C-Terminal Charge-Reversal Derivatization and ... - ACS Publications

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C‑Terminal Charge-Reversal Derivatization and Parallel Use of Multiple Proteases Facilitates Identification of Protein C‑Termini by C‑Terminomics Prasath Somasundaram, Tomas Koudelka, Dennis Linke, and Andreas Tholey* AG Systematische Proteomforschung & Bioanalytik, Institut für Experimentelle Medizin, Christian-Albrechts-Universität zu Kiel, Niemannsweg 11, 24105 Kiel, Germany S Supporting Information *

ABSTRACT: The identification of protein C-termini in complex proteomes is challenging due to the poor ionization efficiency of the carboxyl group. Amidating the negatively charged C-termini with ethanolamine (EA) has been suggested to improve the detection of C-terminal peptides and allows for a directed depletion of internal peptides after proteolysis using carboxyl reactive polymers. In the present study, the derivatization with N,N-dimethylethylenediamine (DMEDA) and (4-aminobutyl)guanidine (AG) leading to a positively charged C-terminus was investigated. C-terminal charge-reversed peptides showed improved coverage of b- and y-ion series in the MS/MS spectra compared to their noncharged counterparts. DMEDA-derivatized peptides resulted in many peptides with charge states of 3+, which benefited from ETD fragmentation. This makes the chargereversal strategy particularly useful for the analysis of protein C-termini, which may also be post-translationally modified. The labeling strategy and the indirect enrichment of Ctermini worked with similar efficiency for both DMEDA and EA, and their applicability was demonstrated on an E. coli proteome. Utilizing two proteases and different MS/MS activation mechanisms allowed for the identification of >400 C-termini, encompassing both canonical and truncated C-termini. KEYWORDS: C-terminus, modification, derivatization, mass spectrometry, EDC, NHS, HCD, CID, ETD, E. coli

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INTRODUCTION The formation of protein fragments as well as of N- and Cterminally truncated proteins by proteolytic processing represents an important irreversible post-translational modification. This mechanism increases the number of proteoforms present in a cell. These newly generated protein variants can exhibit different biological functions compared to their genetically encoded substrates.1,2 Thus, knowledge of the products of protease-catalyzed reactions and their exact cleavage sites are important to understand the biological function of proteolysis. In this respect, the identification of novel-formed N- and C-termini becomes essential. A number of methods are available to identify protein Ntermini 3−6 due to the rather straightforward chemical manipulation of primary and secondary (proline) amino groups. In contrast, the identification of protein C-termini is more difficult,7 mainly due to three reasons. First, proteincompatible methods for chemical derivatization are few in number, which is caused by the low reactivity of carboxyl groups. Therefore, an activation step is necessary, which directly creates the need for protection of free amino groups in the analyte mixture.8 Second, a directed derivatization of protein carboxy terminus in the presence of aspartate and glutamate residues is not straightforward. Third, the ionization efficiency of C-terminal peptides, especially those resulting from tryptic digests devoid of basic residues (lysine, arginine, © XXXX American Chemical Society

and histidine), is reduced compared to basic peptides. This disfavors the ionization of C-terminal peptides compared to coeluting peptides with higher ionization efficiencies.9,10 A possible way to circumvent the latter was introduced recently by Huesgen et al., who presented a novel metalloprotease cleaving N-terminal to lysine and arginine. In comparison to trypsin, this so-called LysargiNase aids in identifying C-termini because proteolysis delivers a C-terminal peptide with an additional basic group at its N-term.11 However, for certain physiological questions, a C-terminal tag is still the better choice by which to perform positional proteomics (e.g., in vivo C-terminal proteolysis events attained from unknown endogenous proteases).12 An alternative strategy that conserves C-termini rather than altering their chemical structure is the so-called combined fractional diagonal chromatography (COFRADIC).13,14 This approach uses the chromatographic isolation of C-terminal from internal peptides, which facilitates their detection. A potential approach to enhance the ionization efficiency of peptides is to introduce positively charged groups by means of chemical derivatization. Although the introduction of fixed positive charge at the N-terminus has been used to facilitate the analysis of MS/MS spectra,15 the so-called charge-reversal Received: February 18, 2016

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DOI: 10.1021/acs.jproteome.6b00146 J. Proteome Res. XXXX, XXX, XXX−XXX

