Proteolytic 18O-Labeling Strategies for Quantitative Proteomics

Transcription

1 See discussions, stats, and author profiles for this publication at: Proteolytic 18O-Labeling Strategies for Quantitative Proteomics ARTICLE in MASS SPECTROMETRY REVIEWS JANUARY 2007 Impact Factor: 7.71 DOI: /mas Source: PubMed CITATIONS 151 READS 74 2 AUTHORS: Masaru Miyagi Case Western Reserve University 106 PUBLICATIONS 3,853 CITATIONS Kode CHANDRASEKHARA Rao Pondicherry University 35 PUBLICATIONS 435 CITATIONS SEE PROFILE SEE PROFILE Available from: Masaru Miyagi Retrieved on: 08 April 2016

2 PROTEOLYTIC 18 O-LABELING STRATEGIES FOR QUANTITATIVE PROTEOMICS Masaru Miyagi* and K.C. Sekhar Rao Case Center for Proteomics, School of Medicine, Case Western Reserve University, Cleveland, Ohio Received 10 May 2006; received (revised) 28 August 2006; accepted 12 September 2006 Published online 3 November 2006 in Wiley InterScience ( DOI /mas A number of proteomic techniques have been developed to quantify proteins in biological systems. This review focuses on the quantitative proteomic technique known as proteolytic 18 O-labeling. This technique utilizes a protease and H 2 18 Oto produce labeled peptides, with subsequent chromatographic and mass spectrometric analysis to identify and quantify (relative) the proteins from which the peptides originated. The technique determines the ratio of individual protein s expression level between two samples relative to each other, and can be used to quantitatively examine protein expression (comparative proteomics) and post-translational modifications, and to study protein protein interactions. The present review discusses various aspects of the 18 O-labeling technique, including: its history, the advantages and disadvantages of the proteolytic 18 O-labeling technique compared to other techniques, enzymatic considerations, the problem of variable incorporation of 18 O atoms into peptides with a discussion on recent advancements of the technique to overcome it, computational tools to interpret the data, and a review of the biological applications. # 2006 Wiley Periodicals, Inc., Mass Spec Rev 26: , 2007 Keywords: 18 O-labeling; quantitative proteomics; comparative proteomics; mass spectrometry; oxygen-18; carboxyl oxygen exchange reaction more, increasingly, quantitative answers are required (e.g., changes in a protein s expression levels). Therefore, development of quantitative proteomic methods is of critical importance for the advancement of proteomic research. Although there are a number of quantitative proteomic methods that have been developed and have been applied to biological samples, further development of these techniques are required to address existing limitations. No technique currently satisfies all the demands required for accurate proteomic determinations. The value of the information obtained in proteomic studies continues to drive the use of these techniques even with their shortcomings. Even though proteolytic 18 O-labeling is not yet a common technique compared to other quantitative proteomic techniques, it has the potential to be useful and to become a widely used technique. There are two major drawbacks to the technique: (1) variable incorporation of 18 O atoms into peptides when trypsin was used as a catalyst (Julka & Regnier, 2004); and (2) lack of broad-based computational tools. However, recent advancements in the 18 O-labeling techniques and development of computational tools show considerable promise to overcome these problems. This review will focus on the recent advances of the proteolytic 18 O-labeling technique. I. INTRODUCTION The completion of the genome sequencing of a multitude of organisms, the emergence of new technologies in mass spectrometry, and the development of computational tools that link mass spectrometry data with protein sequence databases for the identification and covalent structural analysis of proteins have together fostered unprecedented opportunities to study proteins on a large scale. Proteins are identified by using these tools by analyzing peptides that are generated from the proteins by enzymatic hydrolysis or chemical fragmentation. However, the mere identification of a protein expressed in a biological system is not sufficient to answer most biological questions. More and Contract grant sponsor: National Institutes of Health; Contract grant numbers: EY014020, RR016741, RR *Correspondence to: Masaru Miyagi, Case Center for Proteomics, School of Medicine, Case Western Reserve University, Euclid Avenue, BRB 928, Cleveland, OH masaru.miyagi@case.edu II. CURRENT QUANTITATIVE PROTEOMIC METHODS BRIEF OVERVIEW All current proteomic methods that quantify unknown proteins are relative methods: meaning protein amounts determined in a sample are measured relative to the amounts of the same proteins in other samples. Absolute quantification of proteins can be done by using isotopically labeled synthetic peptides and mass spectrometry (Gerber et al., 2003); however, this goal requires foreknowledge of the target proteins and preparation of isotopically-labeled synthetic peptides for each of the target proteins. This need precludes the use of this technique in global quantitative proteomic analysis that seek to identify and quantify unknown proteins in a sample. A number of quantitative proteomic methods have been developed as shown in Figure 1. There are two primary strategies used in current quantitative proteomics: two-dimensional gel electrophoresis (2D-PAGE)-based methods and mass spectrometric methods based on isotope labeling strategies. A brief overview of different quantitative proteomic methods that discusses their strengths and limitations is given below. More Mass Spectrometry Reviews, 2007, 26, # 2006 by Wiley Periodicals, Inc.

