Positional proteomics: preparation of amino-terminal peptides as a strategy for proteome simplification and characterization

We describe a protocol for selective extraction of the amino (N)-terminal-most peptide of a protein or a mixture of proteins after proteolysis. The first stage of the protocol blocks the free amino groups alpha and epsilon (the latter being lysyl residues) on the intact proteins by acetylation. In the second stage, proteolysis of the acetylated proteins yields a mixture of N-terminally acetylated (true N-terminal) and non-acetylated (internal and carboxy-terminal) peptides. Affinity capture of peptides bearing free amino groups using an immobilized amine-reactive reagent removes internal peptides from the mixture. The unbound fraction is highly enriched in N-terminal peptides, which can be analyzed without further treatment. This method is compatible with a range of proteolytic enzymes and fragmentation methods, and should take 2 d to complete. The N-terminal peptides can then be analyzed by mass spectrometry. This low cost, rapid method is readily adopted using off the shelf reagents.     

Capture of peptides with N-terminal serine and threonine: a sequence-specific chemical method for Peptide mixture simplification

The objective of this study was to evaluate a sequence-specific chemistry for the ability to specifically capture peptides that contain N-terminal serine or threonine residues from mixtures. The first step is the oxidation of the 1,2-amino alcohol structure -CH(NH(2))CH(OH)- of peptides containing N-terminal serine or threonine with periodate. The newly formed aldehyde reacts with a labeling reagent containing a hydrazide, RCONHNH(2), to form a hydrazone-peptide conjugate, RCONHN=CH-peptide. Biotin-labeled conjugates can then be isolated by affinity purification with streptavidin. The method described in this report can be useful in simplifying the complex mixtures of peptides that are generated in typical proteomic analysis, where proteins are digested with trypsin and analyzed using liquid chromatography mass spectrometry data. The sequence-specific peptide selection not only reduces the complexity of digest mixtures, but also provides additional information for peptide identification. The targeted peptides are those that have either serine or threonine adjacent to a protease cleavage site. The sequence information should greatly aid in both database matching for protein identification and for de novo sequence determination.     

Selective isolation of N-blocked peptide by combining AspN digestion, transamination, and tosylhydrazine glass treatment

Many eukaryotic proteins are blocked at the α-amino group of their N-terminal with various modifications, thereby making it difficult to determine their N-terminal sequence by protein sequencer. We propose a novel method for selectively isolating the blocked N-terminal peptide from the peptide mixture generated by endoproteinase AspN digestion of N-blocked protein. This method is based on removal of all peptides other than the N-terminal one (non-N-terminal peptides) through their carbonyl group introduced by a chemical transamination reaction. The transamination reaction converts the free α-amino group of the non-N-terminal peptides to a carbonyl group, whereas the blocked N-terminal peptide, which lacks only the free α-amino group, remains unchanged. Silica functionalized with the tosylhydrazino group effectively captures non-N-terminal peptides through their carbonyl group; thus, the blocked N-terminal peptide is selectively recovered in the supernatant. This method was applied to several model proteins, and their N-terminal peptides were successfully isolated and analyzed by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Furthermore, the method was extended to N-terminal analysis of N-free protein by artificially blocking the free α-amino group of its N-terminal with N-succinimidyloxycarbonylmethyl tris(2,4,6-trimethoxyphenyl) phosphonium bromide reagent.     

A method for N-terminal de novo sequence analysis of proteins by matrix-assisted laser desorption/ionization mass spectrometry

A novel method for isolation and de novo sequencing of N-terminal peptides from proteins is described. The method presented here combines selective chemical tagging using succinimidyloxycarbonylmethyl tris(2,4,6-trimethoxyphenyl)phosphonium bromide (TMPP-Ac-OSu) at the N^α-amino group of peptides after digestion by metalloendopeptidase (from Grifola frondosa) and selective capture procedures using p-phenylenediisothiocyanate resin, by which the N-terminal peptide can be isolated, whether or not it is N-terminally blocked. The isolated N-terminal peptide modified N-terminally with TMPP-Ac-OSu reagent produces a simple fragmentation pattern under tandem mass spectrometric analysis to significantly facilitate sequencing.     

