Mitochondrial phosphoproteome revealed by an improved IMAC method and MS/MS/MS

IMAC in combination with mass spectrometry is a promising approach for global analysis of protein phosphorylation. Nevertheless this approach suffers from two shortcomings: inadequate efficiency of IMAC and poor fragmentation of phosphopeptides in the mass spectrometer. Here we report optimization of the IMAC procedure using (32)P-labeled tryptic peptides and development of MS/MS/MS (MS3) for identifying phosphopeptide sequences and phosphorylation sites. The improved IMAC method allowed recovery of phosphorylated tryptic peptides up to approximately 77% with only minor retention of unphosphorylated peptides. MS3 led to efficient fragmentation of the peptide backbone in phosphopeptides for sequence assignment. Proteomics of mitochondrial phosphoproteins using the resulting IMAC protocol and MS3 revealed 84 phosphorylation sites in 62 proteins, most of which have not been reported before. These results revealed diverse phosphorylation pathways involved in the regulation of mitochondrial functions. Integration of the optimized batchwise IMAC protocol with MS3 offers a relatively simple and more efficient approach for proteomics of protein phosphorylation.     

Phosphopeptide detection using automated online IMAC-capillary LC-ESI-MS/MS

IMAC has become a commonly used technique in phosphoprotein analysis because of its affinity for phosphopeptides. However, the commonly used strategy combining offline IMAC enrichment with desalting procedures prior to MS/MS makes this method laborious. Here we report the development of a robust and automatic IMAC-capillary RP HPLC-ESI MS/MS technology platform, by which all procedures needed in phosphopeptide analysis including IMAC enrichment, RP HPLC separation and nanospray MS/MS can be done automatically controlled by the MassLynx program. The platform was optimized by analyzing standard phosphopeptide, and was then applied to the identification of phosphorylation sites of recombinant human telomeric repeat binding factor 1 treated with kinase in vitro, and two phosphorylation sites are defined.     

On-chip microextraction for proteomic sample preparation of in-gel digests

Despite the high sensitivity and relatively high tolerance for contaminants of matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF MS) there is often a need to purify and concentrate the sample solution, especially after in-gel digestion of proteins separated by two-dimensional gel electrophoresis (2-DE). A silicon microextraction chip (SMEC) for sample clean-up and trace enrichment of peptides was manufactured and investigated. The microchip structure was used to trap reversed-phase chromatography media (POROS R2 beads) that facilitates sample purification/enrichment of contaminated and dilute samples prior to the MALDI-TOF MS analysis. The validity of the SMEC sample preparation technique was successfully investigated by performing analysis on a 10 nM peptide mixture containing 2 m urea in 0.1 m phosphate-buffered saline with MALDI-TOF MS. It is demonstrated that the microchip sample clean-up and enrichment of peptides can facilitate identification of proteins from 2-DE separations. The microchip structure was also used to trap beads immobilized with trypsin, thereby effectively becoming a microreactor for enzymatic digestion of proteins. This microreactor was used to generate a peptide map from a 100 nM bovine serum albumin sample.     

Preparation and application of a partially degradable gel in mass spectrometry-based proteomic analysis

In-gel digestion is an attractive route in mass spectrometry-based proteomic analysis, which, however, often suffers from a certain amount of sample loss mainly due to insufficient protein digestion and peptide extraction. To address this, herein we establish a partially degradable gel-assisted protein digestion and peptide recovery method by means of a simple replacement of bis-acrylamide (BA) with bis-acrylylcystamine (BAC). Concretely, the protein sample solubilized using high concentrations of sodium dodecyl sulfate (SDS) and urea were directly entrapped and immobilized into BAC-crosslinked gel by vacuum-dried gel absorption followed by fixation treatment. After removal of SDS and urea by repeated washing, the proteins were subjected to in-gel digestion and the gel was reductively treated. The tryptic peptides were recovered from the partial degradation of the gel and analyzed afterwards by capillary liquid chromatography coupled with tandem mass spectrometry (CapLC-MS/MS). Compared with conventional BA-crosslinked gel method, this new method increased the numbers of identified proteins and unique peptides by 20.2% and 20.4%, respectively. The further statistical analysis demonstrated that the method improved the recovery of tryptic peptides particularly larger and/or hydrophobic peptides, thereby significantly facilitating protein identification. Thus, the newly developed method is a promising alternative for BA-crosslinked gel-based shotgun workflows and has potential application in the related fields of protein chemistry and proteomics.     