Article

Journal of Proteome Research

rotary vacuum centrifuge prior dissolving in 180 μL of 2-(Nmorpholino)ethanesulfonic acid hydrate (MES, 100 mM, pH 5) and divided into 12 aliquots. An equal volume of particular amine (EA, DMEDA, or AG) (75 mM buffered in MES (400 mM, pH 5)) was added, and carboxyl amidation was initiated by adding EDC (50 mM) and NHS (10 mM) dissolved in (i) H2O, (ii) 25% ACN, or (iii) 25% DMSO. After being incubated for 16 h at 25 °C, the reaction solution was acidified to pH 3−4 with 0.1% formic acid (FA). The derivatized peptides were desalted by SPE using Pierce C18 tips and eluted into 70% ACN/0.05% trifluoroacetic acid (TFA). The samples were dried under vacuum and reconstituted in 3% ACN/0.1% TFA to result a 1 μM peptide solution. b). Derivatization of Intact Proteins and Enrichment of C-Termini. Primary amine groups of the reduced and alkylated proteins were reductively dimethylated using formaldehyde (20 mM) and NaBH3CN (20 mM) for 16 h at 25 °C. Afterward, acetone precipitation was performed by adding five volumes of acetone (precooled to −20 °C). The protein pellet was gently washed with acetone (−20 °C) and centrifuged, the supernatant discarded, and the pellet dried under the fume hood. The dried proteins were reconstituted in 120 μL of MES (100 mM) and GndHCl (4 M, pH 5) and divided into eight aliquots. The amidation reaction was initiated as described above, but EDC and NHS stock solutions were prepared in (i) H2O or (ii) 75% DMSO. After incubating for 2 h at 25 °C, fresh EDC and NHS was added followed by a concluding 14 h incubation at 25 °C. In additional experiments using an equimolar mixture of β-casein, β-lactoglobulin, and BSA, the amidation reaction was performed using (i) acetone, (ii) isopropanol (i-PrOH), (iii) ethanol (EtOH), or (iv) methanol (MeOH) with final concentrations of 10% and 30%, respectively. Furthermore, the reaction was performed in the above-mentioned aqueous buffer with 0.25% or 0.5% of the nonionic detergents NP-40 or Rapigest added. Prior to digestion, a buffer exchange was performed by protein precipitation (acetone). After dissolving the dried proteins in HEPES (100 mM, pH 7.5) and GdnHCl (2 M), we diluted the GdnHCl concentration to 0.5 M by adding HEPES (100 mM). Trypsin (1 μg) was added and incubated for 16 h at 37 °C. Free amino groups of neo-N-termini were reductively dimethylated with formaldehyde (20 mM) and NaBH3CN (20 mM) for 6 h at 37 °C. Desalting via Pierce C18 tips was performed with buffers containing HCl rather than FA or TFA. The eluted peptide solutions were divided into two aliquots for samples defined as samples “before depletion” and “after depletion” analysis and dried under vacuum. The “before depletion” samples were dissolved with 3% ACN/0.1% TFA to obtain a 5 μM peptide solution. The “after depletion” samples were reconstituted in filtered polyallylamine solution (PAA) and sonicated shortly for 30 s. Note: the PAA solution required filtering because minimal amounts of coeluted short-chained PAA molecules caused high back-pressures on the trapping column of the LC systems. The crude PAA solution (200 μM in MES (50 mM, pH 5)) was filtered using 10 kDa Amicon membrane cutoff filters (Merck Millipore, Darmstadt, Germany) and washed twice with MES (50 mM, pH 5). The condensation reaction of the neo-Ctermini to the amine-rich resin was initiated by adding EDC (100 mM) and NHS (10 mM) dissolved in (i) H2O or (ii) 75% DMSO in two steps: after a 1 h incubation at 25 °C, fresh EDC and NHS were added, and the reaction was continued for 15 h. Final concentrations of EDC and NHS were 100 and 10 mM,

strategy introduces a positively charged moiety at the Cterminal carboxyl group. This approach has been used to enhance the signal intensities of peptides in MALDI MS experiments (e.g. by applying oxazolone-based chemistry).16,17 For ESI MS, charge-reversal derivatization has, to date, mainly been used to enhance charge states of peptides to improve electron-transfer dissociation (ETD) peptide fragmentation.9,18 In this study, we investigated the charge-reversal derivatization method at the peptide and protein level to analyze protein C-termini. For the later, we applied our strategy to a Cterminomics workflow, recently presented by Schilling et al.,8 which involves the indirect enrichment of C-terminal peptides by depletion of the proteolytically generated internal peptides. In contrast to the original publication, where ethanolamine (EA) was used to amidate the carboxyl groups, we tested two more basic amines, namely N,N-dimethylethylenediamine (DMEDA) and (4-aminobutyl)guanidine (agmatine, AG) (Supplementary Figure 1). The charge-reversal derivatization resulted in more complete y- and b-ion series spectra compared to EA. The increased basicity of DMEDA labeled C-termini resulted in higher charge states, which is beneficial for ETD fragmentation. The potential of the developed approach was shown by the analysis of an Escherichia coli proteome.

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EXPERIMENTAL PROCEDURES

Chemicals

Endoprotease trypsin was purchased from Promega (Madison, WI), and cOmplete Protease inhibitor cocktail was from Roche (Penzberg, Germany). All other proteins, chemicals, and solvents were from Sigma-Aldrich (Steinheim, Germany). Rapigest and Sep-Pak C18 Vac cartridges were from Waters (Milford, MA), Pierce C18 tips were from Thermo Fisher Scientific (Waltham, MA), Amicon Ultra-0.5 centrifugal filter devices were from Merck Millipore (Darmstadt, Germany), and VWR centrifugal filter devices were from VWR International (Rednor, PA). NP-40 was purchased from New England Biolabs. Deionized water (18.2 MΩ/cm) was prepared by an arium611 VF system (Sartorius, Göttingen, Germany). Sample Preparation

Bovine serum albumin (BSA) and bovine β-casein were dissolved in 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES, 100 mM, pH 7.5) and guanidine hydrochloride (GdnHCl) (2 M) at a concentration of 100 pmol/μL. A total of 10 nmol of an equimolar mixture of both proteins were denatured, and disulfide bonds were reduced with dithiotreitol (DTT) (10 mM) for 1 h at 56 °C. The reduced sulfhydryl groups were alkylated iodoacetamide (IAA, 20 mM) for 1 h at 25 °C in the absence of ambient light. Residual IAA was quenched with DTT (10 mM) for 15 min at 25 °C. A pair of different experiments using this stock solution were performed, one using digested proteins (a) and the second using intact proteins (b). a). Derivatization of Digested Protein. Protein stock solutions were diluted with one volume of H2O to reduce the concentration of guanidine, and proteolysis was performed by adding 1 μg of trypsin for 16 h at 37 °C. Subsequently, primary amines were reductively dimethylated with formaldehyde (20 mM) and NaBH3CN (20 mM) for 3 h at 37 °C. The reaction solution was acidified with HCl (5 M) to pH 3−4 and desalted by solid-phase extraction (SPE) using Sep-Pak C18 SPE cartridges. Peptides were washed with HCl (10 mM) and eluted with 70% ACN/HCl (10 mM). Samples were dried in a B


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was achieved by filtration over 10 kDa VWR centrifugal filters and Sep-Pak cartridges. After being freeze-dried, the peptides were dissolved in 100 μL of 3% ACN/0.1% TFA.

respectively. Separation of unbound peptides was achieved by filtration over 3 kDa VWR centrifugal filters with two washing steps (H2O). After acidifying to pH 3−4 with TFA (10%), we performed desalting with Pierce C18 tips as described above. After being dried under vacuum, the peptides were dissolved in 3% ACN/0.1% TFA to yield a 10 μM peptide solution.

