Isobaric Peptide Termini Labeling Utilizing Site-Specific N-Terminal

Apr 29, 2011 - North East Proteome Analysis Facility (NEPAF), Devonshire Building, Newcastle upon Tyne, NE1 7RU, United Kingdom. bS Supporting ...
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Isobaric Peptide Termini Labeling Utilizing Site-Specific N-Terminal Succinylation Christian J. Koehler,† Magnus Ø. Arntzen,†,‡,§ Margarita Strozynski,† Achim Treumann,|| and Bernd Thiede*,† †

The Biotechnology Centre of Oslo, University of Oslo, Gaustadalleen 21, 0349 Oslo, Norway Proteomics Core Facility, Oslo University Hospital-Rikshospitalet and University of Oslo, 0027 Oslo, Norway § Proteomics Core Facility, Norwegian University of Life Sciences, 1432 Ås, Norway North East Proteome Analysis Facility (NEPAF), Devonshire Building, Newcastle upon Tyne, NE1 7RU, United Kingdom

)

‡

bS Supporting Information ABSTRACT: Recently, we introduced a novel approach for protein quantification based on isobaric peptide termini labeling (IPTL). In IPTL, both peptide termini are dervatized in two separate chemical reactions with complementary isotopically labeled reagents to generate isobaric peptide pairs. Here, we describe a novel procedure for the two chemical reactions to enable a cost-effective and rapid method. We established a selective N-terminal peptide modification reaction using succinic anhydride. Dimethylation was used as second chemical reaction to derivatize lysine residues. Both reactions can be performed within 15 min in one pot, and micropurification of the peptides between the two reactions was not necessary. For data analysis, we developed the force-find algorithm in IsobariQ which searches for corresponding peaks to build up peak pairs in tandem mass spectrometry (MS/MS) spectra where Mascot could not identify opposite sequences. Utilizing force-find, the number of quantified proteins was improved by more than 50% in comparison to the standard data analysis in IsobariQ. This was applied to compare the proteome of HeLa cells incubated with S-trityl-L-cysteine (STLC) to induce mitotic arrest and apoptosis. More than 50 proteins were found to be quantitatively changed, and most of them were previously reported in other proteome analyses of apoptotic cells. Furthermore, we showed that the two complementary isotopic labels coelute during liquid chromatography (LC) separation and that the linearity of relative IPTL quantification is not affected by a complex protein background. Combining the optimized reactions for IPTL with the open source data analysis software IsobariQ including force-find, we present a straightforward and rapid approach for quantitative proteomics.

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uantitative proteomics plays an important role in the comparison of biological systems, and different techniques for this purpose are under constant development. The incorporation of stable isotopes to achieve differentially labeled peptides allowing for a comparison of relative abundances is key in most mass spectrometry (MS)-based methods. Isotopic labels can be introduced metabolically, chemically, enzymatically, or via labeled synthetic peptides spiked into the sample. Several approaches have been developed to achieve the quantification on MS or tandem mass spectrometry (MS/MS). Stable isotope labeling by chemical reactions,1 enzymatic oxygen-18,2 or stable isotope labeling by amino acids in cell culture (SILAC)3 are widely used methods. The MS1 quantification is achieved by comparing the area or intensity of the precursor ions of peptides labeled with light and heavy isotopes, whereas the identification of the peptides is accomplished by the MS/MS spectrum. In contrast to SILAC, chemical and enzymatic labeling techniques can be applied to any protein-containing sample. However, this advantage comes at the price of an increased level of experimental noise, which results from the parallel processing of samples (protein level fractionation and digest) occurring prior to the introduction of the label.4 Furthermore, chemical labels with several deuterium atoms may produce a chromatographic shift during peptide separation. When two different states are compared, every peptide in the resulting proteolytic digest will appear r 2011 American Chemical Society

twice, and therefore the overall sample complexity is doubled at the MS level. Isobaric quantification techniques overcome this problem by shifting the quantitative information to the MS/MS level.5 The concept is to introduce chemical labels which result in isobaric peptides and comigrate in the liquid chromatography (LC) separation producing single peaks for isobaric peptide pairs. The quantitative ratio and origin can be differentiated upon peptide fragmentation in the MS/MS spectrum. To obtain the information for identification and quantification, only one precursor ion must be selected and fragmented. Furthermore, as the labeled peptides result in isobaric masses and are pooled, an increase in precursor ion intensity and decrease of the complexity in the MS trace compared to MS-based quantification methods is achieved. The common strategy of the isobaric methods isobaric tagging for relative and absolute quantification (iTRAQ),6 and tandem mass tagging (TMT)7 is the introduction of isobaric chemical tags attached to the amino groups of the peptides which result in reporter ions of distinct masses detected during MS/MS fragmentation. The reporter ions containing the quantitative information are detected in the low-mass region of the MS/MS Received: January 31, 2011 Accepted: April 29, 2011 Published: April 29, 2011 4775

