experimental and control values, as for the label-free techniques.

3.1 Label-Free Quantification As the name suggests, label-free quantification does not require the incorporation of an isotopic label into peptides for quantitation of relative concentrations. Each sample is separately prepared, subjected to individual LC/MS runs, and quantification is performed through analysis of their mass spectra. There are two approaches used for label-free quantification. The first approach is based on the assumption that peptide/protein levels follow a linear relationship with area under the curve of the ion spectra. This method involves quantitation of relative levels by comparing the peak intensity of ions from multiple peptides. But experimental variations in sample preparation and

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FIGURE 1 Quantitative proteomics/peptidomics using label-free and isotopic labeling approaches. In a typical experiment comparing two conditions (control vs. experimental groups), biological replicates are performed; in this example, four biological replicates are indicated. In both approaches, proteins/peptides are first extracted from the samples. For the label-free approach, the samples are directly analyzed on liquid chromatography/mass spectrometry (LC/MS). However, label-free approaches typically run technical replicates of each sample; three technical replicates are indicated here, resulting in 24 total LC/MS runs. Peptides are quantified by the spectral counts or another method, the technical replicates are averaged, and then the biological replicates are averaged and the standard error of the mean (SEM) or standard deviation calculated. The experimental and control groups are compared using statistical tests. For the isotopic labeling approach, the samples are individually labeled with a distinct chemical tag that differs in mass due to the presence of stable isotopes (2H, 13C, 15N, or others). Following the labeling (and quenching of unreacted labels), the samples are pooled and analyzed on LC/MS. With iTRAQ or other reagents that are available in eight different isotopic forms, all of the biological replicates in this example can be tested in a single LC/MS run. The data are analyzed by comparing the peak intensity of each of the isotopic forms, the biological replicates are averaged together, and control and experimental groups compared, as above.

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sample injection procedures can result in run-to-run variations in peak intensities, retention times, and m/z values, which greatly reduce the accuracy of data interpretation (36). To avoid imprecise results, label-free quantification based on peak intensity requires several preprocessing steps. Peak detection by baseline correction and smoothing of raw mass spectra is followed by application of data-normalization algorithms to account for variations in peak intensities between different runs. Subsequent to peak detection, consecutively eluting peaks from the same peptide are merged into features in a process termed peak/feature extraction. Features derived from identical peptides in different samples are aligned with respect to retention times and m/z ratios, following which relative quantitation is performed by comparison of the peak intensities (37). The second approach for label-free quantification involves identifying the total number of MS/MS spectra from a particular protein, based on the direct correlation between the abundance of a protein and the number of fragmentation spectra of peptides derived from that protein. After peptide identification, quantification is carried out by calculating peak areas from extracted ion chromatograms of the corresponding peaks in the LC/MS (38,39). This method has its limitations as well; low-abundance proteins often go undetected, deficiencies in ion-trapping capacity and ion efficiency of the mass spectrometer lead to saturation of spectral counts and consequent imprecise quantitation of high-abundance proteins. Often, the linear relationship between protein abundance and the number of spectra is affected by peptides that do not ionize or fragment efficiently. Peptide sharing or detection of peptides that are common to multiple proteins further complicates identification and quantification by the spectral counting method (40). Furthermore, a large number of falsepositives are generated in analysis of extracted ion chromatograms while applying spectral counting to peptidomics (41). Label-free quantitation has several advantages over isotopic label approaches, but also some disadvantages. Isotopic labels are expensive, so if there is no cost for each LC/MS run, label-free quantification is an economically viable option. However, because of the larger number of LC/MS runs that need to be performed with label-free approaches than with isotopic label approaches (especially multiplex labels), the label-free approaches can be considerably more expensive if there is a cost for each LC/MS run. Also, the number of samples that can be compared by labeling techniques are limited by the number of available labels, whereas there is no such restriction in label-free methods, which can accommodate more samples per experiment (although with an increase in the number of LC/MS runs that need to be performed and analyzed in each experiment). Label-free methods have a higher range for quantitation, making them useful to detect large variations in peptide/protein levels. However, the accuracy of label-free approaches is lower than that of isotopic label approaches, making it difficult to detect small changes in peptide levels using the label-free methods. The total number of

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peptides/proteins identified is typically higher in label-free approaches than with isotopic label approaches because the latter has greater mass spectral complexity due to differently labeled versions of the same peptide (38,42). Overall, there are advantages and disadvantages to each approach, and selection of the optimal approach depends on the scope of the experiment.