Identification of N-myristoylated proteins by reverse-phase high performance liquid chromatography of an azlactone derivative of N-myristoylglycine.

A method is presented for the identification of N-myristoylated proteins. N-Myristoyl transferases have an absolute requirement for a free N-terminal glycine. N-Myristoylglycine is released upon mild acid hydrolysis of myristoylated peptides and proteins and its derivitization to a p-nitrobenzylazlactone with subsequent analysis by reverse phase h.p.l.c. enabled its detection to pmol levels. This facilitated the identification of N-terminal myristate in nmol quantities of purified proteins. Using this method we demonstrate that the alpha-subunit of the GTP-binding protein G0 is N-terminally myristoylated.     

Exploring proteomes and analyzing protein processing by mass spectrometric identification of sorted N-terminal peptides

Current non-gel techniques for analyzing proteomes rely heavily on mass spectrometric analysis of enzymatically digested protein mixtures. Prior to analysis, a highly complex peptide mixture is either separated on a multidimensional chromatographic system or it is first reduced in complexity by isolating sets of representative peptides. Recently, we developed a peptide isolation procedure based on diagonal electrophoresis and diagonal chromatography. We call it combined fractional diagonal chromatography (COFRADIC). In previous experiments, we used COFRADIC to identify more than 800 Escherichia coli proteins by tandem mass spectrometric (MS/MS) analysis of isolated methionine-containing peptides. Here, we describe a diagonal method to isolate N-terminal peptides. This reduces the complexity of the peptide sample, because each protein has one N terminus and is thus represented by only one peptide. In this new procedure, free amino groups in proteins are first blocked by acetylation and then digested with trypsin. After reverse-phase (RP) chromatographic fractionation of the generated peptide mixture, internal peptides are blocked using 2,4,6-trinitrobenzenesulfonic acid (TNBS); they display a strong hydrophobic shift and therefore segregate from the unaltered N-terminal peptides during a second identical separation step. N-terminal peptides can thereby be specifically collected for further liquid chromatography (LC)-MS/MS analysis. Omitting the acetylation step results in the isolation of non-lysine-containing N-terminal peptides from in vivo blocked proteins.     

Analysis of cysteine residues in peptides and proteins alkylated with volatile reagents

Summary Conditions are described for the reduction and alkylation of cysteines in peptides and proteins with volatile reagents by use of triethylphosphine as reductant, bromopropane as alkylating reagent and triethylamine as base. Alkylated samples need only be vacuum dried prior to subsequent analysis steps. Alkylated samples have been acid hydrolyzed and analyzed on an amino acid analyzer with recoveries of cysteine within 10% of the expected value. Alkylated samples have been directly applied to a sequencer membrane, dried on the surface and cysteines identified by sequence analysis without additional wash steps. In addition proteins blotted onto PVDF have been alkylatedin situ and sequenced with identification of cysteines. On the analyzer and sequencer the S-propylcysteine derivative elutes at a unique position allowing for the unambiguous identification of cysteine. Cysteine residues are quantitativly alkylated under the conditions developed. The ease of this procedure allows the routine analysis of cysteine in peptides and proteins without additional, time consuming repurification or dialysis steps.Abbreviations dptu diphenylthiourea - dmptu dimethylphenylthiourea - prop-cys S-propylcysteine     

Investigation of solubilization and digestion methods for microsomal membrane proteome analysis using data-independent LC-MSE