LC−MS/MS Analysis

Sample sets of ß-casein/BSA experiments were analyzed on a Dionex Ultimate 3000 nano-UHPLC RSLC system coupled to a Q Exactive Plus mass spectrometer equipped with a nanospray ion source (Thermo Fisher, Bremen, Germany). A total of 1 μL of the samples derivatized at the peptide level was injected. Due to potential sample loss during the multiple buffer exchange and cleanup steps, 5 μL of the samples derivatized at protein level were injected. Samples were loaded and washed on a trap column (Acclaim Pepmap 100 C18, 10 mm × 300 μm, 3 μm, 100 Å, Dionex) for 5 min with 3% ACN/0.1% TFA at a flow rate of 30 μL/min. Peptide separation was performed using an Acclaim PepMap 100 C18 analytical column (50 cm × 75 μm, 2 μm, 100 Å, Dionex) with a flow rate of 300 nL/min: eluent A (0.05% FA) and eluent B (80% ACN/0.04% FA); linear gradient 5−50% B in 45 min, 50−90% B in 5 min, 90% B for 10 min, 90−5% B in 0.1 min, and equilibrating at 5% B for 10 min. Ionization was performed with 2.4 kV spray voltage applied on a noncoated PicoTip emitter (10 μm tip size, New Objective, Woburn, MA) with the source temperature set to 250 °C. MS data were acquired from 5 to 75 min with MS full scans between 300 and 2000 m/z at a resolution of 70 000 at m/z 200 (automatic gain control (AGC) value of 3e6 and maximum ion injection time (IIT) of 100 ms). The 10 most intense precursors with charge states ≥2+ were subjected to fragmentation with HCD (normalized collision energy (NCE) of 28%; isolation width of 3 m/z; resolution, 17 500 at m/z 200; AGC target of 1e5 and maximum IIT of 100 ms). Dynamic exclusion for 30 s was applied with a precursor mass tolerance of 10 ppm. Lockmass correction was performed on the basis of the polysiloxane contaminant signal of 445.120025 m/z. E. coli proteome samples were analyzed initially on an Ultimate 3000 nano-HPLC coupled online to a LTQ Orbitrap Velos mass spectrometer using a nanospray ion source and XCalibur 2.1 software suite (Thermo Fisher Scientific, Bremen, Germany). Precolumn trapping and separation (Acclaim PepMap 100 C18, 15 cm × 75 μm, 3 μm, 100 Å, Dionex) were performed with identical eluent composition, as described above. The chromatographic parameters were as follows: 5− 10% B in 1 min, 10−40% B in 90 min, 40−95% B in 5 min, 95% B for 10 min, 95−5% B in 0.1 min, and equilibrating at 5% B for 10 min. Ionization was performed with 1.25 kV spray voltage applied on metal-coated PicoTip emitter (30 μm tip size, New Objective, Woburn, MA) and source temperature set to 250 °C. MS data were acquired from 5 to 115 min with MS full scans between 300 and 2000 m/z at a resolution of 60 000 at m/z 400 (AGC target of 3e6; maximum IIT of 1000 ms). For each of the four samples, independent LC−MS experiments were performed with the following MS/MS settings: (i) the five most intense precursors with charge states ≥2+ were subjected to fragmentation with HCD (FTMS; NCE of 40%; isolation width of 3 m/z; resolution, 7500 at m/z 400; AGC target of 1e5 and maximum IIT of 500 ms), (ii) the five most intense precursors with charge states ≥3+ were selected for ETD fragmentation (ITMS; isolation width of 3 m/z; with charge-state-dependent ETD time, reaction time 150 ms, with supplemental activation; isolation width, 2 m/z; AGC target of 1e6 and maximum IIT of 50 ms), and (iii) the 10 most intense

Labeling, Digestion, and C-Termini Enrichment of an E. coli Proteome

a). Protein Extraction and Protein Concentration Determination by BCA Assay. Cytosolic proteins from E. coli (K-12 MG1655, cultured in M9 minimal medium with glucose (15 mM) and harvested at an optical density of 1) were extracted in lysis buffer (HEPES (100 mM, pH 7.4), NaCl (100 mM), and 2× cOmplete protease inhibitor. Glass beads (200 mg, 1:1 w/w of glass beads with Ø 0.1−0.25 mm and Ø 0.25− 0.5 mm diameter; Sigma-Aldrich; washed with 10% acetic acid, twice with water, and precooled to 4 °C) were added to the cell suspension. Cell disruption was performed in five cycles of freezing (30 s, −80 °C ethanol bath) and partial thawing in a sonication bath (45 s, on ice), followed by shaking in a mixer mill (Retsch, MM 400) (3 × 10 s, 30 Hz). The glass beads and cell debris were separated by centrifugation (15 min, 45000g, 4 °C) and the protein concentration determined using the bicinchoninic acid protein assay. b). Protein Denaturation, Disulfide Reduction, Thiol Alkylation, and Amine Protection. The cell lysate was freeze-dried and reconstituted in HEPES (100 mM, pH 7.5) and GdnHCl (4 M). After the dilution of the buffer by adding an equal volume of water, the reduction and alkylation of disulfide bonds was performed with DTT (40 mM) and IAA (80 mM) and residual IAA was quenched with DTT (20 mM). Free amino groups of lysine side-chains and N-termini were reductively dimethylated with formaldehyde (40 mM) and NaBH3CN (40 mM) for 6 h at 37 °C. Buffer exchange was performed by acetone precipitation; however, here the protein pellet was washed with methanol. c). Amidation of Carboxyl Groups. Reconstitution of the dried proteins was attained in MES (100 mM) and GdnHCl (4 M, pH 5) and divided into two aliquots. Both amidating carboxyl groups in the presence of EA or DMEDA and subsequent precipitation of the amidated proteins was conducted as described above. d). Proteolysis and Protection of Neo-N-Termini. Both protein pellets were reconstituted in 100 μL of HEPES (20 mM, pH 7.5) and GdnHCl (2 M) and divided both labeling states further each into two aliquots. After the dilution of the GdnHCl concentration to 0.5 M with HEPES (20 mM), proteolysis was initiated by adding 4 μg of either trypsin (in 1 mM HCl) or chymotrypsin (in 1 mM HCl and 2 mM CaCl2) followed by an incubation for 12 h at 37 °C. Generated neo-Ntermini were reductively dimethylated with formaldehyde (20 mM) and NaBH3CN (20 mM) overnight at 37 °C. e). C-Termini Enrichment by Polymer-Based Depletion of Internal Peptides. A 200 μM stock solution of crude PAA was prepared in MES (200 mM), GndHCl (2 M), and 20% ACN and filtered using 10 kDa Amicon filter devices as described above. Dried peptides were dissolved in 180 μL of filtered PAA solution, and the condensation reaction was initiated by adding EDC (100 mM) and NHS (20 mM) dissolved in the same buffer. After 1 h of incubation at 25 °C, fresh EDC and NHS was added, and the reaction was continued for 23 h at 25 °C. Separation of unbound peptides C