3 & MIYAGI AND RAO FIGURE 1. Classification of current quantitative proteomic techniques. details of the different quantitative proteomic techniques are given in recent review articles (Conrads, Issaq, & Hoang, 2003; Lill, 2003; Righetti et al., 2004). A. 2D-PAGE-Based Methods 2D-PAGE-based methods have been the most commonly used technique in quantitative proteomics. In these methods, protein extracts from different biological samples are separated by 2D-PAGE in parallel. Individual proteins are quantified by comparing the intensities of the same protein spots on the different 2D-gels. The identities of proteins in the spots of interest are determined by performing in-gel enzymatic digestion of the proteins in the spots, followed by analysis of the generated peptides by mass spectrometry. The strength of 2D-PAGE-based methods is in its high resolving power. The technique can separate several thousand proteins in a single gel (Gorg, Weiss, & Dunn, 2004), far surpassing the capability of any other protein separation technique. However, this technique has a number of limitations. The most serious problem intrinsic to current 2D-PAGE techniques is the presence of multiple proteins in any stained spot. Quantification of a protein in a spot is based on the intensity of the spot. This detection step is done prior to the digestion and mass spectrometric analysis of the proteolytically-generated peptides. Multiple proteins in a single spot is a problem because the intensity is not related to one protein (goal) but to all the proteins present in the spot (problem). The mass spectrometric data (mass spectrum) cannot provide the quantitative information without an internal reference protein for each protein in each spot (a technical impossibility). The occurrence of multiple proteins in a single spot appears to happen quite often (30% of 2D-gel spots) (Campostrini et al., 2005). Other limitations of the FIGURE 2. The experimental flow-chart of in vitro stable isotope-labeling method. A: Chemical isotope labeling and (B) proteolytic 18 O-labeling. 122 Mass Spectrometry Reviews DOI /mas

4 PROTEOLYTIC 18 O-LABELING & 2D-PAGE-based methods include difficulties in analyzing membrane proteins (Ghosh et al., 2004) and difficulties in analyzing proteins that have extreme isoelectric points and sizes (Zhan & Desiderio, 2005). Though some laboratories were able to achieve >95% reproducibility in 2D-gels (Zhan et al., 2003) unless these 2Dgels are run by a skilled personnel, the most common problems encountered by many laboratories are the reproducibility of the 2D-gels resulting from gel-to-gel variation and normalization of the spot intensities in the gel sets. To solve these problems, two dimensional-differential gel electrophoresis (2D-DIGE) has been introduced (Unlu, Morgan, & Minden, 1997). In 2D-DIGE technique, proteins from different samples are first labeled with one of the three spectrally-resolvable fluorescent dyes (Cy2, Cy3, and Cy5). Then, the labeled samples are pooled and subjected to 2D-PAGE analysis. Because proteins from different samples are separated in a single gel, this eliminates the gel-to-gel variation that occurs in 2D-PAGE step, the overall technical variation can be significantly reduced. B. Mass Spectrometry Methods Based on Stable Isotope Labeling Strategies Alternate quantitative proteomic methods have been actively sought because of the limitations of 2D-PAGE-based methods. One alternate technique that has been developed is the mass spectrometry based on stable isotope labeling strategy. The technique involves isotope incorporation into proteins or peptides, followed by quantification of the isotopically labeled peptides, using mass spectrometry (with interfaced HPLC), as detailed below. The mass spectrometry based on stable isotope labeling method provides quantitative and identification information. In contrast, the mass spectrometric data from the 2D- PAGE method gives only identification information. The mass spectrometry based on stable isotope labeling method also overcomes some of the drawbacks of 2D-PAGE-based methods such as difficulties in analyzing membrane proteins and proteins that have extreme isoelectric points and sizes. However, the mass spectrometry based on stable isotope labeling method is less sensitive than the 2D-PAGE-based method in detecting minor covalent structural changes, such as post-translational modifications on a protein and different isoforms of a protein. This is because the 2D-PAGE method determines the size and isoelectric point of the proteins, whereas the mass spectrometry based on stable isotope labeling techniques do not. Recent reviews of mass spectrometry based on stable isotope labeling methods include (Julka & Regnier, 2004, 2005; MacCoss & Matthews, 2005; Ong & Mann, 2005; Yan & Chen, 2005). There are two classes of mass spectrometry based on stable isotope labeling techniques: (1) stable isotope labeling of proteins that is achieved metabolically in vivo; (2) and stable isotope labeling that is achieved chemically or enzymatically in vitro. In the first method, stable isotopes are incorporated metabolically in vivo into all proteins. One such method is Stable Isotope Labeling by Amino Acids in Cell Culture (SILAC) (Ong et al., 2002). In SILAC, two groups of cells are grown in culture media; one contains light isotopes in the amino acid(s) and the other contains one or more heavy isotopes incorporated into the amino acid(s) (e.g., L-arginine and 13 C-labeled L-arginine). These light and heavy amino acids are incorporated into proteins during the process of cellular anabolism in the different groups of cells, respectively. Equal numbers, or weights, of cells in the two groups are combined, the proteins are extracted, and the extracted proteins are digested with a protease. The generated peptides are subsequently quantified by mass spectrometry. The metabolic labeling method is expected to have the smallest technical variations compared to other quantitative proteomic methods in the sample-processing step. This is because the two groups of cell samples are combined immediately after harvesting; therefore, any protein/peptide losses in the remaining steps would occur equally for the light and heavy isotope-containing proteins/peptides. This method has been validated to give reasonably accurate and precise results (Molina, Parmigiani, & Pandey, 2005). In addition to SILAC method, other metabolic labeling methods utilizing an isotopically depleted media enriched in 15 N (Oda et al., 1999; Conrads et al., 2001; Washburn et al., 2003) to isotopically label proteins have also been demonstrated to be useful. However, because these methods involve metabolic labeling, they cannot be applied to nonviable samples, such as clinical tissue and body fluid sample. The in vitro stable isotope labeling method is the second class of mass spectrometry based on stable isotope labeling technique. In vitro stable isotope-labeling can be applied universally for any type of sample, which is a major advantage of this technique over