To evaluate the implementation of various denaturants and their efficacy in bottom-up membrane proteomic methods using LC-MS analysis, microsomes isolated from tomato roots were treated with MS-compatible surfactants (RapiGest SF Surfactant from Waters and PPS Silent Surfactant from Protein Discovery), a chaotropic reagent (guanidine hydrochloride), and an organic solvent (methanol). Peptides were analyzed in triplicate sample and technical replicates by data-independent LC-MS(E) analysis. Overall, 2333 unique peptides matching to 662 unique proteins were detected with the order of denaturant method efficacy being RapiGest SF Surfactant, PPS Silent Surfactant, guanidine hydrochloride, and methanol. Using bioinformatic analysis, 103 proteins were determined to be integral membrane proteins. When normalizing the data as a percentage of the overall number of peptides and proteins identified for each method, the order for integral membrane protein identification efficacy was methanol, guanidine hydrochloride, RapiGest SF Surfactant, and PPS Silent Surfactant. Interestingly, only 8% of the proteins were identified in all four methods with the silent surfactants having the greatest overlap at 17%. GRAVY analysis at the protein and peptide level indicated that methanol and guanidine hydrochloride promoted detection of hydrophobic proteins and peptides, respectively; however, trypsin activity in the presence of each denaturant was determined as a major factor contributing to peptide identification by LC-MS(E) . These results reveal the complementary nature of each denaturant method, which can be used in an integrated approach to provide a more effective bottom-up analysis of membrane proteomes than can be achieved using only a single denaturant.     

Amino-terminal amino acid sequences of electron transfer proteins from Gram-negative bacteria as indicators of their cellular localization: the sulfate-reducing bacteria

Abstract Data are presented that all known periplasmic redox proteins from the sulfate reducing bacteria included in the genus, Desulfovibrio have aminoterminal (N-terminal) amino-acid sequences commonly found in other Gram-negative bacteria and are indicative of recognition sites for signal peptides. In contrast, none of the cytoplasmic redox proteins exhibited these unique N-terminal amino-acid sequences. It is proposed that the N-terminal amino-acid residues of a given protein can be used as an indicator of its cellular localization within the bacterial cell.     

Proteome profiling of the green sulfur bacterium Chlorobaculum tepidum by N-terminal proteomics

In this study, we performed the first large-scale identification of N-terminal peptides from the green sulfur bacterium Chlorobaculum tepidum. Combined fractional diagonal chromatography (COFRADIC) was used to isolate protein N-terminal peptides from three different proteome preparations, and following LC-MS/MS analysis, over 621 different proteins were identified by their N-terminal peptides. Our data constitute the largest data set currently available for protein N-termini of prokaryotic photosynthetic organisms.     

In silico analysis of accurate proteomics, complemented by selective isolation of peptides

Protein identification by mass spectrometry is mainly based on MS/MS spectra and the accuracy of molecular mass determination. However, the high complexity and dynamic ranges for any species of proteomic samples, surpass the separation capacity and detection power of the most advanced multidimensional liquid chromatographs and mass spectrometers. Only a tiny portion of signals is selected for MS/MS experiments and a still considerable number of them do not provide reliable peptide identification. In this article, an in silico analysis for a novel methodology of peptides and proteins identification is described. The approach is based on mass accuracy, isoelectric point (pI), retention time (t(R)) and N-terminal amino acid determination as protein identification criteria regardless of high quality MS/MS spectra. When the methodology was combined with the selective isolation methods, the number of unique peptides and identified proteins increases. Finally, to demonstrate the feasibility of the methodology, an OFFGEL-LC-MS/MS experiment was also implemented. We compared the more reliable peptide identified with MS/MS information, and peptide identified with three experimental features (pI, t(R), molecular mass). Also, two theoretical assumptions from MS/MS identification (selective isolation of peptides and N-terminal amino acid) were analyzed. Our results show that using the information provided by these features and selective isolation methods we could found the 93% of the high confidence protein identified by MS/MS with false-positive rate lower than 5%.     

HysTag--a novel proteomic quantification tool applied to differential display analysis of membrane proteins from distinct areas of mouse brain