3.2

Metabolic Labeling

Incorporation of a stable isotopic tag into proteins/peptides in metabolically active cells was first described to quantify protein abundance in yeast (43). Wild-type and mutant cell populations were grown in media containing the naturally abundant isotopes of nitrogen and enriched in 15N, respectively, followed by trypsin digestion and LC-ESI-MS/MS analysis to identify and quantify relative phosphopeptide levels in both populations (43). SILAC (stable isotope labeling by amino acids in cell culture) was later described for the in vivo incorporation of specific amino acids in mammalian cells. The experimental cell population is grown in media containing an isotopic form of an essential amino acid. For proteomics, the amino acids used for incorporation are usually arginine and lysine because these are cleavage sites for trypsin, thus providing an isotopic tag on the C-terminus of every tryptic fragment. Apart from mass difference, the labeled and unlabeled forms of the same peptide in the experimental and control cell populations are chemically identical, which enables their relative quantitation by comparing the peak intensities through standard mass spectrometric techniques. SILAC does not require any peptide labeling steps and confers the advantage of almost complete incorporation of the labeled amino acids, thus precluding any differences in labeling efficiencies between the two samples. In addition, it is very useful to quantify small changes in protein/peptide levels. But not all cells are suitable for SILAC; the requirement for cells to be grown in dialyzed serum makes it difficult to be adopted for those cell lines, which do not grow well under this condition (44,45). A distinct disadvantage of SILAC also arises from the metabolic conversion of [13C6]- and [13C6,15N4]-labeled arginine to proline, which leads to inaccurate estimation of isotopically labeled proline containing peptides. A method based on using the [15N4]-arginine in combination with the “light” form of lysine and [13C6,15N4]-arginine with the [13C6,15N2]-lysine allows for internal control of arginine conversion since both [15N1]-proline and [13C5,15N1]proline will be formed at the same rate under both conditions (46). Unlike mammalian cells, which cannot synthesize all amino acids, SILAC is not well suited to studying the protein/peptide levels of microorganisms, most of which are prototrophic for all amino acids. Application of SILAC in such cases is mostly restricted to auxotrophic strains, which renders the technique ineffective for the proteome analysis of many commercially important microbes. Native SILAC (nSILAC), a recently developed modification of

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the existing technique, has been shown to be useful for the analysis of protein/peptide levels in microorganisms. nSILAC relies on the downregulation of lysine biosynthesis enzymes in the presence of exogenous lysine, thus leading to incorporation of isotopically labeled “heavy” lysine present in the media (47). SILAC is difficult for studies on rodents due to the expense of isotopic label and the need for lengthy exposure times to ensure adequate incorporation of the label into the proteome. Despite the high cost, SILAC had been performed for some specific analyses of protein turnover in rodents (48,49). Nearly all of the published SILAC studies have been for studies on proteomics, although it is technically possible to use this technique for studies on endogenous peptides. One peptidomic study used SILAC to investigate the production of peptide hormones in cell culture (50).