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Analytical Chemistry spectrum and therefore limit the choice of MS analyzer that can be used with these techniques.5 Recently, a discussion about the dampening effect observed in iTRAQ and TMT has emerged.8,9 This phenomenon produces too low estimates of protein abundances when compared to the actual amount in the sample. We recently presented a novel quantification concept based on isobaric peptide termini labeling (IPTL) following a new strategy of isobaric labeling.10 The IPTL approach is based on crosswise chemical labeling of both peptide termini with complementary isotopically labeled reagents. As a result, all N- and C-terminal fragment ions will provide quantification data for each peptide throughout the MS/MS spectrum by forming peak pairs during fragmentation. The presence of several quantification points in each MS/MS spectrum is likely to increase the robustness of the quantification procedure. Furthermore, IPTL can be performed with any mass spectrometer as the quantification points are distributed through the whole mass range and no chromatographic shift of the peptides was observed as both peptides contain the same number of deuterium atoms. Here, we present a straightforward rapid-IPTL approach where the chemical reactions were optimized and resulted in a cost-effective rapid-IPTL method which incorporates both labels in a one-pot reaction. The new rapid-IPTL approach was first established with standard proteins and confirmed by the analysis of the complex proteome of HeLa cells exposed to the antimitotic inhibitor S-trityl-L-cysteine (STLC). For data analysis, we used the software tool IsobariQ11 and implemented force-find as a novel algorithm for improved quantitative data analysis.

’ METHODS Cell Culture, Induction of Mitotic Arrest and Apoptosis, SDSPAGE, and Lys-C Digestion. Cell culture, induction of

mitotic arrest (16 h), and apoptosis (40 h) by exposure to STLC, sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDSPAGE), and Lys-C digestion were performed as previously described.10 For the background experiment, a complex tryptic peptide mixture of Mycobacterium tuberculosis protein total lysate was prepared as previously described.12 N-Terminal Peptide Succinylation. A solution of 20 mM succinic anhydride (Sigma-Aldrich, Oslo, Norway) or tetradeuterated succinic anhydride-d4 (Larodan Fine Chemicals AB, Malm€o, Sweden) was freshly prepared in 100 μL of 50 mM sodium acetate buffer, pH 7.6. Then, 20 μL of succinic anhydride solution was added to the purified and dried Lys-C peptide digests. After thorough mixing, the sample was incubated for 5 min at room temperature. Dimethylation of Lysine Residues. 30 μL of 200 mM triethylammonium bicarbonate was directly mixed with the solution of N-terminal succinylated peptides. Then, 2 μL of 4% formaldehyde or dideuterated formaldehyde-d2 in water, respectively, was added and mixed before 2 μL of 600 mM sodium cyanoborohydride was added to each sample and incubated for 5 min. Subsequently, 8 μL of 1% ammonium hydroxide was added and incubated for 1 min before adding 4 μL of 5% formic acid. Finally, the samples were ZipTip μ-C18 purified, combined, and prepared for analysis by mass spectrometry. MALDI-TOF/TOF-MS. An Ultraflex II (Bruker Daltonics, Bremen, Germany) matrix-assisted laser desorption ionization tandem time-of-flight (MALDI-TOF/TOF) mass spectrometer was used after external calibration as previously described.13 NanoLCLTQ Orbitrap Mass Spectrometry. The dried peptides were dissolved in 10 μL of 1% formic acid, 2% acetonitrile