Reduction of non-specific binding in Ga(III) immobilized metal affinity chromatography for phosphopeptides by using endoproteinase glu-C as the digestive enzyme

The selectivity of immobilized metal affinity chromatography (IMAC) systems for the purification of phosphopeptides is poor. This is particularly a problem with tryptic digests of proteins where a large number of acidic peptides are produced that also bind during IMAC. The hypothesis examined in this work was that the selectivity of IMAC columns for phosphopeptides could be increased by using endoproteinase glu-C (glu-C) for protein digestion. Glu-C cleaves proteins at acidic residues and should reduce the number of acidic residues in peptides. This method was successfully applied to a mixture of model proteins and bovine milk. The percentage of phosphorylated peptides selected from proteolytic digests of the milk sample was increased from 40% with trypsin to 70% with glu-C. Additionally, this method was coupled with stable isotope coding methods to quantitatively compare the concentration of phosphoproteins between samples.     

Fe3+ immobilized metal affinity chromatography with silica monolithic capillary column for phosphoproteome analysis

Immobilized metal affinity chromatography (IMAC) is a commonly used technique for phosphoproteome analysis due to its high affinity for adsorption of phosphopeptides. Miniaturization of IMAC column is essential for the analysis of a small amount of sample. Nanoscale IMAC column was prepared by chemical modification of silica monolith with iminodiacetic acid (IDA) followed by the immobilization of Fe3+ ion inside the capillary. It was demonstrated that Fe3+-IDA silica monolithic IMAC capillary column could specifically capture the phosphopeptides from tryptic digest of alpha-casein with analysis by MALDI-TOF MS. The silica monolithic IMAC capillary column was manually coupled with nanoflow RPLC/nanospray ESI mass spectrometer (muRPLC-nanoESI MS) for phosphoproteome analysis. The system was validated by analysis of standard phosphoproteins and then it was applied to the analysis of protein phosphorylation in mouse liver lysate. Besides MS/MS spectra, MS/MS/MS spectra were also collected for neutral loss peak. After database search and manual validation with conservative criteria, 29 singly phosphorylated peptides were identified by analyzing a tryptic digest of only 12 mug mouse liver lysate. The results demonstrated that the silica monolithic IMAC capillary column coupled with muRPLC-nanoESI MS was very suitable for the phosphoproteome analysis of minute sample.     

Quantitative Assessment of In-solution Digestion Efficiency Identifies Optimal Protocols for Unbiased Protein Analysis