the in vivo metabolic labeling method described above. However, higher technical variations in the sample-processing step are expected, as compared with the in vivo metabolic labeling method. This variation is because the two sample groups for the in vitro technique cannot be mixed until the isotope labeling has been accomplished, mixing usually takes place after the digestion of the proteins. The in vitro stable isotope labeling method can be classified further into two classes as shown in Figures 1 and 2: chemical isotope labeling and proteolytic 18 O-labeling. A major difference between these two methods is where isotope labeling is performed. Isotope coded affinity tag (ICAT) (Gygi et al., 1999) is the commonly used chemical isotope labeling strategies. In the ICAT labeling method, the two different protein mixtures representing the two different cell states are treated with isotopically light (e.g., H, 12 C) and heavy (e.g., D, 13 C) ICAT reagents, respectively, by covalently reacting with the cysteinyl residues in proteins. The two protein mixtures are combined, proteolyzed, and analyzed by liquid chromatography tandem mass spectrometry. ICAT technique has been reported to give reasonably good reproducibility (18.6% coefficient of variation determined by analyzing the same Escherichia coli proteome repeatedly) (Molloy et al., 2005). Recently, an alternative approach analogous to ICAT called isobaric tags for relative and absolute quantitation (itraq) has been developed (Ross et al., 2004). itraq technology involves the parallel protein extraction, routine enzymatic digestion, and subsequent labeling of peptides with isobaric tags. itraq reagents covalently react with all the amino terminus and lysine side chain of all peptides. Fragmentation of the tag attached to the peptides generates a low molecular mass reporter ion that is unique to the tag used to label each of the digests. Measurement of the intensity of these reporter ions, enables relative quantification of the peptides in each digest Mass Spectrometry Reviews DOI /mas 123

5 & MIYAGI AND RAO and hence the proteins from where they originate. Using the itraq technology, more than two samples can be analyzed at a time (DeSouza et al., 2005). In contrast, the proteolytic 18 O-labeling method achieves isotope labeling concurrent with the proteolytic digestion of proteins in both sample groups; one sample incorporates 16 Oby digestion in H 2 16 O solvent, and the other sample incorporates 18 O by digestion in H 2 18 O. In the proteolytic 18 O-labeling method, the variability of the yield of each isotopically labeled peptide depends only on the proteolytic digestion reaction. In contrast, the chemical isotope labeling method has two steps in which a variability in yield can occur: an isotope labeling step and a proteolytic digestion step. Therefore, it is expected that the proteolytic 18 O-labeling method has a smaller technical variations than the chemical isotope labeling method in the sampleprocessing step. One note to the above discussion; isotope labeling can be performed on proteins prior to the digestion step in a modified chemical isotope labeling method. This modification allows a mixing of the two samples before the proteolytic digestion; therefore, the variability that might arise from different efficiencies in the proteolytic step is eliminated. However, it is well recognized that complete labeling of targeted functional groups at the protein level is not easy because of problems with the accessibility of the reagent to the targeted sites that exist even on denatured proteins. Therefore, variable degrees of labeling could occur on the identical target sites on the proteins in two different sample proteins. The principle of quantification of 16 O- and 18 O-labeled peptides by mass spectrometry is schematically shown in Figure 3. Although the example is for 16 O- and 18 O-labeled peptides, the same principle can be applied for other mass spectrometry based on stable isotope labeling techniques. The same peptides generated from the same protein from two different sample groups differ only in their molecular weights: one has 16 O in the C-terminal carboxyl groups of the peptides (no change in the oxygen isotope), and the other incorporates 18 O. Because the peptide labeled with 16 O and the same peptide labeled with 18 O co-elute from the chromatography step in the LC/MS/MS analysis (Reynolds, Yao, & Fenselau, 2002), a constant difference in molecular weight (2 Da in the case of a single 18 O atom incorporation, and 4 Da with two 18 O atom incorporation) exists in the 16 O- and 18 O-labeled peptides; that difference can be readily distinguished in the mass spectrum. The relative abundance of the two peptides can be determined by comparing the peak intensities of the 16 O- and 18 O-labeled peptides in the mass spectrum, the relative abundance also equals the relative abundance of the protein from which the peptide was generated in the original samples. The identity of the peptide, which in most cases identifies the protein, can be determined by subjecting one of the peptide ions ( 16 O- or 18 O-labeled peptide) to tandem mass spectrometry to obtain the amino acid sequence from a MS/MS spectrum. The quantification in the proteolytic 18 O-labeling technique is based on the assumption that the ionization efficiencies of the 16 O- and 18 O-labeled peptide are identical. It is also important to note that C-terminal peptides of proteins are different than other proteolytically generated peptides in these labeling experiments. In most cases, the C-terminal peptides of proteins do not have any incorporated 18 O atoms because of the lack of lysine or arginine at their C-termini when trypsin is used. However, 18 O atom incorporation into C-terminal peptides of proteins bearing lysine or arginine does occur. It is likely that data from C-terminal peptides will falsely indicate large changes in protein expression. Therefore, the sequence location of the peptide should always be monitored to eliminate the possibility of misleading protein expression levels. Qian et al. (2005) have validated the 18 O-labeling technique by labeling two equal aliquots of plasma samples by trypsin. When the 1:1 ( 16 O-labeled: 18 O-labeled peptide) mixture was analyzed by a fourier transform ion cyclotron resonance mass spectrometer the average ratio of 891 peptides identified was , demonstrating the accuracy of 18 O-labeling method even in highly complex human plasma samples. In the 18 O-labeling strategy, the difference in molecular weight between the 16 O- and 18 O-labeled peptide is only 2 Da (single 18 O atom incorporated peptide) or 4 Da (two 18 O atom incorporated peptide), therefore the isotope envelopes of the 16 O- and 18 O-labeled peptide overlap. Therefore, it is required to correct the intensities of the observed 18 O-labeled peptide peak by subtracting the isotopic contribution of the 16 O-labeled peptide to the 18 O-labeled peptide peak from the observed 18 O-labeled peptide peak (Rao, Carruth, & Miyagi, 2005a). The