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Figure 1. Derivatization efficiencies analyzed for the proteins β-casein and BSA. (A) On the peptide level: the protein mixture of β-casein and BSA was derivatized with EA, DMEDA, or AG in parallel after proteolysis under aqueous, ACN-, or DMSO-containing buffer conditions. Only peptides identified from a minimum of two out of three technical replicates with an average standard deviation lower than 30% were considered into the calculations. (B,C) Comparison of average derivatization efficiencies under aqueous or DMSO-containing buffer conditions of (B) internal peptides and (C) C-terminal peptides after the derivatization of protein mixture of β-casein and BSA on protein level and digestion.

Proteome Discoverer output files (in .txt format) are accepted (among others). For the access of the information about the Cand N-terminus of a protein, the.fasta and.xml of each identified protein is downloaded automatically from UniProt. These files are used to compare the peptide sequence of interest (identified by Proteome Discoverer) to the C- and N-terminus of the protein (information obtained from UniProt). Finally, the following output is appended to the original input file: (i) peptide start and end sequence within the protein, (ii) Nterminus with signal peptide sequence, (iii) N-terminus without signaling peptide, and (iv) C-terminus. Items (ii)−(iv) were exported as boolean operators. Venn diagrams were generated using the Venn Diagram Plotter (Pacific Northwest National Laboratory; http://omics. pnl.gov/software/VennDiagramPlotter.php) and the online tool Venny (Oliveros, J. C., http://bioinfogp.cnb.csic.es/ tools/venny/).

precursors with charge states ≥2+ were selected for CID fragmentation (ITMS; NCE of 35%; isolation width, 2 m/z; AGC and maximum IIT identical to ETD parameters). Dynamic exclusion for 90 s was applied with a precursor mass tolerance of 10 ppm with lockmass correction (445.120025 m/z). Additionally, we analyzed the same samples on a Dionex Ultimate 3000 nano-UHPLC RSLC/Q Exactive Plus system as described above but with minor altered conditions: trapping of peptides was performed for 4 min, and elution was attained by a linear gradient from 5 to 40% B in 90 min and 40−90% B. MS data were acquired from 10 to 115 min. Other MS2 parameters were identical to those described above. Data Analysis and Interpretation

Database searches were performed with Proteome Discoverer 1.4.1.14 using the search algorithm SequestHT (both Thermo Fisher). Enzyme settings were semispecific trypsin or chymotrypsin with a maximum of two missed cleavages. Carbamidomethylation of cysteine thiol groups and dimethylation of lysine-ε-amino groups were set as static modifications. Oxidation of methionines, dimethylation of N-termini, amidation (with specific amine) of aspartates, glutamates, and C-termini were set as variable modifications. For the E. coli data set, modifications on Asp and Glu were set as static. The precursor mass tolerance was 10 ppm, fragment mass tolerances were 0.02 Da (HCD in Orbitrap (LTQ Velos and Q Exactive)) and 0.5 Da (CID and ETD in Iontrap (LTQ Velos only)). Peptide spectrum match (PSM) validation was performed by Percolator (default values). Protein and peptide lists were exported as text files using only high-confidence matches with a FDR ≤ 0.01, with a minimum of one unique peptide per protein. Derivatization efficiencies were calculated as ratio of the peak area of fully derivatized peptides divided by the sum of peak areas of all corresponding partial and fully derivatized peptide species. To differentiate C-terminal peptides from internal ones, we developed a GAMBAS3 script in-house. As input data,

Raw Data Repository

All mass-spectrometry proteomics data (E. coli experiments) have been deposited to the ProteomeXchange Consortium19 via the PRIDE partner repository with the data set identifier PXD003342. Instrument data were converted to mzIdentML files using ProCon as a conversion tool.20

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RESULTS AND DISCUSSION The aim of this study was to test the compatibility of different derivatization conditions and reagents with an established Cterminomics approach and to study the MS and MS/MS behavior of the derivatized peptides. Agmatine (AG), a decarboxylated analogon of arginine, and the asymmetric dimethylated amine N,N-dimethylethylenediamine (DMEDA) were used, and the success in identifying modified C-termini was compared to ethanolamine (EA), which yields uncharged C-termini. A second goal was to introduce the use of complementary proteases into a C-terminomics workflow used for the analysis of an E. coli proteome. D