3.3 Chemical Labeling Stable isotopic labels can be introduced into reactive groups present in peptides/proteins through chemical modifications. For peptidomics, the most commonly targeted reactive group is the free amine present on the N-terminus of most peptides and also on the side-chain of lysine residues. For proteomics, free amines have been used as well as the side-chain of cysteine residues. In this section, we review different chemical labeling strategies and their applicability in peptidomics. There are several advantages of chemical labeling over the metabolic labeling approach. First, a wide range of samples can be analyzed, without the need to first culture cells in a special medium. Animal tissues can be used for the analysis, which would be prohibitively expensive using metabolic labeling (and impossible for human tissue). Second, some isotopic tags allow for multiple samples to be compared in the same LC/MS run. This allows for several different replicates to be analyzed at the same time, providing a greater estimate of the reproducibility of the data. The ideal chemical isotopic tag is one that will react with every protein or peptide present in the sample and be reactive enough that the protein/peptide becomes fully labeled. However, the intrinsic reactivity of the compound should not be so high that it degrades upon storage under proper conditions. Also, the ideal tag will be stable upon mass spectrometry and not readily break apart under standard conditions. The ideal chemical tag will either be commercially available or relatively easy to synthesize, and the price should be reasonable. At a minimum, the tag needs to be available in a heavy and light form, but ideally would be available in a range of masses so that multiple replicates can be combined. It is also important that peptides labeled with the heavy and light forms of the tag coelute from HPLC; otherwise, the quantitation is less accurate when

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comparing different spectra (due to ion suppression). Some of the isotopic tags come close to the properties of the ideal reagent, although all have their drawbacks (described below).

3.3.1 ICAT Isotope-coded affinity tags (ICATs) were first used for the quantitative comparison of protein expression in Saccharomyces cerevisiae. The three basic components of an ICAT reagent are a thiol-reactive group (usually iodoacetamide) to specifically target cysteine residues present in the peptide/protein, an isotopically “light” or “heavy” linker, and a biotin moiety. The protein samples are treated with the ICAT prior to enzymatic digestion, following which the peptides are purified on an avidin affinity column. ICAT was originally synthesized as a deuterated tag containing none or eight deuteriums and yielded peptide pairs differing by a mass shift of 8 Da on the mass spectrum. One disadvantage is that deuterium interacts with hydrophobic surfaces weakly as compared to hydrogen, causing deuterated peptides to elute earlier on a reverse-phase column (deuterium isotope effects). This difference in retention times of the labeled peptide pairs can lead to inaccurate quantitation (51). A modified version of the ICAT (clCAT) was developed later, featuring an acid-cleavable biotin tag and substitution of 1H with 12C and 2H and 13C in the isotopically “light” and “heavy” forms, respectively. Peptides thus labeled, coelute during MS, which results in more accurate identification. Removal of the biotin tag after peptide purification leads to relatively cleaner fragmentation spectra, thus further improving the accuracy and rate of identification (52). Several other versions of the ICAT labeling technique have emerged in recent years to widen its potential (e.g., for the quantification of phosphopeptides). Phosphoprotein isotope-coded affinity tag (PhIAT) is based on the labeling of phosphorylated serine and threonine residues by deuterated and nondeuterated ethanedithiol. Peptide complexes are subsequently enriched on an avidin affinity column and analyzed by mass spectrometry to detect, identify, and quantify differences in phosphopeptide levels (53,54). IGOT (isotope-coded glycosylation-site-specific tagging) combines affinity chromatography and proteolytic labeling (further discussed in this review) for the identification of N-linked glycoproteins. Consequent to isolation of glycopeptides on a lectin affinity column, 18O tag is stably incorporated into glycosylation sites by N-glycosidase action. The 18O peptides are further identified by mass spectrometry (54). ICAT has also been adapted for absolute quantification purposes. The metal-coded affinity tag method introduces different lanthanide ions instead of stable isotope into peptides, which enables their absolute quantification by inductively coupled plasma mass spectrometry (ICP-MS) (55,56). Appending a fluorophore to an ICAT reagent also facilitates absolute quantification by fluorescence detection. Fluorescence isotope-coded affinity tag, as this

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method has been named, further aids in purification of labeled peptides by immunoprecipitation with anti-fluorescein antibody (57). Being a relatively rare residue, labeling of cysteines by ICAT reduces complexity in the sample mixture (51), making it a convenient methodology for proteomics. The same property makes it inefficient for peptidomic analyses, since most endogenous peptides do not contain cysteine residues.