required correction of the 18 O-labeled peptide peak intensity may decrease the accuracy of the method as isotopic contribution of the 16 O-labeled peptide to the 18 O-labeled peptide peak increases (Rao, Carruth, & Miyagi, 2005a). Whereas in the other isotope labeling methods such as SILAC and itraq, the light and heavy peptide ions are separated by 6 10 Da. So the isotopic contribution from the light peptide ion to the heavy peptide ion is very small, therefore it can be ignored. It should also be noted that significant degree of chemical back exchange of the carboxyl oxygen of 18 O-labeled peptides to oxygen-16 could occur in H 2 16 O solvent, especially at extreme ph. The degree of carboxyl oxygen exchange in mild acidic conditions (0.1 5% v/v formic acid) at 48C within 24 h has been reported to be negligible (Stewart, Thomson, & Figeys, 2001), even so it is recommended to dry the labeled peptides and store at low temperature if the peptides are not analyzed immediately. III. HISTORY OF USE OF 18 O-LABELING IN PROTEIN QUANTIFICATION The use of oxygen-18 ( 18 O) labeling in protein quantification has its origin in the work of Sprinson and Rittenberg, who studied the mechanism of the product inhibition in the chymotrypsin-catalyzed amide bond hydrolysis reaction (Sprinson & Rittenberg, 1951). They found that the incubation of carbobenzoxy-phenylalanine (a product of the amide substrate) with chymotrypsin in H 2 18 O solvent resulted in incorporation of 18 O atoms into the carboxyl group of the carbobenzoxyphenylalanine. This 18 O atom incorporation reaction into the C-terminal carboxyl group of a peptide is essentially a carboxyl oxygen exchange reaction-as described later. The study clearly demonstrated that the products of the chymotryptic amide bond 124 Mass Spectrometry Reviews DOI /mas

6 PROTEOLYTIC 18 O-LABELING & FIGURE 3. Principle of quantification by mass spectrometry. Each 16 O-labeled peptide from one sample and the corresponding 18 O-labeled peptides from the other sample (A) co-elute from in the LC/MS/MS analysis (B). By comparing the peak heights of the 16 O- and 18 O-labeled peptide in the mass spectrum, the relative abundance of the two peptides (and thus the relative abundance of the protein in the two samples from which the peptides were generated) can be determined (C). By subjecting one of the peptide ions to MS/MS analysis, the identity of the protein can be determined from the amino acid sequence and database search tools (D). Mass Spectrometry Reviews DOI /mas 125

7 & MIYAGI AND RAO hydrolysis reaction are not only competitive inhibitors of chymotrypsin, but are also substrates for the carboxyl oxygen exchange reaction. The first use of 18 O-labeling in peptide quantification was performed by Desiderio and Kai (1983). They enzymatically incorporated 18 O atoms into the carboxyl termini of leucine- and methionine-enkephalin, and used them as 18 O-labeled internal standards to quantify endogeneous leucine- and methionineenkephalin in thalamus tissue by field desorption mass spectrometry. Although this technique was developed to quantify peptides, the principle of quantification is the same as the principle by which proteolytic 18 O-labeling is used in protein quantification (described later). However, that initial research did not spur and further development of the technique applied to protein quantification until relatively recently. Other uses of proteolytic 18 O-labeling not related to quantification have also been developed, including: identification of the C-terminal peptide of a protein (Rose et al., 1983), identification of chemically cross-linked (Back et al., 2002) and chemically modified (Sun & Anderson, 2005) peptides, and facilitation of the assignment of peptide product ions produced by collision-induced dissociation (Takao et al., 1991; Schnolzer, Jedrzejewski, & Lehmann, 1996). Mirgorodskaya et al. (2000) expanded the use of proteolytic 18 O-labeling to protein quantification, and suggested the potential use of this technique in comparative proteomics. Shortly after this report, Stewart, Thomson, and Figeys (2001) reported a more detailed study of the labeling conditions pertinent to proteomics. The first proteomic application of this technique was reported by Yao et al. (2001), who quantified proteins in two serotypes of adenovirus. Since then, a number of articles have described this technique and/or reported its biological applications. This technique has steadily grown in quantitative proteomic studies. IV. ENZYMOLOGY OF PROTEOLYTIC 18 O-LABELING REACTION The proteolytic enzymes catalyze the following two hydrolytic reactions: ð1þ RC 16 ONHR 0 þ H 18 2 O protease! RC 16 O 18 O þ þ H 3 NR 0 ð2þ RC 16 O 18 O þ H 18 2 O protease! RC 18 O 18 O þ H 16 2 O Hydrolysis of a protein in H 2 18 O by a protease results in the incorporation of one 18 O atom into the carboxyl terminus of each proteolytically generated peptide. This mechanism involves a nucleophilic attack by a solvent water molecule on the carbonyl carbon of the scissile peptide bond (reaction 1). Following this hydrolysis reaction, all proteases could incorporate one more 18 O atom into the carboxyl terminus of the proteolytically generated peptide. This second incorporation results in two 18 O atoms being incorporated into the carboxyl terminus of the peptides (reaction 2). The second 18 O atom-incorporation reaction (reaction 2) is essentially the reverse reaction of peptide-bond hydrolysis or the peptide-bond formation reaction (protease catalysts accelerate the forward and reverse reactions). There are in fact numerous experimental studies that verify the occurrence of the proteasecatalyzed peptide-bond formation reaction (for example, Bongers & Heimer, 1994). The only difference between reaction 2 and the peptide-bond formation reaction is the reacting reagent and the nucleophilic atom on that reagent. In the carboxyl oxygen exchange reaction, the reagent is water, which has an oxygen atom nucleophile, where as in the peptide bond formation reaction, the reagent is an amine component (H 2 N R) with a nitrogen nucleophile. Therefore, there will be complete incorporation of two 18 O atoms by the protease with sufficient time for the reaction to reach equilibrium in H 2 18 O (under the appropriate conditions). It should be noted that complete two 18 O atom incorporation in this review means that >98% of the two oxygen atoms in the C-terminal carboxyl group of a peptide are replaced with the oxygen atoms from solvent water used regardless of the content of H 2 18 O in the solvent water. The 98% oxygen replacement is sufficient in our practice, considering that an overall technical variation of the method is significantly higher than the 2% variation generated in this step. Note: in this review, reaction 2 will be referred to as the carboxyl oxygen exchange reaction, which indicates the actual outcome of the reaction. There is a difference in the efficiency of the first (reaction 1) and second (carboxyl oxygen exchange reaction, reaction 2) 18 O- atom labeling