A novel isotopically labeled cysteine-tagging and complexity-reducing reagent, called HysTag, has been synthesized and used for quantitative proteomics of proteins from enriched plasma membrane preparations from mouse fore- and hindbrain. The reagent is a 10-mer derivatized peptide, H(2)N-(His)(6)-Ala-Arg-Ala-Cys(2-thiopyridyl disulfide)-CO(2)H, which consists of four functional elements: i) an affinity ligand (His(6)-tag), ii) a tryptic cleavage site (-Arg-Ala-), iii) Ala-9 residue that contains four (d(4)) or no (d(0)) deuterium atoms, and iv) a thiol-reactive group (2-thiopyridyl disulfide). For differential analysis cysteine residues in the compared samples are modified using either (d(4)) or (d(0)) reagent. The HysTag peptide is preserved in Lys-C digestion of proteins and allows charge-based selection of cysteine-containing peptides, whereas subsequent tryptic digestion reduces the labeling group to a di-peptide, which does not hinder effective fragmentation. Furthermore, we found that tagged peptides containing Ala-d(4) co-elute with their d(0)-labeled counterparts. To demonstrate effectiveness of the reagent, a differential analysis of mouse forebrain versus hindbrain plasma membranes was performed. Enriched plasma membrane fractions were partially denatured, reduced, and reacted with the reagent. Digestion with endoproteinase Lys-C was carried out on nonsolubilized membranes. The membranes were sedimented by ultra centrifugation, and the tagged peptides were isolated by Ni(2+) affinity or cation-exchange chromatography. Finally, the tagged peptides were cleaved with trypsin to release the histidine tag (residues 1-8 of the reagent) followed by liquid chromatography tandem mass spectroscopy for relative protein quantification and identification. A total of 355 unique proteins were identified, among which 281 could be quantified. Among a large majority of proteins with ratios close to one, a few proteins with significant quantitative changes were retrieved. The HysTag offers advantages compared with the isotope-coded affinity tag reagent, because the HysTag reagent is easy to synthesize, economical due to use of deuterium instead of (13)C isotope label, and allows robust purification and flexibility through the affinity tag, which can be extended to different peptide functionalities.     

A fast and reproducible method for albumin isolation and depletion from serum and cerebrospinal fluid

Analysis of serum and plasma proteomes is a common approach for biomarker discovery, and the removal of high‐abundant proteins, such as albumin and immunoglobins, is usually the first step in the analysis. However, albumin binds peptides and proteins, which raises concerns as to how the removal of albumin could impact the outcome of the biomarker study while ignoring the possibility that this could be a biomarker subproteome itself. The first goal of this study was to test a new commercially available affinity capture reagent from Protea Biosciences and to compare the efficiency and reproducibility to four other commercially available albumin depletion methods. The second goal of this study was to determine if there is a highly efficient albumin depletion/isolation system that minimizes sample handling and would be suitable for large numbers of samples. Two of the methods tested (Sigma and ProteaPrep) showed an albumin depletion efficiency of 97% or greater for both serum and cerebrospinal fluid (CSF). Isolated serum and CSF albuminomes from ProteaPrep spin columns were analyzed directly by LC‐MS/MS, identifying 128 serum (45 not previously reported) and 94 CSF albuminome proteins (17 unique to the CSF albuminome). Serum albuminome was also isolated using Vivapure anti‐HSA columns for comparison, identifying 105 proteins, 81 of which overlapped with the ProteaPrep method.     

An isotope coding strategy for proteomics involving both amine and carboxyl group labeling

This paper describes a heavy isotope coding strategy for the analysis of all types of tryptic peptides, including those that are N-terminally blocked and from the C-terminus of proteins. The method exploits differential derivatization of amine and carboxyl groups generated during proteolysis as a means of coding. Carboxyl groups produced during proteolysis incorporate 18O from H218O. Peptides from the C-terminus of proteins were not labeled with 18O unless they contained a basic C-terminal amino acid. Primary amines from control and experimental samples were differentially acylated after proteolysis with either 1H3- or 2H3-N-acetoxysuccinamide. When these two types of labeling were combined, unique coding patterns were achieved for peptides arising from the C-termini and blocked N-termini of proteins. This method was used to (1) distinguish C-terminal peptides in model proteins, (2) recognize N-terminal peptides from proteins in which the amino terminus is acylated, and (3) identify primary structure variations between proteins from different sources.     