3.3.2 iTRAQ/TMT/mTRAQ Since most peptides contain a free N-terminus and one or more internal lysines, labeling amine groups will permit detection of a large number of peptides, thus making it a useful strategy for peptidomics. Isobaric tags for relative and absolute quantification (iTRAQ) and tandem mass tags (TMTs) use the tandem mass spectra of labels identical in mass and structure (also known as isobars/isotopomers) for the quantification of peptides/proteins. The labels comprise a reporter group, balance group, and an amine-reactive N-hydroxysuccinimide group, which derivatizes N-terminal amine and side-chain amines of lysine residues present in peptides/proteins. Differential isotopic composition of carbon, nitrogen, and oxygen atoms keeps the combined mass of the reporter and balance groups constant throughout the isobaric tags. Introduction of isobaric tags produces isotopically labeled peptides with identical mass and chromatographic properties. These peptides coelute in MS, but the reporter group dissociates as a signature ion with distinct m/z in the MS/MS mode, thus allowing for the identification and subsequent quantification of the parent peptides (58,59). The first iTRAQ reagents used a fourfold multiplex strategy (reporter ion m/z from 114 to 117) that allowed multiplexing of experimental samples, a property further enhanced by the development of eight isobaric variants (reporter ion m/z from 113 to 121) (60). TMTs were initially developed as a 2-plex technique, and have been recently expanded to a 6-plex (reporter ion m/z from 126–131) and 8-plex version (with two variants of the TMT127 and TMT129 labels) for improved multiplexing ability (61,62). Although iTRAQ 4-plex has been shown to have higher peptide identification rates over iTRAQ 8-plex as well as TMT 6-plex (63), the ability of the latter to facilitate simultaneous analysis of a large number of samples cannot be overlooked, especially while performing high-throughput studies. Because the iTRAQ/TMT-labeled peptides have the same nominal mass when analyzed by the first stage MS, there is lower mass spectral complexity as well as increased sensitivity. This allows for the identification of a greater number of ions by MS/MS (58). In spite of these advantages, isobaric labeling suffers from certain disadvantages. Peptide identification and quantification require tandem mass spectra of peptides, which are usually fewer in number as compared to the total number of spectra generated during mass spectrometry. Thus, not all peptides that are seen on MS can be quantified. Furthermore, the iTRAQ/TMT reagents are expensive and difficult to synthesize.

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There are several examples of peptidomic studies using iTRAQ labeling. One study identified substrates of prolyl peptidases in mouse models, and another explored the effect of diurnal variation on the endogenous peptide composition of human parotid saliva (64,65). Recently, mTRAQ (MRM tags for relative and absolute quantification) was developed by Applied Biosystems as a proteomic tool to be used specifically in conjunction with MRM. Derived from iTRAQ reagents, these aminereactive nonisobaric compounds were initially synthesized as light and heavy versions, differing only in their composition of 13C and 15N atoms. Peptides labeled with the light form (mass 141 Da) and heavy form (mass 145 Da, identical to iTRAQ 117) differ by 4 Da. This generates unique MRM transitions (precursor/product pair) for the same peptide labeled with the two mTRAQ reagents, which causes labeled peptide pairs to be monitored independently within a complex mixture. MRM combined with mTRAQ allows for detection of peptides present in low amounts in complex mixtures while avoiding the use of deuterated standards for absolute quantitation. This method has been successfully applied for absolute quantitation in proteomic studies by using one label to tag the synthetic peptide standard and the other to label the protein mixture (66). mTRAQ has also been used for the relative quantification of proteins, particularly biomarkers (67–69). Recent development of a triplex version of mTRAQ (mass 149 Da) permits multiplexing of samples (70). Though mTRAQ has been used to validate methods for enrichment of endogenous peptides in human plasma and their identification (71), it has not been widely used for peptidomics.