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in water, and 3 μL was injected into an Ultimate 3000 nanoLC system (Dionex, Sunnyvale CA, U.S.A.) connected to an LTQ Orbitrap XL mass spectrometer (ThermoScientific, Bremen, Germany). Column dimensions, LC buffer and gradient, and other instrument parameters were as previously described.14 Data Analysis Using DtaSuperCharge and Mascot. Raw LTQ Orbitrap XL data were processed using DtaSuperCharge15 version 1.37 to generate peak lists in *.mgf format. A database search was performed using Mascot in-house version 2.2.1 to search from Swiss-Prot (57.13) human entries (20 349 sequences). Lys-C as enzyme, no missed cleavage site, a tolerance of 10 ppm for the precursor ion and 0.6 Da for the MS/MS fragments, and methionine oxidation and N-terminal protein acetylation as variable modifications were selected. The two corresponding modifications, N-terminal succinylation/lysine dimethylation-d4 or N-terminal succinylation-d4/lysine dimethylation, respectively, were set as variable modifications for the complex data set. To estimate the N-terminal specificity of the rapid-IPTL chemistry, modifications were set as fixed in the following combinations: N-terminal succinylation/lysine succinylation, N-terminal succinylation-d4/lysine succinylation-d4, N-terminal dimethylation/lysine dimethylation, and N-terminal dimethylation-d4/lysine dimethylation-d4. To gain information about side reactions, additional succinylations and succinylationd4, respectively, at cysteines, serines, threonines, and tyrosine were allowed as variable modifications on top of the normal settings. Automatic decoy database searches were performed in Mascot and revealed a false discovery rate for peptide matches above an identity threshold of 1.94%, 1.96%, and 2.16%, respectively, for the three replicates of the Lys-C digested proteins of HeLa cells. IsobariQ Improvements. The force-find algorithm was developed to mine the spectra where Mascot failed to identify opposite sequences. IsobariQ first creates a virtual hit to the MS/ MS spectrum where the IPTL labeling is opposite to the Mascot identified sequence. This virtual dual-sequenced match is then quantified to see how many quantification points in the MS/MS spectrum could be detected. If the number of quantification points found in a spectrum is above a user-defined threshold, the opposite sequence is believed to be present. The ion score of such a force-found pair is not recalculated, in contrast to the pairs where both sequences were identified by Mascot. Quantitative Data Analysis Using IsobariQ. The import wizard of IsobariQ was used to load the data sets into the main module. No sequence suggestion below a Mascot ion score of 20 was allowed. Proteins were loaded if a minimum of two peptides and the significance of p < 0.05 were achieved. The minimal number of ratios for force-find hits was set to 4. The data were quantified with the following options: Protein scoring was set to MudPit and the MS/MS tolerance to 0.6 Da. Bold peptides (Mascot option) were required for the quantification, and unique and razor peptides were used. The “normalize rawfiles independently” checkbox was checked. Two rounds of quantification were performed, once including force-find hits and once without. The data were quantified using the VSN normalization method,16 and the results were exported to Excel.

’ RESULTS The Rapid-IPTL Approach. IPTL is based on crosswise isotopic labeling of both peptide termini which results in isobaric peptides after pooling. An outline of the rapid-IPTL approach is shown in Figure 1. First, the proteins are digested 4776

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Analytical Chemistry

Figure 1. Flowchart of rapid-IPTL. First, the proteins are digested with Lys-C. Second, the peptide R-N-termini are modified with succinic anhydride (succinyl, state A) and succinic anhydride-d4 (succinyl-d4, state B), respectively. Third, dimethylation of the free ε-amino group of lysines using sodium cyanoborohydride and formaldehyde is performed. Here, the peptides from state B are dimethylated with formaldehyde, whereas the peptides from state A are dimethylated with formaldehyded2. The pooled doubly labeled peptides result in isobaric masses, and corresponding peptides of the two states result in single peaks in MS mode acquisition. The relative quantitative abundance of the peptides derived from the two different states can be detected by the ion intensities of peptide fragment ions in the MS/MS spectrum, occurring in pairs with 4 Da mass shifts. Finally, IsobariQ can be used to determine the ratios of the relative peptide quantity after protein identification with Mascot.

with endoproteinase Lys-C to generate peptides with lysines at the C-terminal end. The peptide R-N-termini are subsequently modified with succinic anhydride in one state and succinic anhydride-d4 for the other state. The second chemical modification is a dimethylation of the free ε-amino group of lysines, performed using reductive amination with sodium cyanoborohydride and formaldehyde.17,18 The peptides modified with succinic anhydride-d4 are dimethylated using formaldehyde, whereas the peptides labeled with succinic anhydride are modified with formaldehyde-d2 to produce dimethyl-d4 at the C-terminus (Figure 1). The doubly labeled peptides with single lysines from both combined states resulted in isobaric masses, and the corresponding peptide pairs of the two states result in single peaks in MS mode. The relative quantitative abundance of the peptides derived from the two different states can be determined by the ion intensities of peptide fragment ions in the MS/MS spectrum, occurring in pairs with mass shifts of 4 Da. The N-terminal fragment ions such as b-ions of the peak pairs with lower masses are derived from labeling with succinic anhydride, and the corresponding higher masses are from succinic