The majority of mass spectrometry-based protein quantification studies uses peptide-centric analytical methods and thus strongly relies on efficient and unbiased protein digestion protocols for sample preparation. We present a novel objective approach to assess protein digestion efficiency using a combination of qualitative and quantitative liquid chromatography-tandem MS methods and statistical data analysis. In contrast to previous studies we employed both standard qualitative as well as data-independent quantitative workflows to systematically assess trypsin digestion efficiency and bias using mitochondrial protein fractions. We evaluated nine trypsin-based digestion protocols, based on standard in-solution or on spin filter-aided digestion, including new optimized protocols. We investigated various reagents for protein solubilization and denaturation (dodecyl sulfate, deoxycholate, urea), several trypsin digestion conditions (buffer, RapiGest, deoxycholate, urea), and two methods for removal of detergents before analysis of peptides (acid precipitation or phase separation with ethyl acetate). Our data-independent quantitative liquid chromatography-tandem MS workflow quantified over 3700 distinct peptides with 96% completeness between all protocols and replicates, with an average 40% protein sequence coverage and an average of 11 peptides identified per protein. Systematic quantitative and statistical analysis of physicochemical parameters demonstrated that deoxycholate-assisted in-solution digestion combined with phase transfer allows for efficient, unbiased generation and recovery of peptides from all protein classes, including membrane proteins. This deoxycholate-assisted protocol was also optimal for spin filter-aided digestions as compared with existing methods.MS-based proteomics is an indispensable technology for the characterization of complex biological systems, including relative or absolute protein expression levels and protein post-translational modifications. The most popular method for analyzing medium to high complexity protein samples in large-scale proteomics relies on protein digestion by using the endoprotease trypsin. Analysis and sequencing of tryptic peptides by liquid chromatography-tandem MS (LC-MS/MS)¹ then enables identification and determination of protein expression levels based on the peptide ion abundance level or the (fragment) ion intensities of identified peptides. This peptide-centric approach thus strongly relies on efficient, unbiased and reproducible protein digestion protocols. Efficiency is required to maximize the number of detectable peptides per protein (coverage) to distinguish unique proteins within protein families with similar sequences and/or sequence variants, and to detect post-translational modifications. Unbiased generation of peptides is required for the resulting data set to most accurately reflect the relative (stoichiometry) and absolute protein abundance in a sample. A particular protocol should be unbiased with respect to abundance, molecular weight, hydrophobicity and protein class. Membrane proteins for example are often suspected to be underrepresented. For MS-based proteomics approaches several critical steps can be distinguished: (a) disruption and solubilization of cells and protein complexes, (b) protein denaturation and enzymatic proteolysis, (c) MS-compatible peptide recovery, which normally entails removal of reagent leftovers and desalting before MS analysis, (d) adequate peptide separation (achieved by liquid chromatography), and (e) MS peptide analysis and sequencing (MS/MS), including the chosen data acquisition strategy.Comparative evaluations of digestion protocols generally consist of qualitative studies using standard tandem mass spectrometry. These approaches may reveal efficiency (i.e. more identifications), but are unable to reveal digestion protocol induced bias with respect to peptide and protein abundance, including membrane proteins. In addition, most data-dependent acquisition workflows are intrinsically biased, which is detrimental for making comparisons. The aim of the present study was to systematically assess efficiency and bias of trypsin-based protocols applying both standard qualitative and label-free quantitative MS approaches.The in-gel digestion protocol for proteomics, established over 15 years ago (1), has been the cornerstone method affording robust protein identifications from many sample types. Although sodium dodecyl sulfate (SDS) interferes with trypsin digestion and hampers LC-MS analysis, this powerful detergent can still be used to achieve complete protein solubilization as gel-separation is an effective way to remove interfering substances. Gel-based approaches are however not optimal for protein samples of increasing complexity and dynamic range (2). Inherent and practical limitations include, for example, concentration-dependent, incomplete peptide recovery and error-prone handling procedures (3–6). This hampers throughput, reproducibility and unbiased protein analysis, which in recent years has prompted a shift toward the application and optimization of in-solution digestion procedures.Previous comparative studies revealed that for in-solution digestions, the acid labile and MS-compatible detergent RapiGest performed most favorably compared with buffer only, urea, other detergents and organic solvents (7–9). Sodium deoxycholate (SDC), naturally found in mammalian bile (10), has emerged as a cheaper MS-compatible detergent for in-solution digestion (11). Unlike other detergents, SDC was found to enhance trypsin activity almost fivefold at a concentration of 1% (12). Like RapiGest, SDC can also be removed by acidification, but potentially without detrimental peptide loss if a phase separation protocol involving organic solvent is applied (12).An alternative strategy is to perform protein digestion on spin filter devices, introduced a few years ago by Manza and co-workers (13), and further developed by Wisniewski et al. (14). This approach allows the use of SDS to first achieve complete protein solubilization followed by removal of the detergent through repeated washes with urea (14). This is an effective way to remove interfering chemicals and small molecules after protein solubilization, and before digestion, without substantial sample loss. Although this protocol is touted to be a highly effective and universal method for any type of sample, digestion is performed using urea or buffer only and has so far not been evaluated in combination with detergents such as SDC.For our comparative study we selected protocols and methods based on spin filter-aided and standard in-solution digestion that were previously reported optimal and we also report novel optimized protocols. We investigated several experimental parameters including reagents for protein solubilization and denaturation (SDS, SDC, urea), spin filter aided removal of SDS before digestion (urea, SDC, buffer), trypsin digestion conditions (buffer, RapiGest, SDC, urea), and methods for removal of detergents before analysis of peptides (acid precipitation or phase separation with ethyl acetate).Mitochondria are organelles carrying out key metabolic processes fundamental for cellular function (15). The mitochondrial proteome is predicted to contain up to a thousand proteins (16) and is very heterogeneous with a wide range of protein pI, molecular weight and hydrophobicity values (17). We selected mitochondrial preparations to serve as model sample of medium complexity, containing a favorable combination of peptide and protein classes, including soluble and insoluble membrane-anchored or integral proteins.Using standard qualitative as well as data-independent quantitative LC-MS/MS workflows we demonstrate that SDC-based protocols combined with phase separation are the most optimal for both in-solution and filter-aided tryptic digestion, yielding the highest efficiency and lowest bias. This workflow enabled quantitative and objective assessment of various protein digestion conditions, identifying optimal protocols for efficient and unbiased protein analysis.     