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Journal of Proteome Research Peptide-Level Modification

efficiencies for internal and C-terminal peptides are shown in panels B and C of Figure 1, respectively. Derivatization in aqueous solutions led to moderate modification efficiencies: for Asp and Glu of internal peptides, an average yield of 46.1% was determined using EA as coupling reagent, while for DMEDA, a labeling efficiency of 54% was achieved. Protein C-terminal peptides were found to be derivatized on their C-terminal amino acid and in present on Asp and Glu side-chains, with an average of 73% in case of EA and with 87% efficiency using DMEDA. In accordance with the results obtained at the peptide level, AG, with 33%, exhibited the lowest efficiency for the internal peptides, while, interestingly, the C-terminal peptides showed a comparable high-derivatization yield of 79%. The addition of DMSO improved the labeling efficiencies on internal peptides compared to those in aqueous solutions: in the case of EA, an average yield of 58% was achieved, while DMEDA showed the highest improvement with a reaction completeness of 69%. The labeling efficiency with AG was also improved to 52% with the use of DMSO, which is comparable to EA in this case. Meanwhile, the derivatization efficiencies for the C-terminal peptides were slightly reduced for EA and AG in the presence of DMSO. To test potential increase of yields of derivatization by changing other reaction conditions, we performed several control experiments. For example we performed the amidation reaction using higher contents of organic solvents (e.g. acetone, i-PrOH, EtOH, and MeOH with concentrations of 10% and 30%, respectively). Indeed, a recent publication of Zhang et al. delivered hints for an improved derivatization upon the increase of organic solvent content of derivatization buffer.10 However, the achieved derivatization efficiencies (28−64% for internal and 46−80% for C-terminal carboxyl groups (Supplementary Figure 2A) were similar to those we observed in aqueous buffers in our experiments. Furthermore, we investigated the influence of nonionic detergents (NP-40 and Rapigest). These detergents should both help denaturing the protein structures, thus increasing the accessibility of otherwise shielded carboxyl groups, and secondly aid in keeping the proteins in solution. For internal carboxyl groups, no significant improvement was achieved (40−64%) compared to organic solvents (Supplementary Figure 2B). The derivatization efficiencies for Cterminal carboxyl groups were even decreased (36−44%). Additionally, the overall protein sequence coverages were lower for detergent treated samples. The latter is potentially caused by the still-insufficient removal of the detergents by precipitation and by sample loss encountered with this process. Conclusively, lower labeling efficiencies were observed at the protein level when compared to the peptide level, probably as a result of remaining secondary and tertiary protein structure, which is likely to be less prevalent at the peptide level. This is also supported by the fact that protein termini were labeled with higher efficiencies than internal peptides. For example, we observed that both the C-terminal α-carboxyl group as well as side-chain functionalities of Asp and Glu residues were more efficiently derivatized (Figure 1C) compared to internal sidechain carboxyls (Figure 1B). The lower derivatization efficiency of intact proteins is not unexpected. Even derivatization methods usually delivering high yields at peptide level (e.g. reductive dimethylation or Nhydroxysuccinimide activation-based labeling of amino groups) are in most cases incomplete when performed at the protein level.24 Whereas for the amino-directed derivatization reactions (e.g., the NHS-chemistry-based methods), the molecules to be

A prerequisite for labeling procedures are high-derivatization yields and the avoidance of side reactions. Due to the low reactivity of the carboxyl function, an activation step has to be applied, enabling the reaction with a free amine to form an amide bond. In our study, we used the EDC−NHS-based activation.21 To prevent unwanted cross-reactions with free amino groups, it is necessary to block these prior to derivatization. For this purpose, we chose the well-established reductive dimethylation protocol for the derivatization of Nterminal and ε-amino groups (Lys), both at the peptide and at the intact protein level.22 Different derivatization conditions were tested using tryptic digests of two model proteins, β-casein and BSA. Derivatization efficiencies with EA, DMEDA, and AG were calculated using the precursor ion areas obtained from proteome discoverer. The yield was determined by dividing the precursor ion area of the completely labeled peptide species (including modifications of side chains of Asp and Glu) by the sum of the precursor ion areas of the fully and partially labeled species. If no fully labeled species was observed, the yield was given a zero value (Supplementary Table S1). Carboxyl-directed labeling of peptide mixtures in aqueous solutions showed good derivatization efficiencies, with an average yield of 82% using EA and 87% for DMEDA (Figure 1A) In contrast, peptide modification with AG led to an average derivatization efficiency of 28%, which might be due to steric reasons. EDC and other carbodiimides undergo rapid hydrolysis in aqueous solutions,23 which may lead to reduced derivatization yields. Therefore, we modified the established derivatization protocol, which is usually conducted in aqueous buffer solution, by applying EDC and NHS dissolved in up to 80% ACN or DMSO. These stocks were added as a 1:1 mixture (v/v) to the peptides. The final concentration of organic solvent was ∼30%. Compared to results in purely aqueous conditions, higher derivatization efficiencies were achieved with ACN containing reaction media. In the case of EA, peptides were modified with an average yield of 88%, while 92% efficiency was achieved with DMEDA. Interestingly, AG derivatization was also improved by applying ACN to the reaction media. However, with an average efficiency of 48%, the derivatization with this reagent was still less efficient compared to EA and DMEDA. For DMSO, the derivatization efficiency also increased compared to aqueous conditions, but the effect was less than for ACN. Protein-Level Modification and Enrichment of C-Termini

For the application of the charge-reversal strategy of protein Ctermini in complex mixtures, the derivatization at the intact protein level rather than at the peptide level is necessary. Here a number of additional factors affecting the derivatization become important (e.g., the limited accessibility of the C-termini caused by protein tertiary or quaternary structures). To investigate protein level derivatization efficiencies, we used an equimolar mixture of ß-casein and BSA (Supplementary Table S2). For reductive dimethylation, control experiments performed in aqueous buffer revealed an average derivatization efficiency of 93%. Because the derivatization efficiencies for carboxyl groups at peptide level was shown to be more efficient in aqueous buffers containing organic solvents, we also tested the influence of the later on intact protein carboxyl derivatization. In initial experiments, we observed precipitation even at relatively low concentrations (∼30%) of acetonitrile. Therefore, we only tested the addition of DMSO. The derivatization E


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Figure 2. MS/MS-spectra of the C-terminal peptide LVVSTQTALA of BSA. Peptide species N-terminally dimethylated and carboxyl-amidated with (A) EA, (B) DMEDA, or (C) AG were subjected to fragmentation using the HPLC-Orbitrap Velos by CID (ITMS, NCE of 35%) and HCD (FTMS, NCE of 40%): (D) EA, (E) DMEDA, or (F) AG, respectively.