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anhydride-d4. For C-terminal fragment ions such as y-ions, the lower mass ions of peak pairs are derived from dimethylated peptide fragments and the higher mass ions were peptide fragments modified with dimethyl-d4. Consequently, the relative ratio of a fragment peak pair gives reciprocal results for the different ion series and it is necessary to allocate peak pairs to ion species. One-Pot Reaction by N-Terminal Succinylation and Subsequent Dimethylation. The original IPTL approach has been performed with the specific labeling of the ε-group of lysines with methoxy-4,5-dihydro-1H-imidazole (MDHI) at pH 10.5 followed by derivatization of the R-amino terminal group using succinic anhydride at pH 6.8. Because of the difference in pH, both reactions cannot be combined in a one-pot reaction, and a micropurification step was required in between. To simplify this, we have searched for alternative chemical reactions using reagents which are available as isotopes deviating with at least 4 Da. First, we tested the reaction of peptides with succinic anhydride in different buffers and identified 50 mM sodium acetate enabling the specific labeling of only the N-terminus. As an example, the peptides of the Lys-C digest of human transferrin (Figure 2A) were selectively modified with succinic anhydride resulting in a mass shift of 100 Da of all peptides within 5 min (Figure 2B). Isotopic labeling by dimethylation is commonly used with a slightly higher pH than that employed for N-terminal succinylation. Therefore, the first reaction mixture for N-terminal succinylation was diluted with triethylammonium bicarbonate (pH 9.3) and dimethylation was completed within 10 min using formaldehyde and sodium cyanoborohydride. As shown in Figure 2C, the Lys-C peptides of N-terminal succinylated human transferrin have been further modified by dimethylation using formaldehyde-d2, resulting in an additional mass shift of 32 Da for all peptides. When the unmodified Lys-C digest of human transferrin was compared with the doubly modified digest, a mass shift of 132 Da was observed for all peptides (Figure 2, part A vs part C). Specificity of N-Terminal Succinylation. Succinylation is known to modify peptides at R (N-terminus) and ε (lysine) amino groups in phosphate butter at pH 78.19 However, we have tested different buffers for succinylation and found that R-amino groups are selectively modified using sodium acetate buffer at pH 7.6. To estimate the impact of side reactions for N-terminal succinylation and subsequent dimethylation of lysines, both reactions for rapid-IPTL of labeled transferrin were analyzed separately and masses of possible side reactions were checked. The most intense mass peak observed in the peptide mass fingerprint corresponded to the peptide YLGEEYVK with an unmodified single charged mass of m/z 1000.5 (Figure 2A) and resulted in the most intense peak after rapid-IPTL labeling with a mass of m/z 1132.6, modified with N-terminal succinylation/ lysine dimethylation-d4 (Figure 2D) or N-terminal succinylation-d4/lysine dimethylation (Figure 2E). The peak of the unmodified peptide at m/z 1000.5 Da disappeared completely in both spectra. Single (m/z 1100.5 and 1104.5, respectively) or double succinylation (m/z 1200.5 and 1208.5, respectively) were both not detected (Figure 2, parts D and E). Single dimethylation (m/z 1028.5 and 1032.5, respectively) was not observed either in both spectra. However, the presence of the peaks at m/z 1056.6 (Figure 2D) and 1064.6 (Figure 2E) indicated that a small proportion of peptides (0.9998 (Figure 4). Although the ratios were linear over the whole dynamic range when the data was quantified with the force-find algorithm (Figure 4A), the extreme ratios (20:1, 10:1, 1:10, 1:20) were not found when the data were quantified without the force-find option (Figure 4B). Effect of a Complex Background on Rapid-IPTL. As negative effects occurring from a complex background have been described on isobaric labeling techniques like iTRAQ and TMT,8,9 we compared a low-complexity standard against the same standard spiked with a complex peptide mixture to estimate a potential effect on rapid-IPTL. Different ratios (20:1, 5:1, 3:1, 1:1, 1:3, 1:5, and 1:20) of rapid-IPTL-labeled BSA were once analyzed as a neat sample and once mixed with a complex peptide mixture of an unlabeled M. tuberculosis protein lysate12 increasing the background complexity in the sample (Supplementary Figure 4 in the Supporting Information). To ensure MS/MS events for quantification, an inclusion list of the masses for the labeled BSA peptides was used for precursor ion selection. The quantitative analysis using IsobariQ revealed that no significant difference was found between the ratios acquired from the sample with and without matrix background (Figure 4C).