Comparison of detergent‐based sample preparation workflows for LTQ‐Orbitrap analysis of the Escherichia coli proteome

This work presents a comparative evaluation of several detergent‐based sample preparation workflows for the MS‐based analysis of bacterial proteomes, performed using the model organism Escherichia coli. Initially, RapiGest‐ and SDS‐based buffers were compared for their protein extraction efficiency and quality of the MS data generated. As a result, SDS performed best in terms of total protein yields and overall number of MS identifications, mainly due to a higher efficiency in extracting high molecular weight (MW) and membrane proteins, while RapiGest led to an enrichment in periplasmic and fimbrial proteins. Then, SDS extracts underwent five different MS sample preparation workflows, including: detergent removal by spin columns followed by in‐solution digestion (SC), protein precipitation followed by in‐solution digestion in ammonium bicarbonate or urea buffer, filter‐aided sample preparation (FASP), and 1DE separation followed by in‐gel digestion. On the whole, about 1000 proteins were identified upon LC‐MS/MS analysis of all preparations (>1100 with the SC workflow), with FASP producing more identified peptides and a higher mean sequence coverage. Each protocol exhibited specific behaviors in terms of MW, hydrophobicity, and subcellular localization distribution of the identified proteins; a comparative assessment of the different outputs is presented.     

Method optimisation for peptide profiling of microdissected breast carcinoma tissue by matrix-assisted laser desorption/ionisation-time of flight and matrix-assisted laser desorption/ionisation-time of flight/time of flight-mass spectrometry

Appropriate methods for the analysis of microdissected solid tumour tissues by matrix-assisted laser desorption/ionisation-time of flight-mass spectrometry (MALDI-TOF MS) are not yet well established. Optimisation of sample preparation was performed first on undissected tissue slices, representing approximately 200 000 cells, which were solubilised either in urea containing buffer, trifluoroethanol/NH4HCO3, 0.1% sodium dodecyl sulphate (SDS) or in 0.1% RapiGest solution, then trypsin digested and analysed by MALDI-TOF MS. Solubilisation in 0.1% SDS resulted in detection of the highest number of sample specific peak signals. Interestingly, there was little overlap in detectable peaks using the different buffers, implying that they can be used complementarily to each other. Additionally, we fractionated tryptic digests on a monolithic high-performance liquid chromatography column. Fractionation of tryptic digest from whole tissue sections resulted in a four-fold increase in the total number of peaks detected. To prove this principle, we used 0.1% SDS to generate peptide patterns from 2000 microdissected tumour and stromal cells from five different breast carcinoma tumours. The tumour and stroma specific peaks could be detected upon comparison of the peptide profiles. Identification of differentially expressed peaks by MALDI-TOF/TOF MS was performed on fractionated tryptic digests derived from a whole tissue slice. In conclusion, we describe a method that is suitable for direct peptide profiling on small amounts of microdissected cells obtained from breast cancer tissues.     

Identification of membrane proteins from mammalian cell/tissue using methanol-facilitated solubilization and tryptic digestion coupled with 2D-LC-MS/MS

The core prerequisites for an efficient proteome-scale analysis of mammalian membrane proteins are effective isolation, solubilization, digestion and multidimensional liquid chromatography-tandem mass spectrometry (LC-MS/MS). This protocol is for analysis of the mammalian membrane proteome that relies on solubilization and tryptic digestion of membrane proteins in a buffer containing 60% (vol/vol) methanol. Tryptic digestion is followed by strong cation exchange (SCX) chromatography and reversed phase (RP) chromatography coupled online with MS/MS for protein identification. The use of a methanol-based buffer eliminates the need for reagents that interfere with chromatographic resolution and ionization of the peptides (e.g., detergents, chaotropes, inorganic salts). Sample losses are minimized because solubilization and digestion are carried out in a single tube avoiding any sample transfer or buffer exchange between these steps. This protocol is compatible with stable isotope labeling at the protein and peptide level, enabling identification and quantitation of integral membrane proteins. The entire procedure--beginning with isolated membrane fraction and finishing with MS data acquisition--takes 4-5 d.     