BSA with the different amides (Supplementary Figure 3). Comparing the sum of all peptide peak areas assigned to BSA before and after depletion, we observed an approximate reduction in the overall peptide peak intensities by 2 orders of magnitude, which mainly results from the depletion of internal peptides. This procedure was successful regardless of the amine or the work protease used. However, independent from the derivatization strategy applied, we observed high backpressures on the trapping column of our LC systems after using the high amounts of PAA suggested in the original protocol.8 To circumvent this problem, we performed the enrichment steps with 10% of the recommended PAA concentration, which was clarified via a 10 kDa Amicon centrifugal filtration device prior to use to deplete shorter PAA chains from the polymer solution. This procedure reduced the problems of precolumn clogging when charge-reversal derivatization was applied in aqueous buffer. However, in the presence of DMSO again, high back-pressures on the trapping column were observed. We assume that DMSO increases the permeability of residual short and medium molecular PAA chains through the cellulose membranes of the used Microcon centrifugal filters.

attached to the protein can be preactivated, in the case of carboxyl group derivatization, the activation occurs at the protein first, followed by the addition of the molecule to be conjugated. It has been shown that the activation of protein carboxyl functions by EDC can lead to the formation of Nacylurea side products (Timkovich).25 This can lead to additional problems reducing the overall yields of derivatization. Although the incomplete derivatization of protein C-termini is still a problem to be solved, a more important factor is the impact of the charge-reversal strategy on the identification of Cterminal peptides in comparison to their uncharged or even completely underivatized counterparts. Interestingly, the Ctermini of β-casein and BSA were identified with more peptide spectrum matches (PSM) using DMEDA or AG. This can be explained by the fact that these more basic reagents induce additional charges to the C-terminal peptide, which improves their ionization efficiency and therefore increases their signal intensity compared to the signals from internal peptides with underivatized neo-C-termini. However, internal peptides containing Asp or Glu residues also showed increased ionization efficiency after derivatization with DMEDA or AG. Therefore, to benefit from the increase in ionization efficiency of chargereversed C-terminal peptides, enrichment of these peptides, or vice versa (depletion of internal peptides), can be applied. Moreover, enrichment is necessary to reduce the number of internal peptides, which hamper the identification of the protein C-termini in complex samples.8 The use of high molecular weight (above 50 kDa) linear polyallylamine (PAA) polymers has been suggested to bind internal peptides covalently via their free C-termini to the polymers amino groups.8 Thus, we compared the enrichment efficiencies for Cterminal peptides using the PAA resin on the model protein

MS and MS/MS Behavior of Amidated Peptides

C-terminal charge-reversal of peptides altered their behavior in the mass spectrometry in both MS and MS/MS mode. It is in agreement to previous ESI-MS experiments involving chargereversal derivatization investigated by Frey and co-workers, who also observed an increase in signal intensities of derivatized peptides compared to underivatized peptides.18 A comparison of the intensities of the extracted ion chromatograms of Cterminal peptides, derivatized either by ethanolamine or DMEDA at peptide level using only BSA as model protein, showed no clear trends. In many cases (for example, for the Cterminal peptide of BSA (Supplementary Figure 4)), a F


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Figure 3. Comparison between identified internal peptides and truncated and canonical C-terminal peptides of E. coli after derivatization and indirect enrichment of C-terminal peptides. All measurements were conducted on the HPLC Orbitrap Velos platform (n = 3).

observed for charge-reversed peptides when the a1-ion was within the mass range (e.g., above 100 m/z for FTMS).

significant increase of intensity for the basic group DMEDA could be observed compared to its EA-derivatized counterpart; a similar effect was observed for agmatine. However, this is not a general phenomenon. In more complex mixtures (e.g., the E. coli proteome described below; data not shown), we also observed lower intensities for peptides carrying charged reversal modification compared to those with the uncharged ethanolamine modification. This is most likely influenced by the physicochemical properties and the number of coeluting peptides. Additionally, the more basic functions in the derivatized peptides can also cause a broader charge-state distribution.10 However, this effect can be beneficial when ETD fragmentation is used, which favors charge states of ≥3+.26 For MS/MS spectra, the results showed a clearer trend. In tandem MS using the Orbitrap Velos, fragmentation behavior was dependent on the activation type applied. For example, activation of the doubly charged precursor of the C-terminal peptide LVVSTQTALA of BSA by resonance excitation in the linear ion-trap and collision-induced decomposition (CID) showed poor spectral quality and yielded mainly higher b-ions when the peptide was modified with EA (Figure 2A). Upon introduction of the basic DMEDA (Figure 2B) and AG (Figure 2C) moieties, nearly complete b- and y-ion series were observed. This was in contrast to beam-type activation of the same peptide using higher-energy collisional decomposition (HCD) in the dedicated HCD octopole, which yielded mainly y-ions for the charge-reversed peptides (Figure 2E,F), whereas for the EA derivative, predominantly b-ions were observed (Figure 2D). Similar observations for an instrument dependent fragmentation behavior were previously described for tryptic peptides, which (aside from the basic N-terminus) contain a basic group at the C-terminus in the form of a Lys- or Argresidue.27−29 Although b-ions are generally underrepresented in beam-type activated CID experiments using quadrupole qTOFinstruments, this effect was even stronger in case of HCD fragmentation in the Orbitrap Velos. For reductively dimethylated peptides the b1-ions have been shown previously to lose carbon monoxide to yield a more stable alkylated immonium a1-ion during tandem MS.30 This characteristic feature was also