reactions. The first 18 O atom is incorporated from H 2 18 O upon the proteolytic cleavage of a peptide bond; therefore, only one cycle of the enzyme reaction is required to incorporate the first 18 O atom into the peptide. On the other hand, the carboxyl oxygen exchange reaction is required to occur several times on the peptide to achieve the complete incorporation of two 18 O atoms because there is a 50% chance of replacing the newly incorporated 18 O atom during the reaction, this replacement does not lead to any net single 18 O atom incorporation. A 50% chance assumes that the reaction rate to replace the 18 O atom and to replace a 16 O atom are the same (no isotopic effect). The yield of 18 O atom incorporation into a peptide after five enzymatic carboxyl oxygen exchange reactions is %, which is an acceptable yield in proteomic experiments, considering the overall variations for such a technique. If the reaction continues, then both the oxygen atoms in the C-terminal carboxyl group of the peptide should eventually come to equilibrium with oxygen atoms from H 2 18 O. Because the major problem of the proteolytic 18 O-labeling technique applied to quantitative proteomics is the variable incorporation of 18 O atoms into peptides, it is essential to control the carboxyl oxygen exchange reaction (reaction 2) as a key strategy in overcoming this problem. The variable 18 O atom incorporation problem is described in the next section. V. A MAJOR PROBLEM INHERENT TO 18 O-LABELING TECHNIQUE This technique often suffers from the generation of a mixture of isotopic isoforms that result from the variable incorporation of either one or two 18 O atoms ( 18 O 1 / 18 O 2 ) into each peptide species. This variable 18 O atom incorporation problem is because of the second 18 O atom incorporation reaction (carboxyl oxygen exchange reaction), that is extremely slow under the conditions 126 Mass Spectrometry Reviews DOI /mas

8 PROTEOLYTIC 18 O-LABELING & commonly used, and requires multiple enzymatic reactions (as discussed above) that lead to a variable exchange within the timeframe of the proteolytic reaction. Unfortunately, the rates of exchange are not predictable and vary from peptide to peptide. This variable 18 O atom incorporation complicates the quantification of the peptides, and increases the error in the calculation of the 16 O- to 18 O-labeled peptide ratios (Julka & Regnier, 2004; Ong & Mann, 2005). This complication is illustrated in Figure 4. When the unlabeled 16 O-labeled peptide and the same peptide labeled with variable numbers of 18 O atoms are mixed, the mass spectrum is complex (Fig. 4A). To accurately quantify the relative abundance of the 16 O- and 18 O-labeled peptides, the following three peaks must be used for the calculation: (1) the monoisotopic peak of the unlabeled 16 O-labeled peptide ( 16 O), (2) the monoisotopic peak of the peptide labeled with one 18 O( 18 O 1 ), and (3) the monoisotopic peak of the peptide labeled with two 18 O atoms incorporated ( 18 O 2 ). The intensity of 16 O-labeled peptide peak indicates the amount of the protein in the unlabeled sample, whereas the added monoisotopic intensities of the 18 O 1 and 18 O 2 peak indicate the amount of protein in the labeled samples. The equations needed to calculate the results from experiments in which there is a mixture of one and two 18 O atoms in the labeled peptide are very complicated (Johnson & Muddiman, 2004). The results from these mixed isotope experiments are expected to have a greater variability compared to the results obtained for a peptide that was completely labeled with two 18 O atoms. This variable 18 O atom incorporation mainly attributes to the difficulty of obtaining the accurate intensities of the 18 O-labeled peptide monoisotopic ions ( 18 O 1 and 18 O 2 ion) in the mass spectrum. For example, a lower precision in the 18 O 1 ion arises because of the 16 O þ 2 ion, which superimposes on the 18 O 1 ion (same m/z value). Likewise, the 18 O 2 ion is interfered by the superimposed 18 O 1 þ 2 ion, as well as by the 16 O þ 4 ion. On the other hand, when the 16 O unlabeled peptide and the same peptide labeled completely with two 18 O atoms are mixed, the observed mass spectrum will be simpler (Fig. 4B). Importantly, we need to use only two peaks, the monoisotopic ion of 16 O-labeled peptide ( 16 O) and the monoisotopic ion of two 18 O atom-labeled peptide ( 18 O 2 ), to calculate the relative abundance of the 16 O- and 18 O-labeled peptides. This complete two 18 O atom incorporation is expected to improve the precision of the measurement significantly because of the smaller number of peaks in the calculation, and the small intensity of the interfering 16 O þ 4 peak compared to the intensity of the 18 O 2 peak. To overcome the problem of the variable 18 O atom incorporation into peptides, it is apparent that either an invariable single 18 O atom incorporation or a complete two 18 O atom incorporation must be achieved. To attain a predominate single 18 O atom incorporation, the carboxyl oxygen exchange reaction (Reaction 2) needs to be slowed down significantly (virtually stopped). On the other hand, to achieve a complete two 18 O atom incorporation, the carboxyl oxygen exchange reaction must be accelerated significantly. Finding reaction conditions to achieve these results would be a significant step towards improving the quantitative proteomics methods that utilize proteolytic 18 O-labeling. Proteases can be classified into two groups: (1) class-1 proteases that hydrolyze the N-terminal side of the peptide bond of a specific amino acid such as peptidyl-lys-metallopeptidase (Lys-N, cleaves the X-Lys bond) and endoproteinase Asp-N (cleaves the X-Asp bond); and (2) class-2 proteases that hydrolyze the C-terminal side of the peptide bond of a specific amino acid such as lysyl endopeptidase (Lys-C, cleaves the Lys- X bond) and trypsin (cleaves the Arg-X and Lys-X bonds). Proteases in the former class are expected to be suited to achieve single 18 O atom incorporation, and proteases in the latter class are expected to be suited for complete two 18 O atom incorporation, for the reasons illustrated in Figure 5. Lys-N represents the class 1 proteases and Lys-C the class 2 proteases in Figure 5. The cases of Lsy-N and Lys-C are discussed next. In the following discussion, the terminology used below for catalytic subsites of proteases was proposed previously (Berger & Schechter, 1970). The subsites in a protease are numbered from the catalytic site, S Sn towards the N-terminus of the substrate, and S Sn 0 towards the C-terminus. The substrate amino acid residues that the subsites in a protease bind are numbered P Pn, and P Pn 0, respectively, as follows. FIGURE 4. Mass spectra observed in variable and complete two 18 O atoms incorporation. (A) variable incorporation and (B) complete two 18 O atom incorporation. Mass Spectrometry Reviews DOI /mas 127