A solid phase extraction-based platform for rapid phosphoproteomic analysis

Protein phosphorylation is among the most common and intensely studied post-translational protein modification. It plays crucial roles in virtually all cellular processes and has been implicated in numerous human diseases, including cancer. Traditional biochemical and genetic methods for identifying and monitoring sites of phosphorylation are laborious and slow and in recent years have largely been replaced by mass spectrometric analysis. Improved methods for phosphopeptide enrichment coupled with faster and more sensitive mass spectrometers have led to an explosion in the size of phosphoproteomic datasets. However, wider application of these methods is limited by equipment costs and the resultant high demand for instrument time as well as by a technology gap between biologists and mass spectrometrists. Here we describe a modified two-step enrichment strategy that employs lysC digestion and step elution from self-packed strong cation exchange (SCX) solid phase extraction (SPE) columns followed by immobilized metal ion affinity chromatography (IMAC) and LC–MS/MS analysis using a hybrid LTQ Orbitrap Velos mass spectrometer. The SCX procedure does not require an HPLC system, demands little expertise, and because multiple samples can be processed in parallel, can provide a large savings of time and labor. We demonstrate this method in conjunction with stable isotope labeling to quantitate peptides harboring >8000 unique phosphorylation sites in yeast in 12 h of instrument analysis time and examine the impact of enzyme choice and instrument platform.     

Microcolumn capture and digestion of proteins combined with mass spectrometry for protein identification

A procedure has been developed for protein identification using mass spectrometry (MS) that incorporates sample cleanup, preconcentration, and protein digestion in a single-stage system. The procedure involves the adsorption of a protein, or protein mixture, from solution onto a hydrophobic resin that is contained within a microcolumn. Sample loading is accomplished by flowing the protein solution through the microcolumn, where the protein adsorbs to the hydrophobic surface. The protein is digested while still bound to the hydrophobic surface by flowing a buffered trypsin solution through the column bed. The peptide fragments are subsequently eluted for detection by MALDI or ESI-MS. The procedure is demonstrated using dilute protein samples containing high concentrations of salt, urea, and modest amount of sodium dodecyl sulfate relative to protein. Peptide fragments are also detected by MS from a 500 nM bacteriorhodopsin solution digested in a microcolumn. In this case, a combined cyanogen bromide/trypsin digestion was performed in-column. The procedure is applied to the MALDI-MS/MS identification of proteins present in an individual fraction collected by ion exchange HPLC separation of E. coli total cell extract. An additional application is illustrated in the analysis of a human plasma fraction. A total of 14 proteins, which were present in the sample at sub-micromolar concentrations, were identified from ESI-MS/MS. The microcolumn digestion procedure represents the next step toward a system for fully automated protein analysis through capture and digestion of the adsorbed protein on hydrophobic surfaces.     

Immobilized metal affinity chromatography revisited: pH/acid control toward high selectivity in phosphoproteomics

Despite recent advances in instrumentation and analytical strategies for identification and quantitation of protein phosphorylation, a highly specific enrichment protocol is still a challenge in large-scale studies. Here, we report a simple pH/acid control method that addresses the poor specificity seriously criticized in IMAC. Detailed evaluation of the capture and release mechanism in IMAC revealed that pH, buffer and salt yield a complex interplay in enrichment of phosphopeptides, yet they play individual roles in recovery and specificity. A revised one-step IMAC method with low sample loss and high specificity can be rationally designed by controlling salt, pH and the structure and concentration of organic acid. Without methyl esterification, the one-step IMAC enrichment with single LC-MS/MS identified 386 phosphoproteins in 550 mug of non-small-cell lung cancer cell lysate with 96% specificity. Additional fractionation by SDS-PAGE from 4 mg of cell lysate revealed the comprehensive proteome map, identifying 2747 phosphorylation sites from 2360 nondegenerate phosphopeptides and 1219 phosphoproteins with a false discovery rate of 0.63%. To our knowledge, this pH/acid-controlled IMAC procedure provides higher specificity than any other one-step IMAC purification procedure. Furthermore, the simple and reproducible IMAC protocol can be adapted to other solid supports, fully automated or manual, for large-scale identification of the vastly under-explored phosphoproteome.     