C-Terminome Analysis of E. coli

To elucidate the potential benefits of the charge-reversal strategy for the C-terminome analysis, we performed the derivatization and enrichment procedure on a cytosolic proteome extract originated from E. coli. In this experiment, only EA and DMEDA were compared; AG was omitted because the derivatization efficiencies were significantly lower with this amine. After proteome extraction, proteins were first reductively dimethylated and then derivatized under aqueous conditions, as described above, due to the problems encountered with the use of DMSO and subsequent polymer-based enrichment. To cover a broad range of C-terminal peptides, we performed tryptic and chymotryptic digests of the labeled proteome in parallel. The C-terminal peptides were enriched by the depletion of the internal peptides as described above. Initially, we analyzed the enriched peptide fraction using nano-HPLC coupled to a LTQ Orbitrap Velos with ETD support. We applied three different fragmentation techniques (CID, HCD, and ETD) to cover as many different physicochemical peptide properties as possible and to determine whether the fragmentation type, the work protease, and the amine used for labeling had an influence on peptide identifications. Triplicate LC−MS/MS measurements were performed for each fragmentation technique. The numbers of identified internal peptides, along with the canonical as well the truncated C-termini, are shown in Figure 3 for each combination of fragmentation technique (HCD, ETD, and CID), along with the protease (trypsin and chymotrypsin) and the amine used for C-terminal labeling. The increase in basicity and hence the charge state of peptides after DMEDA labeling clearly made these peptides more favorable for ETD fragmentation; consequently, more peptides were identified with ETD than CID or HCD. In contrast, EA-labeled peptides performed very poorly with ETD, most likely due to the lower number of peptides with charge states of 3+ or higher. It can be seen that for both CID and HCD, a larger number of EAG


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Journal of Proteome Research modified peptides were identified than with DMEDA. This is not surprising because CID, and in particular HCD, has been shown previously to provide more peptide identifications than ETD for doubly charged peptides on an Orbitrap Velos.31 Interestingly, amide formation using the noncharged (EA) and the charge-reversal (DMEDA) reagents and their performance, in terms of number of peptides identified, was also dependent on the work protease (chymotrypsin or trypsin). DMEDA was more favorable for tryptic peptides and outperformed EA regardless of the fragmentation technique applied. In contrast, EA yielded more unique peptide identifications when chymotrypsin was used as the work protease. This was less ambiguous when redundant peptide identifications of C-termini (in terms of being identified by ETD, CID, or HCD) were removed, resulting in 302 protein C-termini (canonical and truncated C-terminal peptides combined) (Supplementary Figure 5). For the EA-derivatized sample, 214 C-termini were identified in total, while 123 were originating from tryptic, and 91 were additionally identified from chymotryptic cleavage events. By using DMEDA, we identified 189 protein C-termini; of these were 141 from tryptic, and only 48 were contributed by chymotryptic digestion. During protein digestion, trypsin ensures that the C-terminal peptide contains a basic residue that facilitates their MS/MS-fragmentation behavior and identification. However, the C-termini of proteins processed with trypsin can contain any residue that can hamper peptide identification because they are optimized for fully tryptic peptides. DMEDA effectively introduces a charge at the peptide C-terminus that, depending on the peptide, can help it to ionize and give a more complete b- and y-ion fragmentation series facilitating its identification. This benefit was not observed for chymotrypsin. Nevertheless, it is clear that the use of multiple enzymes is advantageous for the analysis of protein C-termini and has been shown to deliver deeper proteomic coverage than when using trypsin alone.32,33 This is particular true for C- (and N-) terminomics approaches, in which the identification of a certain protein is usually based on a single peptide. The parallel use of different proteases circumvents this problem by generating more than one peptide species containing the C-terminus of the protein. This increases the confidence of identification. In this regard, the use of chymotrypsin in parallel to trypsin is a good choice. Other commonly used proteases (e.g., Glu-C or Asp-N) cannot be used due to blocking of the cleavage sites by modifications introduced in the procedure or deliver no complementary information to trypsin. In addition to utilizing the LTQ Orbitrap Velos for the identification of C-termini, we also analyzed the same E. coli samples on a more sophisticated machine: an UHPLC coupled to an Orbitrap Q Exactive Plus. With the higher LC resolving power of the UHPLC and the faster MS scan rates, we were able to identify more-or-less the same number of protein Ctermini (310) in considerably less time compared to the HPLC-Velos platform (302) utilizing HCD, CID, and ETD (Supplementary Figure 6). Finally, we merged the identified C-termini from all experiments on both the HPLC-Velos and the UHPLC-Q Exactive platforms: a comparable number of protein C-termini were identified using EA (311) and DMEDA (279) (Figure 4). The labeling strategies were very complementary, however, with a high overlap. Taken together, we were able to identify 424 C-termini from E. coli proteome using the combination of multiprotease digestion and two different carboxyl group

Figure 4. Identified protein C-termini in E. coli proteome after merging results from HPLC-Orbitrap Velos and UHPLC-Q Exactive. (A) Comparison between EA- and DMEDA-derivatized C-termini (both canonical and truncated species combined). (B) Illustration of the relations between canonical and truncated C-termini after the merging of EA and DMEDA data sets.