3.3.3 TMAB The trimethylammonium butyrate (TMAB) isotopic tags, which are synthesized as amine-reactive N-hydroxysuccinimide derivatives, were first described by Regnier and coworkers (72). The chemical name of the TMAB-N-hydroxysuccinimide reagent (TMAB-NHS) is [3-(2,5-dioxopyrrolidin-1-yloxycarbonyl)propyl] trimethylammonium chloride. The label incorporates a quaternary amine so that the labeled peptide retains a positive charge in place of the free amine (unlike acetylation and related labels described in the next section). TMAB-NHS was initially synthesized in a light and heavy form, containing all hydrogens (D0) and nine deuteriums (D9), respectively. The multiplexing capability of TMAB labels was greatly improved upon synthesis of additional D3, D6, and 13C, D9 (referred to as D12-TMAB) forms (Figure 2). Although TMAB-NHS labels are not commercially available, they are relatively easy to synthesize from inexpensive reagents (73). Peptides labeled with TMAB coelute, presumably due to clustering of deuterium in a polar environment, which reduces its affinity for the reverse-phase column matrix and hence minimizes isotope effects (72). The 3 Da mass difference

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FIGURE 2 Quantitative peptidomics using trimethylammoniumbutyrate (TMAB) labels. (A) Chemical structures of the labeling reagents, which are reactive N-hydroxysuccinimide (NHS) esters that combine with free amines to form an amide bond. The methyl groups on the amine contain various ratios of hydrogen/deuterium (1H, or 2H) and 12C/13C. The form containing 12C and only H was initially named H9-TMAB, and renamed D0-TMAB. The form with one deuterium on each of the three methyl groups is D3-TMAB, the form with two deuteriums on each methyl group is D6-TMAB, and the form with three deuteriums on each methyl group is D9-TMAB. The form with nine total deuteriums and the 12C substituted with 13C is named D12-TMAB because the mass is 12 Da heavier than the D0 form. (B) Up to five different samples can be labeled with the various TMAB tags, as shown. (C and D) Representative results from an experiment of HEK293T cells treated for 1 h with 50 nM bortezomib, a proteasome inhibitor used as a drug to treat multiple myeloma and other cancers. In this experiment, two replicates of control cells were labeled with D3 and D9TMAB, while three replicates of drug-treated cells were labeled with D0, D6, and D12-TMAB. The peptide in panel C was not altered by bortezomib treatment; this peptide was subsequently identified by tandem MS analysis as Ac-AAKVFESIGKFGLALA (an N-terminal fragment of prohibitin) labeled with two TMAB tags. The peptide in panel D was greatly elevated by bortezomib treatment; this peptide was identified as Ac-AGQAFRKFLPLFD (an N-terminal fragment of heat shock 10 kDa protein 1, also known as chaperonin 10) labeled with one TMAB tag.

between the labeled peptides allows most to be sufficiently resolved on MS (Figure 2C), although some peptides are not adequately resolved and can be difficult to quantify (Figure 2D). This problem can be eliminated if only three tags are used: D0, D6, and D12, providing a 6 Da difference between each isotopic form.

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Another disadvantage of using TMAB labels is that the isotopic tag disintegrates during collision-induced dissociation in MS/MS mode, causing neutral loss of trimethylamine (74). This can complicate peptide identification if the investigator is not aware of this; however, if a high enough energy is used for collision-induced dissociation, the dissociation of the trimethylamine group is complete and so this is not a major problem for MS/MS identification. The Mascot computer program includes TMAB labels and neutral loss of trimethylamine in their list of modifications (under the name GIST) for the D0, D3, D6, and D9-TMAB tags (although not the D12-TMAB tag). TMAB labels have been extensively used in peptidomics, especially for the study of neuropeptides. Using mice models deficient in endopeptidases such as prohormone convertase 1/3 (PC1/3) and carboxypeptidase E (CPE), TMAB peptidomics has been useful for exploring the role of these enzymes in neuropeptide biosynthesis as well as the identification and quantitation of several neuropeptides. Numerous peptides derived from secretory pathway proteins (classical neuropeptides) have been identified in the pituitary as well as brain regions of CPE-deficient (Cpefat/fat) and PC1/3-deficient mice (75–79). Since neuropeptides function as cell–cell signaling molecules and regulate a number of biological processes, it is of interest to quantify their levels under specific physiological conditions. Quantitative peptidomics using TMAB labeling has been applied to measuring changes in relative peptide levels in the brain under conditions of food deprivation, exercise, and administration of drugs such as cocaine and morphine (12,80,81). Differential isotopic labeling with TMAB has been employed for the study of neuropeptides derived from cytosolic proteins, particularly hemopressin, which is a hemoglobin-derived peptide and a known CB1 cannabinoid receptor antagonist. Novel forms of hemopressin with CB1 agonist activity were discovered and their regulation in Cpefat/fat mice was examined (82,83). Apart from identifying these “nonclassical” neuropeptides (28), TMAB-labelingbased peptidomic approaches have unveiled a possible third category of peptides, which elicit their biological effects within the cell in which they are produced. Many such “intracellular peptides” have been detected and the enzymatic pathway for their synthesis has been elucidated in human cell lines (84–86). In addition, TMAB tags have been used to determine substrate specificity of enzymes (16).