Enhancement of the efficiency of phosphoproteomic identification by removing phosphates after phosphopeptide enrichment

Immobilized metal affinity chromatography (IMAC) and titanium oxide (TiO2) chromatography are simple, widely used, and cost-effective methods to enrich phosphopeptides, but the sample loading buffer composition, desalting procedure, and control of loading amount are critical to avoid nonspecific interactions and to achieve efficient phosphopeptide enrichment. Although the combination of MS3 analysis and high-resolution mass spectrometry (MS) is helpful to identify phosphopeptides, the quality of many MS/MS spectra having a neutral loss peak of phosphate is still too poor to allow sequence identification, and this results in many false-negative as well as false-positive identifications. Here, we present a novel strategy, which is based on the use of alkaline phosphatase to remove phosphates and analysis of phospho/dephosphopeptide retention times to increase the reliability of identification. The use of phospho/dephosphopeptide retention time ratios allows the identification of phosphopeptides with high confidence with the aid of a focused database of dephosphopeptides. This approach was very effective to identify multiple phophorylations in tryptic peptides. A 'true' phosphorylation data set should contain about 90% phospho-Ser and a few percent phospho-Tyr, and this ratio can be used as a quality criterion for evaluation of data sets. By applying this efficient approach, we were able to identify more than one thousand phosphopeptides.     

Automated in-solution protein digestion using a commonly available high-performance liquid chromatography autosampler

A completely automated peptide mapping liquid chromatography/mass spectrometry (LC/MS) system for characterization of therapeutic proteins in which a common high-performance liquid chromatography (HPLC) autosampler is used for automated sample preparation, including protein denaturation, reduction, alkylation, and enzymatic digestion, is described. The digested protein samples are then automatically subjected to LC/MS analysis using the same HPLC system. The system was used for peptide mapping of monoclonal antibodies (mAbs), known as a challenging group of therapeutic proteins for achieving complete coverage and quantitative representation of all peptides. Detailed sample preparation protocols, using an Agilent HPLC system, are described for Lys-C digestion of mAbs with intact disulfide bonds and tryptic digestion of mAbs after reduction and alkylation. The automated procedure of Lys-C digestion of nonreduced antibody, followed by postdigestion disulfide reduction, produces both the nonreduced and reduced digests that facilitate disulfide linkage analysis. The automated peptide mapping LC/MS system has great utility in preparing and analyzing multiple samples for protein characterization, identification, and quantification of posttranslational modifications during process and formulation development as well as for protein identity and quality control.     

Global phosphoproteome of HT-29 human colon adenocarcinoma cells

Phosphorylation events in cellular signaling cascades triggered by a variety of cellular stimuli modulate protein function, leading to diverse cellular outcomes including cell division, growth, death, and differentiation. Abnormal regulation of protein phosphorylation due to mutation or overexpression of signaling proteins often results in various disease states. We provide here a list of protein phosphorylation sites identified from HT-29 human colon adenocarcinoma cell line by immobilized metal affinity chromatography (IMAC) combined with liquid chromatography (LC)-tandem mass spectrometry (MS/MS) analysis. In this study, proteins extracted from HT-29 whole cell lysates were digested with trypsin and carboxylate groups on the resulting peptides were converted to methyl esters. Derivatized phosphorylated peptides were enriched using Fe(3+)-chelated metal affinity resin. Phosphopeptides retained by IMAC were separated by high performance liquid chromatography (HPLC) and analyzed by electrospray ionization-quadrupole-time-of-flight (ESI-Q-TOF) mass spectrometry. We identified 238 phosphorylation sites, 213 of which could be conclusively localized to a single residue, from 116 proteins by searching MS/MS spectra against the human protein database using MASCOT. Peptide identification and phosphorylation site assignment were confirmed by manual inspection of the MS/MS spectra. Many of the phosphorylation sites identified in our results have not been described previously in the scientific literature. We attempted to ascribe functionality to the sites identified in this work by searching for potential kinase motifs with Scansite (http://scansite.mit.edu) and obtaining information on kinase substrate selectivity from Pattern Explorer (http://scansite.mit.edu/pe). The list of protein phosphorylation sites identified in the present experiment provides broad information on phosphorylated proteins under normal (asynchronous) cell culture conditions. Sites identified in this study may be utilized as surrogate bio-markers to assess the activity of selected kinases and signaling pathways from different cell states and exogenous stimuli.     