’ DISCUSSION Important benefits of isobaric labeling methods for peptide quantification in comparison to nonisobaric labeling methods are the preservation of the original sample complexity on the MS level. Furthermore, MS/MS spectra contain less noise than MS spectra, because the peaks exclusively originate from the isolated precursor mass window. Approaches for quantification using isobaric labels include either reporter-ion-based methods such as iTRAQ, TMT, DiLeu, CILAT or crosswise peptide terminal labeling using IPTL.5 An additional feature of IPTL is that the fragment ion series result in peak pairs with a common mass difference due to the double labeling which all can be used for quantification. Therefore, many quantification points are generated in each MS/MS spectrum, and consequently, a statistical assessment of the measured ratios is feasible. Furthermore, IPTL is not limited to certain mass analyzers and MS/MS fragmentation techniques due to the distribution of the ratios over the complete mass range in the MS/MS spectrum. For the originally published IPTL approach, MDHI/succinic anhydride-d4 and MDHI-d4/succinic anhydride, respectively, were used as reagents to accomplish isobaric-labeled peptides. Thereby, the first reagent (MDHI) labeled the peptides selectively at the ε-amino group of lysines at the C-terminus and succinic anhydride used as the second reagent labeled the remaining free R-amino groups at the N-terminus. Both reagents were available as isotopes with the same mass difference of 4 Da, chosen to avoid interference with the isotopic envelope of the molecules. However, both labeling steps required their specific buffer with different pH values which required a cleanup step between the reactions increasing sample handling time and potential experimental errors. With the development of rapid-IPTL, we showed that the N-terminal succinylation can be performed in a few minutes and combined with dimethylation in a one-pot reaction. This resulted in an improved IPTL approach for rapid and cost-effective quantitative proteome analyses.

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A prerequisite for IPTL is the selective modification of one peptide terminus with the first chemical reaction. Considering the reactivity of amino acids and cleavage reagents, amines are preferred targets for this purpose. Although a small difference of the pKa of R- and ε-amino groups exists,22 methods for the selective modifications of either R- or ε-amino groups of peptides are limited. MDHI and guanidination with O-methylisourea at high pH are effective reagents for modifications of the ε-amino group. However, 13C15N2-methylisourea provides only a 3 Da difference to the natural analogue, which can lead to interferences of the isotopic clusters.23 Specific labeling reagents to R-amino groups have been reported using nicotinyl-N-hydroxysuccinimide (Nic-NHS),24 phenyl isocyanate (PIC),25 and derivates of N-tris(2,4,6-trimethoxyphenyl)phosphonium bromide (TMPP).26 Using Nic-NHS, only partial labeling of the N-terminus of peptides was observed at pH 5.0 and the N-terminal specificity was lost at higher pH.24 Heavy isotopes with at least 4 Da difference to the natural molecule have not been reported for TMPP. PIC reacts quantitatively with peptide N-terminal amines at neutral pH, and the pentadeuterated reagent can be used for quantitative comparisons.25 However, N-terminal succinylation is a cost-effective alternative to these reagents and succinic anhydride is available as 2H4 and 13C4 isotopes, both suitable for relative quantification. Furthermore, reductive dimethylation using formaldehyde-d2 is an effective and relatively cheap isotopic-labeling reaction as a counterpart to the N-terminal succinylation in IPTL. Drawbacks of chemical labeling techniques are incomplete modifications and side reactions. However, we showed here that N-terminal succinylation can be performed with high yield and specificity (Figure 2). Furthermore, even though deuterated reagents are used to achieve the labeling, no chromatographic shift of the peptides in the LC run was observed (Figure 4). Rapid-IPTL was suitable for a complex proteome analysis using a human HeLa cell line incubated with STLC for 16 and 40 h. The antimitotic agent STLC blocks HeLa cells in the M phase of cell cycle and subsequently leads to apoptosis after 40 h of treatment. We identified 1054 proteins in at least two of the three replicates, but only 437 (41%) proteins could be quantified by IsobariQ with the common approach. To determine the correct ratio for a quantified peptide, the annotation of the fragment ions is mandatory due to the reciprocal results for N- and C-terminal ion pairs. Originally, IsobariQ was limited to the output of Mascot as the light and the heavy fragment ions must be annotated in order to find a peak pair. To overcome this limitation, the force-find algorithm was developed and included into IsobariQ. This algorithm enables IsobariQ to search for the counterpart of annotated peaks to build a set of quantifiable peak pairs. When this force-find functionality is applied, the quantification rate of our data set significantly increased from 41% to 64% as 678 of the 1054 identified proteins were quantified. Importantly, we have shown that extreme quantitative changes can only be determined reliably with force-find in IsobariQ. As peaks detected by force-find have not been annotated by Mascot, we implemented a user definable threshold into the algorithm that a minimum number of peaks must be found within a MS/MS spectrum in order to be accepted for quantification and to decrease the probability of finding random background peaks. Many proteomic studies merge the MS data of all bands in an SDSPAGE lane into one data set for quantitative analysis. Consequently, only one quantitative ratio per protein is obtained. Previously, we have used differential SDSPAGE gel 4780