3.3.4 Amine Labeling by Other Reagents A relatively inexpensive method of labeling peptides is to tag their free amines with isotopically light and heavy forms of commercially available reagents. One of the first reagents to be tested was acetic anhydride, which can be purchased with the normal isotopic distribution of atoms (i.e., containing mostly hydrogen) or with six atoms of deuterium (although only three are

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incorporated into the labeled peptide). An advantage of these labels is that they are relatively inexpensive and commercially available. A disadvantage is that the heavy and light forms of the labeled peptides generally do not completely coelute from the LC due to deuterium isotope effects, which can result in imprecise quantification (14,87). Another problem is that free amines are turned into amides, and unless there is another positively charged amino acid within the peptide (Arg or His), the peptide will not be positively charged. Because most LC/MS is performed in the positive-ion mode, neutral peptides will not be detected. Other amine-labeling reagents with similar problems are N-acetoxysuccinimide (available with three deuteriums) and propionic anhydride (available with five deuteriums). Both of these labels suffer from isotopic effect of deuterium on chromatographic retention times, similar to that observed with acetic anhydride (72,88). In addition, the free amines of the N-termini and lysine side-chains are covered by the modifying reagent, rendering the peptide neutral if there is no internal arginine or histidine. Free amines of peptides/proteins can also be labeled with succinic anhydride, available with natural distribution of atoms (i.e., mostly hydrogen and 12 C) or with four deuteriums such that succinylation causes a mass difference of 4 Da per tag incorporated into the peptide (9,89,90). In addition, succinic acid with four 13C atoms can be purchased and easily converted into the anhydride. The use of 13C eliminates the problem with non-coelution observed with the deuterated and nondeuterated forms of acetylated peptides. However, the extra expense of the 13C-labeled reagent is not necessary because the deuterated and nondeuterated forms of the succinylated peptides show fairly good coelution. Labeling with succinic anhydride labeling thus provides for more accurate quantitation than acetic anhydride (72,91). Isotopic labeling by succinylation has been used for the identification and quantitation of N-linked glycoproteins as well. After proteolytic removal of nonglycosylated peptides, glycopeptides are isotopically labeled by succinic anhydride. Labeled peptides are then cleaved by peptide-N-glycosidase and analyzed by MS (92). One of the main disadvantages of succinic anhydride labeling is that peptides without arginine or histidine residues go undetected in the positive-ion mode, as with acetylated peptides. Furthermore, succinylated peptides have a reduced signal intensity in the mass spectrum compared to acetylated peptides, possibly due to the increased number of acidic groups in the labeled peptide (74). Another strategy for stable isotope labeling of N-terminus and lysine side-chain amines is reductive amination by nondeuterated and deuterated formaldehyde. Dimethylation of peptides occurs by the formation of a Schiff base, followed by reduction with cyanoborohydride. Peptides thus labeled differ by a mass of 4 Da for each labeled pair. The labeled peak pairs show minimum isotope effects on reverse-phase separation and hence coelute (93). Dimethylation using formaldehyde with the natural isotopic abundance (hydrogen and 12C) versus a heavy form with two deuteriums and one atom of 13C

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results in a mass difference of 6 Da between the labeled peptides, which enhances resolution by MS (94). This technique has been adapted for multiplexing in proteomic studies by combining light and heavy forms of formaldehyde with normal (hydrogen) versus deuterated cyanoborohydride (95).