Cyclization of N-terminal S-carbamoylmethylcysteine causing loss of 17 Da from peptides and extra peaks in peptide maps

Enzymatic digests of proteins S-alkylated with iodoacetamide may contain peptides with N-terminal S-carbamoylmethylcysteine. These can be partly converted to a form with 17 Da lower mass and increased HPLC retention. Proof by synthesis supported by MS/MS and NMR spectroscopy was used to show that N-terminal S-carbamoylmethyl-L-cysteine can cyclize, losing NH3 to form an N-terminal residue of (R)-5-oxoperhydro-1,4-thiazine-3-carboxylic acid. The abbreviation Otc is proposed for the (R)-5-oxoperhydro-1,4-thiazine-3-carbonyl residue. The rate of cyclization is significant in 0.1 M NH4HCO3 at 37 degrees C, with the half-life of the acyclic form being 10-12 h for several peptides tested. This is similar to the rate at which N-terminal pyroglutamate forms from N-terminal glutamine.     

Improvement of an in-gel tryptic digestion method for matrix-assisted laser desorption/ionization-time of flight mass spectrometry peptide mapping by use of volatile solubilizing agents

The combination of matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS), in-gel enzymatic digestion of proteins separated by two-dimensional gel electrophoresis and searches of molecular weight in peptide-mass databases is a powerful and well established method for protein identification in proteomics analysis. For successful protein identification by MALDI-TOF mass spectrometry of peptide mixtures, critical parameters include highly specific enzymatic cleavage, high mass accuracy and sufficient numbers and sequence coverage of the peptides which can be analyzed. For in-gel digestion with trypsin, the method employed should be compatible both with enzymatic cleavage and subsequent MALDI-TOF MS analysis. We report here an improved method for preparation of peptides for MALDI-TOF MS mass fingerprinting by using volatile solubilizing agents during the in-gel digestion procedure. Our study clearly demonstrates that modification of the in-gel digestion protocols by addition of dimethyl formamide (DMF) or a mixture of DMF/N,N-dimethyl acetamide at various concentrations can significantly increase the recovery of peptides. These higher yields of peptides resulted in more effective protein identification.     

The EIPeptiDi tool: enhancing peptide discovery in ICAT-based LC MS/MS experiments

Background  

Isotope-coded affinity tags (ICAT) is a method for quantitative proteomics based on differential isotopic labeling, sample digestion and mass spectrometry (MS). The method allows the identification and relative quantification of proteins present in two samples and consists of the following phases. First, cysteine residues are either labeled using the ICAT Light or ICAT Heavy reagent (having identical chemical properties but different masses). Then, after whole sample digestion, the labeled peptides are captured selectively using the biotin tag contained in both ICAT reagents. Finally, the simplified peptide mixture is analyzed by nanoscale liquid chromatography-tandem mass spectrometry (LC-MS/MS). Nevertheless, the ICAT LC-MS/MS method still suffers from insufficient sample-to-sample reproducibility on peptide identification. In particular, the number and the type of peptides identified in different experiments can vary considerably and, thus, the statistical (comparative) analysis of sample sets is very challenging. Low information overlap at the peptide and, consequently, at the protein level, is very detrimental in situations where the number of samples to be analyzed is high.     

Phosphatase-directed phosphorylation-site determination: a synthesis of methods for the detection and identification of phosphopeptides

Phosphopeptides can be difficult to detect and sequence by mass spectrometry (MS) due to low ionization efficiency and suppression effects in the MS mode, and insufficient fragmentation in the tandem MS (MS/MS) mode, respectively. To address this problem, we have developed a technique called Phosphatase-directed Phosphorylation-site Determination (PPD), which combines on-target phosphatase reactions, MALDI MS/MS of IMAC beads on target, and hypothesis-driven MS (HD-MS). In this method, on-target dephosphorylation experiments are conducted on IMAC-bound phosphopeptides, because dephosphorylated peptides have, in general, higher MS sensitivities than the corresponding phosphopeptides. The detected dephosphorylated peptides are sequenced by MS/MS, which identifies the potentially phosphorylated peptide and the total number of Ser, Thr, or Tyr residues that could hypothetically be phosphorylated within that peptide. On the basis of this information, a mass list containing every possible phosphorylation state of each observed peptide (where 1 HPO(3) = 80 Da) is used to direct MALDI-MS/MS on the phosphorylated peptides bound to IMAC beads at each theoretical mass from the list. If the peptide is present, the resulting MS/MS spectrum reveals the exact site(s) of phosphorylation in the peptide. We have demonstrated the applicability of PPD to the detection of in vivo phosphorylation sites on the Drosophila Stem Loop Binding Protein (dSLBP), and the complementarity of this new technique to conventional MS phosphorylation site mapping methods, since the phosphorylation sites in dSLBP could not be detected by other methods.