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Analytical Chemistry migration of proteins to identify protease substrate on a proteomic scale,21 an approach that was substantially refined with the PROTOMAP methodology developed in Benjamin Cravatt’s laboratory.27 In this study, we reanalyzed our data using the detailed IsobariQ export information to requantify protein ratios into protein subgroups based on the information contained in a gel slice. Thus, a significant increase of regulated proteins was found even though these proteins’ overall abundance did not change. Many proteins are cleaved by caspases during apoptosis resulting in protein fragments with lower molecular masses, and consequently also the amount of the cleaved full-length protein is reduced. Actually, many known caspase substrates were found in our study using the gel slice information, such as, e.g., the cytoskeletal proteins filamin A, filamin B, lamin A/C, myosin-9, spectrin R-chain, spectrin β-chain, and vimentin. Consequently, quantifying every slice individually was a prerequisite in order to obtain a more complete view of relative changes, an information which might be lost if whole gel lanes are merged for data analysis. Recently, it was pointed out that reporter-ion-based isobaric labeling techniques can lead to inaccurate results for extreme (>5-fold) ratios in complex samples.8,16 As the quantification signals in IPTL are specific for each peptide, one would not expect that this ratio compression occurs in IPTL-labeled samples, independent of sample complexity. Indeed, a series of BSAcontaining samples in different dilutions yielded the same linear quantification covering a 20-fold range of concentrations in the presence and in the absence of a tryptic digest of M. tuberculosis. In conclusion, IPTL is based on crosswise isotopic labeling of peptide termini which results in isobaric peptides after pooling. With the development of N-terminal succinylation and subsequent dimethylation, we present an improved chemistry for the recently introduced isobaric quantification strategy IPTL. The reaction time and costs were significantly reduced in comparison to the original approach. Furthermore, we show that both chemical modifications can be performed successively in a one-pot reaction without an intermediate cleanup procedure reducing sample handling and thus the potential of experimental errors. We demonstrate that the rapid-IPTL approach is linear across a 20fold range of concentration ratios, both in the presence and in the absence of a complex proteome background. Furthermore, we show that rapid-IPTL can be applied to perform a proteomic study with a complex biological data set. In combination with the data analysis software IsobariQ, we were able to quantify 64% of the proteins applying the new force-find algorithm, increasing the number of quantified proteins by more than 50%.

’ ASSOCIATED CONTENT

bS

Supporting Information. Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*Phone: þ47-22840533. Fax: þ47-22840501. E-mail: bernd.thiede@ biotek.uio.no.

’ ACKNOWLEDGMENT The authors thank Gustavo de Souza (The Gades Institute, University of Bergen) for providing the M. tuberculosis samples.

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This work was supported by the FUGE, FUGE-Øst, and Inven2 to B.T. C.J.K. and B.T. are the inventors of “Quantitative proteomics using isobaric peptide termini labeling” which was made as employees of the University of Oslo and constitutes the basis for a patent application. A.T. is working for NEPAF, a contract proteome analysis facility funded by the regional development agency ONE Northeast and by the European Regional Development Fund.

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