9 & MIYAGI AND RAO FIGURE 5. The second 18 O atom incorporation reaction by Lys-N and Lys-C. Peptides, H-Lys-Xaa-Xaa- 16 OH, and H-Xaa-Xaa-Lys- 16 OH represent Lys-N and Lys-C peptide, respectively. A: Weak and productive binding between Lys-N peptides and Lys-N. B: Strong and non-productive binding between Lys-N peptides and Lys-N, C: Strong and productive binding between Lys-C peptides and Lys-C, (D) weak and non-productive binding between Lys-C peptides and Lys-C. First is the case of Lys-N. Lys-N-generated peptides have a lysine residue at their N-terminus, but no specific amino acids at their C-terminus (H-Lys ---- Xaa-OH). For this protease, Lys-N peptides must bind to the S Snsite of Lys-N in order for an 18 O atom to be incorporated into the C-termini-as illustrated in Figure 5A. However, the binding affinity between a Lys-N peptide s C-terminal amino acids (Xaa) and the S1 site is assumed to be significantly lower than the binding affinity between a Lys-N peptide s N-terminal lysine residue and the S1 0 site, because Lys-N s strong specific affinity to the lysine residue belongs to the S1 0 site. Therefore, it is expected that Lys-N peptides tend to bind to the S Sn 0 site (Fig. 5B, strong/nonproductive binding), not to the S Sn site (Fig. 5A, weak/ productive binding). Thus, a slower incorporation of the second 18 O atom is expected because of the smaller proportion of Lys-N peptide binding to the S1 ----Snsite. For this reason, a single 18 O atom incorporation is most likely to be achieved, if the binding between Lys-N peptides C-terminal amino acids and the S1 site can be inhibited sufficiently. Second is the case of Lys-C. Lys-C-generated peptides have a lysine residue at their C-terminus, but no specific amino acids at their N-terminus (H-Xaa Lys-OH); that result is the reverse arrangement compared to the Lys-N peptides. Lys-C peptides are expected to bind strongly to the protease s S Sn site (Fig. 5C) because of the S1 site s strong affinity to the C-terminal lysine residue of the Lys-C peptides. Binding of the Lys-C peptides to the protease s S1 ----Sn site is the productive binding site to yield 18 O-labeling. On the other hand, the non-productive binding is the binding of Lys-C peptides to S Sn 0 site (Fig. 5D), is expected to be weak. Thus, unlike the case of Lys-N, productive binding is the dominant binding in the case of Lys-C. It is, therefore, expected that the second 18 O atom incorporation reaction by Lys-C is significantly more efficient compared to Lys- N, and, because of the higher affinity of the S1 site for the peptide, less susceptible to inhibition. Thus, complete two 18 O atom incorporation might be achieved with Lys-C, if conditions could be found so that the reaction is further accelerated. VI. RECENT ADVANCES There are two approaches to overcome the problem of variable incorporation of 18 O atoms: (1) develop a true single 128 Mass Spectrometry Reviews DOI /mas

10 PROTEOLYTIC 18 O-LABELING & 18 O atom-labeling technique, and (2) develop a true two 18 O atom-labeling technique. Recent advances in developing these two labeling techniques are reviewed below. A. Single 18 O Atom-Labeling Technique To achieve predominant single 18 O atom incorporation, the carboxyl oxygen exchange reaction (reaction 2) must be virtually stopped, as discussed above. Conditions to achieve single 18 O atom incorporation have been explored with Lys-N (Rao, Carruth, & Miyagi, 2005a). Presumably, class 1 proteases, such as Asp-N, could be used in a single 18 O-labeling technique; however, no reports have been published, that developed this specific technique with these proteases. Rao, Carruth, and Miyagi (2005a) have found that Lys-N incorporates only a single 18 O atom at a ph >9.5 by yet-unknown mechanisms, as summarized below. In Figure 6, peptide ions with two 18 O atoms ( 18 O 2 ) are abundant in the three peptides derived from apomyoglobin at ph 6.0. However, incorporation of the second 18 O atom is not observed at ph 10.0, as evidenced by the exact match of the relative intensities of the isotope peaks in the mass spectrum compared to their theoretical abundances for one 18 O atom incorporated into the peptide (Fig. 6, three bottom panels). These results suggest that the second 18 O atom incorporation is ph-dependent and that only one 18 O atom is incorporated at ph 9.5. Note that the original article also presents results at ph 8.0, 9.0, and 9.5 to further support this conclusion. A detailed description of the calculation of the relative abundance of 16 O- and 18 O-labeled peptides can be found in this report (Rao, Carruth, & Miyagi, 2005a). A major advantage of the single 18 O-labeling technique (for example, using Lys-N) is the elimination of a variable incorporation of 18 O atoms into peptides. Another advantage of this technique is that there is no enzyme-catalyzed 18 O backexchange reaction when the two protein digests are mixed. This mixing of digests assumes that the mixture is kept at a ph >9.5. Another advantage of using enzymes such as Lys-N compared to trypsin is that a less-complex protein digest is produced. This less complexity is because the Lys-N enzyme cleaves only at lysine residues, whereas trypsin cleaves at arginine and lysine residues to produce more peptides. A validation study of the Lys-N-based technique has been reported by Rao et al. (2005b). In this study, an equal amount of protein from human retinal pigment epithelium cells in culture was digested with Lys-N, either in H 2 16 OorinH 2 18 O. After the digestion, the two digests were mixed 1:1 (wt:wt) and analyzed with LC/MS. Theoretically, the 16 O- and 18 O-labeled peptide ratios of all of the generated peptides should be 1. For most of the peptides, the measured ratios were within However, approximately 17% of the peptides gave ratios below 0.5 or above 2.0; those data imply false results for 17% of the peptides from biological samples analyzed by this method. Further analysis by the authors suggested that the Lys-N-based method provided inaccurate results for peptides that were the products of cleavage at one or more of the following amino acid sequences: (1) Lys- X 0 3 -Lys (two lysine residues separated by three other amino acids), (2) Glu-Lys, and (3) Pro-Lys. This difference in the digestion efficiency is the major drawback of the Lys-N-based technique. It is, therefore, important to exclude peptides generated from one or more of these sequences in biological applications. Another drawback of this technique is the small mass difference (2 Da) between 16 O- and the same 18 O-labeled peptide, that difference necessitates the use of a high-resolution mass spectrometer such as a quadrupole/time-of-flight type instrument. Therefore, the most utilized mass spectrometer in the field of proteomics, a quadrupole-based ion trap type instrument, cannot be used because it does not have sufficient resolution. B. Two 18 O Atom-Labeling Technique Class 2 proteases are predicted to be suitable for the two 18 O atom-labeling technique, as discussed previously. To achieve complete two 18 O atoms incorporation, the carboxyl oxygen exchange reaction (Reaction 2) must be accelerated significantly. Optimization of reaction