derivatizing reagents. In total, we identified 669 proteins including those, which were only identified with unique internal peptides (245 proteins or 37%). The relatively low number of proteins without identified C-term confirms the high efficiency of the depletion of internal peptides. From the 424 C-termini, 267 (63%) matched to genetically encoded canonical protein C-termini, while the remaining 157 (37%) C-termini are potentially truncated forms. The later could be generated as a result of proteolytic processing within the cell (Supplementary Table S3). For example, 50S ribosomal protein L31 (P0A7M9), which is encoded as a 70 amino acid consisting protein, was identified with the encoded protein Cterminus and a truncation variant with a C-terminal Lys at residue 62. This site has been previously shown to be processed by protease VII yielding a truncated protein lacking residues 63−70.34 Another example is the 30S ribosomal protein S6 (P02358), which has been isolated in five different isoforms that differ only in the number of C-terminal glutamate residues. The long form (6 Glu residues) is the canonical form; however, the short form with two Glu is encoded by the rpsF gene, and additional Glu are added post-translationally by the RimK enzyme.35 The protein with additional 2, 3, and 4 C-terminal glutamic acid residues attached was identified in our study but not the canonical form that is in the database. The large percentage of noncanonical protein C-termini that was identified in our study (∼37%) is comparable to that previously reported by Schilling and co-workers (∼40%). Here, we also identified more E. coli C-termini (424) than the previously reported 196 by Schilling et al.8 by combining two different work proteases (chymotrypsin and trypsin) and two compounds to amidate the carboxyl groups (EA and DMEDA).

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CONCLUSION C-terminal charge-reversal derivatization using DMEDA or the arginine analogue agmatine in general provides better fragmentation efficiencies compared to the use of peptides H


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different C-terminal forms) and other post-translational modifications may contribute to this phenomenon. Therefore, the parallel application of different approaches, including such a capability to analyze other protein modifications (e.g., via ETD), may become important in the future for elucidating the yet-undiscovered manifold biological functions of protein Ctermini.

derivatized with ethanolamine, which yield noncharged Ctermini. The introduction of positively charged groups at the Cterminal carboxyl group mimics the behavior of tryptic peptides and thus can improve peptide identification of C-terminal peptides lacking basic residues in their sequence. While high labeling efficiencies were achieved at peptide level, lower derivatization efficiencies on intact proteins were observed, probably due to lower accessibility of certain carboxyl groups by steric hindrance in their surrounding protein environment. The use of organic solvents or detergents improved carboxyl derivatization efficiencies only to a limited extent. Although the achievement of high derivatization yields at the intact protein level remains a yet-unsolved general problem, one can question the need for complete derivatization, at least for internal Asp- and Glu-carboxyl groups in this C-terminomics approach. Because internal peptides are depleted via a carboxyl-reactive polymer reacting with neo-Ctermini formed after proteolytic digestion, additional free Aspand Glu-residues can serve as additional sites improving depletion efficiency. Only unprotected acidic residues close to the C-terminus would thus contribute to a loss of information. However, this can be overcome to a certain extent by the formation of different peptide species using multiple proteases, as suggested in our approach. Overall, rather than the achievement of complete derivatization of all carboxyl residues, a more important feature would be an increased selectivity for the derivatization of the C-terminal α-carboxyl function. Indeed, we observed a slightly increased efficiency of derivatization of α-carboxyl groups compared over internal Asp and Glu ε-carboxyl groups, which is certainly influenced by factors as shielding of the latter by the three-dimensional protein structure. Nevertheless, finding appropriate methods for increasing the selectivity toward the C-terminal α-carboxyl function will be an important goal for future developments. The use of complementary proteases and three different fragmentation techniques improves the identification of Ctermini in complex mixtures (e.g., full proteomes), enabling the identification of peptides with different properties. This increases the confidence of identifications. Furthermore, DMEDA-derivatized peptides resulted in many peptides with charge states of 3+, which is beneficial when using ETD fragmentation. This makes the charge-reversal strategy particularly useful for the analysis of protein C-termini, which may also be post-translationally modified (e.g. phosphorylated), or for the analysis of secreted membrane proteins, in which Oglycosylation close to the transmembrane domain is known to modulate shedding. Combining the use of charge-reversal derivatization, multiprotease digestion, and three different MS/MS activation techniques, we identified more E. coli C-termini (424) than reported previously by Schilling et al. (196).8 Zhang and coworkers identified 369 high-confidence protein C-termini from an E. coli cell lysate also using a modified Schilling protocol.10 In another C-terminomics study Liu, et al. used positive enrichment of C-termini from Thermoanerobacter tengcongensis based on oxazolone chemistry and biotinylation; they identified 183 C-termini using chymotryptic digestion and CID fragmentation.7 It is obvious that in all of these studies, including ours, the number of identified C-termini is well below the number of proteins identified in classical full proteome analyses. Factors such as further C-terminal processing leading to ensembles of rugged C-termini (thereby diluting the concentration of the

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jproteome.6b00146. Figures showing a scheme of C-terminomics workflow; derivatization efficiencies analyzed for a protein mixture of β-casein, β-lactoglobulin, and BSA derivatized at the protein level with EA; comparison of peptide depletion efficiencies after derivatization of BSA at protein level; a comparison of extracted ion chromatograms and MS spectra of the C-terminal peptide LVVSTQTALA of BSA after peptide level derivatization; overlap of identified unique protein C-termini in E. coli after derivatization and indirect enrichment of C-terminal peptides; and identified protein C-termini from E. coli proteome. (PDF) A table showing derivatization efficiencies of β-casein and BSA at the peptide level. (XLSX) A table showing derivatization efficiencies of β-casein and BSA at protein level. (XLSX) A table showing identified C-terminal peptides in E. coli. (XLSX)

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AUTHOR INFORMATION

Corresponding Author

*Phone: 49-(431)-597-2335; fax: 49-(431)-597-1887; e-mail: a. [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Kai Stürmer for performing control experiments, Christian Treitz for providing the E. coli samples and Liam Cassidy for helpful discussions. This work was supported by the Cluster of Excellence “Inflammation at Interfaces” and by the SFB877, project Z2, both funded by the Deutsche Forschungsgemeinschaft (DFG).

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REFERENCES

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