Characterization of low abundant membrane proteins using the protein sequence tag technology

About 25% of open reading frames in fully sequenced genomes are estimated to encode transmembrane proteins that represent valuable targets for drugs. However, the global analysis of membrane proteins has been proven to be problematic, e.g., because of their very amphiphilic nature. In this paper, we show that the recently published Protein Sequence Tag (PST) technology combined with an efficient sample preparation is a powerful method to perform protein analysis of highly enriched membrane fractions. The PST approach is a gel-free proteomics tool for the analysis of proteins, which relies on a "sampling" strategy by isolating N-terminal protein sequence tags from cyanogen bromide cleaved proteins. The identification of these N-terminal PST peptides is based on LC-MS/MS. The effectiveness of the technology is demonstrated for a membrane fraction, which was isolated from crude mitochondria of yeast after alkaline sodium carbonate treatment. The PST approach performed on this fraction analyzed 148 proteins, whereas 84% are identified as membrane proteins. More interestingly, among these membrane proteins 56% are predicted to be of low abundance. These encouraging results are an important step toward the development of a quantitative PST approach (qPST) for the differential display of membrane protein analysis.     

Protease profiling using a fluorescent domino peptide cocktail

Five hexapeptides were prepared containing, in a domino-type arrangement, all 25 possible dipeptides between (1) aromatic, (2) hydrophobic, (3) positively charged, (4) negatively charged, and (5) small and polar amino acids. The peptides were fluorescence labeled at the N-terminus with a (7-coumaryl)oxyacetyl group, allowing the selective detection of N-terminal cleavage products. The five peptides were used as a cocktail reagent in an HPLC analysis. The cocktail produced specific cleavage patterns, or fingerprints, for a variety of proteases. This domino peptide cocktail can be used as a general reagent for protease identification and functional profiling.     

Comprehensive peptidome analysis of mouse livers by size exclusion chromatography prefractionation and nanoLC-MS/MS identification

Peptidome analysis has received increasing attention in recent years. Cancer diagnosis by serum peptidome has also been reported by peptides' profiling for discovery of peptide biomarkers. Tissue, which may have a higher biomarker concentration than blood, has not been investigated extensively by means of peptidome analysis. Here, a method for the peptidome analysis of mouse liver was developed by the combination of size exclusion chromatography (SEC) prefractionation with nano-liquid chromatography-tamdem mass spectrometry (nanoLC-MS/MS) analysis. The extracted peptides from mouse liver were separated according to their molecular weight using a size exclusion column. MALDI-TOF MS was used to characterize the molecular weight distribution of the peptides in fractions eluted from the SEC column. The low molecular weight (LMW) (MW < 3000 Da) peptides in the collected fractions were directly analyzed by LC-MS/MS which resulted in the identification of 1181 unique peptides (from 371 proteins). The high molecular weight (HMW) (MW > 3000 Da) peptides in the early two fractions from the SEC column were first digested with trypsin, and the resulted digests were then analyzed by LC-MS/MS, which led to the identification of 123 and 127 progenitor proteins of the HMW peptides in fractions 1 and 2, respectively. Analysis of the peptides' cleavage sites showed that the peptides are cleaved in regulation, which may reflect the protease activity and distribution in body, and also represent the biological state of the tissue and provide a fresh source for biomarker discovery.     

Selective enrichment of sialic acid-containing glycopeptides using titanium dioxide chromatography with analysis by HILIC and mass spectrometry

The terminal monosaccharide of cell surface glycoconjugates is typically a sialic acid (SA), and aberrant sialylation is involved in several diseases. Several methodological approaches in sample preparation and subsequent analysis using mass spectrometry (MS) have enabled the identification of glycosylation sites and the characterization of glycan structures. In this paper, we describe a protocol for the selective enrichment of SA-containing glycopeptides using a combination of titanium dioxide (TiO(2)) and hydrophilic interaction liquid chromatography (HILIC). The selectivity of TiO(2) toward SA-containing glycopeptides is achieved by using a low-pH buffer that contains a substituted acid such as glycolic acid to improve the binding efficiency and selectivity of SA-containing glycopeptides to the TiO(2) resin. By combining TiO(2) enrichment of sialylated glycopeptides with HILIC separation of deglycosylated peptides, a more comprehensive analysis of formerly sialylated glycopeptides by MS can be achieved. Here we illustrate the efficiency of the method by the identification of 1,632 unique formerly sialylated glycopeptides from 817 sialylated glycoproteins. The TiO(2)/HILIC protocol requires 2 d and the entire procedure from protein isolation can be performed in <5 d, depending on the time taken to analyze data.