conditions to achieve complete two 18 O atom incorporation has been explored with trypsin and Lys-C. Zang et al. (2004) showed that the trypsin-catalyzed carboxyl oxygen exchange reaction is more efficient at ph 6.75 than at ph 8.50, based on experiments that measured the changes of isotopic ions of the labeled peptides after 20 h of labeling reactions. Staes et al. (2004) showed that complete two 18 O atom incorporation with trypsin can be achieved after an overnight reaction at ph 4.5. This experiment suggests that trypsin expresses moderately high carboxyl oxygen exchange activity at an acidic ph, despite showing almost a complete suppression of the amidase activity at these acidic conditions. A recent study done by Hajkova, Rao, and Miyagi (2006) has found that the optimum ph for the carboxyl oxygen exchange reaction catalyzed by Lys-C and trypsin are 5.0 and 6.0, respectively, which are significantly shifted toward acidic phs compared to the ph range of their highest amidase activities (Fig. 7). The authors also determined the steady state kinetics parameters for both proteases at two different ph values; one at the ph optimum for their carboxyl oxygen exchange activities (ph 5 6), and the other at the ph of their highest amidase activities (ph 8 9). The catalytic activities (kcat/km) of Lys-C and trypsin at the acidic phs were 2.5-fold and 17-fold higher than those at the alkaline phs, respectively. In addition to the ph of the reaction, some research groups have found that carboxyl oxygen exchange reaction by trypsin can be increased in aqueous solutions that contains organic solvent such as in 30% acetonitrile (Brown & Fenselau, 2004), 20% methanol (Blonder et al., 2005), and 50% methanol (Nelson et al., 2006). Taken together, it is evident that the carboxyl oxygen exchange reaction can be significantly accelerated by performing the reaction at an acidic ph or in aqueous solutions that contains organic solvent. Validation studies to assess the accuracy of the two 18 O atom-labeling technique, using Lys-C and trypsin at acidic ph or in aqueous solutions containing organic solvent, are yet to be reported. The study done by Hajkova, Rao, and Miyagi (2006) also found that the activity of the carboxyl oxygen exchange reaction of Lys-C (kcat/km) at ph 5.0 was 17-fold higher than that of trypsin at ph 6.0; those results indicated that the use of Lys-C as a catalyst in the 18 O-labeling experiment has an advantage over trypsin in terms of efficiency of 18 O-labeling. However, it is important to note that Lys-C produces peptides that are longer than those produced by trypsin. A high-resolution mass spectrometer might be required for the analysis of the Lys-C-generated Mass Spectrometry Reviews DOI /mas 129

11 & MIYAGI AND RAO FIGURE 6. ph dependency of the second 18 O atom incorporation by Lys-N. Apomyoglobin was digested by 18 Lys-N in H 2 O at ph 6.0 (100 mm sodium phosphate) or ph 10.0 (100 mm glycine-naoh) at 258C for 18 h and the resulting digests were analyzed with LC/MS. A:(Mþ 3H) 3þ ions of KALELFRNDIAA, (B)(Mþ 3H) 3þ ions of KHPGDFGADAQGAMT, and (C) (Mþ 4H) 4þ ions of KVEADIAGHGQEVLIRLFTGHPETLE, at ph 6.0 or ph The bottom spectra are the theoretical spectra of the isotopes calculated for each corresponding peptide that contained one 18 O atom. Reproduced from Rao, Carruth, and Miyagi (2005a) with permission from the American Chemical Society. 130 Mass Spectrometry Reviews DOI /mas

12 PROTEOLYTIC 18 O-LABELING & FIGURE 7. Effects of ph on the initial velocities of the carboxyl oxygen exchange and amidase activities of Lys-C and trypsin. For the carboxyl oxygen exchange activity, a 10 mm solution of Ac-Lys-OH in aqueous buffers at different ph was incubated with 1 mm of Lys-C or trypsin at 258C in 95% H 2 18 O buffered at various ph. The duration of the reaction was 5 min for Lys-C and 20 min for trypsin. For the amidase activity, Ac-Lys- NH 2 (10 mm) was incubated with 0.2 mm of Lys-C or 1 mm of trypsin at 258C in 95% H 2 18 O at various ph. The duration of the reaction was 5 min for Lys-C and 20 min for trypsin. The reaction product, Ac-Lys- 18 OH, was quantified by stable isotope dilution tandem mass spectrometry. A: ph dependency of the carboxyl oxygen exchange and amidase activities of Lys-C. B: ph dependency of the carboxyl oxygen exchange and amidase activities of trypsin. *-: carboxyl oxygen exchange activity, * : amidase activity. All reactions were carried out in triplicate. Reproduced from Hajkova, Rao, and Miyagi (2006) with permission from the American Chemical Society. peptides because of the tendency of longer peptides to produce higher charge states in electrospray than smaller peptides. Another way to increase the 18 O-labeling efficiency, using class 2 proteases, is to use a protease immobilized to a resin. This immobilization allows the investigator to increase significantly the molar ratio of protease-to-substrate to increase the labeling efficiency to produce complete two 18 O atom incorporation. In contrast, increasing protease concentration in the solution is problematic because the protease remains in the digest after the labeling and will interfere with the subsequent mass spectrometry. This interference is because autodigestion of the highconcentration solution protease will produce peptides from the enzyme itself. There are several reports that used immobilized trypsin to incorporate 18 O atoms into peptides (Brown & Fenselau, 2004; Chen et al., 2005; Qian et al., 2005; Hood et al., 2005a). However, the 18 O-labeling reaction was performed at alkaline ph in these studies. The efficiency of the 18 O-labeling could be further increased by performing the 18 O-labeling at an acidic ph (ph 6.0). Another advantage of using immobilized proteases is that no protease-catalyzed oxygen back exchange reaction is expected (after mixing of the samples prior to mass spectrometry) because the immobilized proteases are readily removed from the peptides after the labeling reaction. For these reasons, immobilized proteases will be more frequently used in 18 O-labeling experiments. However, it is important to note that the use of immobilized proteases can increase the variability of the process, because an extra step is added to the experimental procedure. Proteolytic 18 O-labeling, using the class 2 proteases, can be decoupled from protein digestion (Yao, Afonso, & Fenselau, 2003). In other words, the two samples being compared can be digested in H 2 16 O first, and the labeling step can take place after evaporating the H 2 16 O and replacing it with either H 2 16 Oor H 2 18 O. The decoupling procedure has the advantage of separating the digestion and labeling steps; thus, conditions can be optimized separately for each step (e.g., the digestion at ph 8 and the labeling at ph 6 for trypsin). An advantage of the two 18 O-labeling technique over the single 18 O-labeling technique is that there is a larger mass difference (4 Da) between the 16 O- and 18 O-labeled peptides, assuming that a complete two 18 O atom incorporation is achieved. Complete two 18 O atom incorporation should allow the use of a low-resolution mass spectrometer for the analysis. In fact, a quadrupole-based ion trap instrument has been tested and shown to give accurate quantitative results (Heller et al., 2003; Sakai et al., 2005). Note, however, that it is always better to have a baseline separation between isotopic peaks to accurately obtain the intensities of each isotopic peak. Therefore, a high-resolution instrument is still recommended. The achievement of complete two 18 O atom incorporation allows to use only two peaks the monoisotopic peak of a 16 O-labeled peptide ( 16 O) and the monoisotopic peak of a two 18 O-labeled peptide ( 18 O 2 ) to Mass Spectrometry Reviews DOI /mas 131