3.4

Proteolytic Labeling

Isotopic labeling of peptides at the carboxy-terminus by 18O occurs during enzymatic digestion of the parent protein. Differential labeling of peptides is achieved by the action of proteases such as trypsin and endoprotease GluC, which catalyze the exchange of two 16O for two 18O atoms, thus causing a mass shift of 4 Da. Proteolytic labeling was first applied to MS-based comparative proteomics in 2001 by measuring the relative signal intensities of the labeled and unlabeled peak pairs from two serotypes of adenovirus (96,97). This technique confers significant advantages over metabolic and chemical labeling strategies; it is simple, inexpensive, free from side reactions, and applicable to virtually any type of sample. In spite of these advantages, proteolytic labeling suffers from variable incorporation of the isotopic label. Incomplete exchange of 18O can occur, resulting in [16O, 18O] or both [18O] oxygen atoms incorporated at the C-terminus. A complex isotopic distribution arising from the mixture of unlabeled and variably labeled peptides makes it difficult to correctly ascertain the 16O/18O ratios, thus complicating analysis (98,99). Slow exchange of the second 18O atom often leads to incomplete labeling, which can be circumvented by carrying out 18O incorporation under acidic pH (100). The isotopic label can be lost by acid-catalyzed nonenzymatic back exchange when the two protein digests are mixed together for analysis, though this is not observed under mildly acidic conditions typically employed for LC runs during mass spectrometry (101). Protease-catalyzed back exchange can also lead to loss of the label (102), but using a Lys-N enzyme for protein digestion, which introduces a single 18O at the C-terminus and hence does not alter 16O/18O ratios, significantly addresses this issue (103). Incorporation of a single 18O atom also eliminates variable incorporation of the label, but the resultant mass shift of only 2 Da can go undetected if the mass spectrometer is not adequately high resolution. A recently developed method for 18O-based quantification of peptides uses catalysis with hydrochloric acid instead of standard proteases for labeling. Unlike proteolytic labeling, acid catalysis labels all carboxyl groups present in a peptide (Asp, Glu, carboxymethylated Cys, C-terminal carboxyl groups). Thus, most peptides labeled by this strategy have a mass difference >4 Da, making data analysis simpler and more reliable than conventional proteolytic labeling approaches (104). Proteolytic labeling has not yet been applied to peptidomic analysis, which is carried out in the absence of enzyme digestion. However, sample preparation under 18O-enriched conditions, allowing for the endogenous peptide pool

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to incorporate the isotopic label during the general turnover process, can potentially answer critical questions regarding stability of the peptides predominant in the peptidome.

4 CONCLUDING REMARKS A large variety of approaches have been developed for quantification of proteins, and some of these have been developed for analysis of peptides. Each of the methods has advantages and disadvantages, and as a result the choice of approach depends largely on the experimental goals and details. Remarkable progress has been made over the past decade in the development of reagents and mass spectrometry instruments. It is now possible to accurately measure the levels of peptides present in small amounts of tissue or cells, and to identify many of these peptides from MS/MS spectra. In looking back at techniques used several decades ago, what can now be done in a few days would have taken decades of work from large teams of scientists, first to purify each peptide and then to determine the sequence. Several Nobel prizes were awarded for discoveries of bioactive peptides (insulin, vasopressin, thyrotropin-releasing hormone, gonadotropin-releasing hormone, and others), based on the importance of the discovery as well as the many years of work involved with the isolation and sequencing of each of these peptides. With current methods, it would have taken days or weeks to identify the peptides. However, it still takes considerable time to learn the function of peptides found by peptidomic approaches. The use of quantitative methods to learn which peptides are regulated by particular treatments can help focus further studies to those peptides that are highly regulated, and therefore likely to play important biological roles. Thus, quantitative peptidomics is an advance over nonquantitative peptidomic methods.

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