13 & MIYAGI AND RAO calculate the relative abundance of the 16 O- and 18 O-labeled peptides. Complete two 18 O atom incorporation is expected to significantly reduce the variability of the measurement as discussed before. It is important to note that a protease-catalyzed oxygen back-exchange reaction will occur when the two digests (one digested in H 2 16 O and the other in H 2 18 O) are mixed, unless the protease is completely inactivated before the mixing. Staes et al. (2004) demonstrated the importance of inhibiting the oxygen back-exchange reaction, and showed that the reduction of trypsin s disulfide bonds followed by alkylation of the reduced cysteine residues can completely inactivate trypsin. Therefore, the inactivation process is required for accurate measurement. Trypsin has been the protease of choice among the class-2 proteases because trypsin produces relatively short peptides with a basic amino acid residue at each C-terminus (arginine or lysine) (Paizs & Suhai, 2005). This basic nature makes the peptides wellsuited for tandem mass spectrometric analysis. However, other proteases can also be used. Lys-C has a 17-fold higher carboxyl oxygen exchange activity compared to trypsin, and generates a less complex peptide mixture. Therefore, Lys-C could be advantageous over trypsin, depending on the purpose of the experiment. Endoproteinase Glu-C and chymotrypsin have also been used (Reynolds, Yao, & Fenselau, 2002). However, these two proteases have not been preferable enzymes in proteomic studies, probably because these proteases produce peptides that are not easily sequencable by collision-induced tandem mass spectrometry compared to peptides produced by trypsin and Lys-C. VII. BOTTLENECKS OF THE 18 O-LABELING TECHNIQUE Although there are many advantages of the 18 O-labeling technique, there are two major problems that must be solved before this technique can be fully accepted by the proteomics community. One problem is that there is no validated proteinextraction procedure that is compatible with all of the following experimental steps: protein S-alkylation, digestion, 18 O-labeling, and the chromatographic separation of the generated peptides. The other problem is that there is no general computational tool that can be used by all mass spectrometers and with all database search tools to automatically quantify proteins in these labeling experiments. A. Extraction and Solubilization of Proteins Sodium dodecyl sulfate (SDS) (2%) or urea (8 M) has been traditionally used in the extraction and solubilization of proteins from biological samples. SDS is a well-known detergent that solubilizes proteins very efficiently. However, proteases in general have limited proteolytic activities in SDS solution, and the effect of SDS on the 18 O-labeling efficiencies by any protease has not been reported. Another drawback of using SDS as a protein solubilization agent is that it interferes with chromatographic techniques. Even though most of proteases retain their activities at a low urea concentration (1 M), the efficiency of protein solubilization by urea is less than SDS. Therefore, it cannot be used for the small tissue samples. Thus, better protein extraction and solubilization methods must be developed. Protein solubilization remains a challenge for any shotgun proteomics strategy. B. Computational Tools The interpretation of the mass spectrometry data from labeling experiments requires the following steps: (1) a database search to identify proteins from the MS/MS spectra of 16 O- and/or 18 O- labeled peptides with a tool such as Mascot, (2) extraction of the precursor ion of each 16 O- and 18 O-labeled peptide from the data, and (3) calculation of the relative intensity of each 16 O- and 18 O- labeled peptide ion. The third data-analysis step requires knowledge of the amino acid sequence of the peptide determined by the database search. Amino acid sequence is needed to subtract out the intensity of the M þ 4 isotope ion of the 16 O-labeled peptide to obtain an accurate measurement of the intensity resulting from only the two 18 O atom-incorporated peptide ion (in the case of the single 18 O-labeling technique, the intensity of the Mþ2 isotope of the 16 O-labeled peptide must be calculated and subtracted from the one 18 O atom-incorporated peptide peak). It takes a significant amount of time to do this calculation manually (Rao et al., 2005b). Therefore, the use of computational tools that can automate the process will greatly enhance the throughput of an 18 O-labeling experiment. Fernandez-de-Cossio et al. (2004) developed the webaccessible computational tool, Matching, to automatically calculate the relative intensity of an 16 O- and 18 O-labeled peptide ion. However, the software is not currently linked with a database search tool, and, therefore, the amino acid sequence of the identified peptide and its mass spectrum must be individually loaded before any data processing. Halligan et al. (2005) developed a stand-alone computational tool named Zoom- Quant that quantifies the mass spectra of 16 O- and 18 O-labeled peptides from an ion trap instrument of a particular manufacturer (Thermo Electron Corporation, Waltham, MA) (Hicks et al., 2005). The software is linked with a database search tool Sequest. Another stand-alone computational tool named STEM, is linked with a database search tool Mascot is used for the data processing of 18 O-labeling experiments (Shinkawa et al., 2005). The software that has been developed so far is specific to a particular instrument and is specific to a particular database search tool. Therefore, there is a definite need to develop a user-friendly computational tool that can integrate with any kind of data available from a variety of mass spectrometers. VIII. BIOLOGICAL APPLICATIONS A major use of the proteolytic 18 O-labeling technique has been in comparative proteomics. A number of studies have been published. The sample types are diverse: a virus proteome (Yao et al., 2001), proteomes of cultured cells (Brown & Fenselau, 2004; Blonder et al., 2005; Chen et al., 2005; Rao et al., 2005b), serum (Hood et al., 2005b; Qian et al., 2005), and tissues (Zang et al., 2004; Hood et al., 2005a). Comparative proteomics will continue to be the major area of the application of proteolytic 18 O-labeling. 132 Mass Spectrometry Reviews DOI /mas