Use of deuterium-labeled lysine for efficient protein identification and peptide de novo sequencing

Here, we describe a method for protein identification and de novo peptide sequencing. Through in vivo cell culturing, the deuterium-labeled lysine residue (Lys-d4) introduces a 4-Da mass tag at the carboxyl terminus of proteolytic peptides when cleaved by certain proteases. The 4-Da mass difference between the unlabeled and the deuterated lysine assigns a mass signature to all lysine-containing peptides in any pool of proteolytic peptides for protein identification directly through peptide mass mapping. Furthermore, it was used to distinguish between N- and C-terminal fragments for accurate assignments of daughter ions in tandem MS/MS spectra for sequence assignment. This technique simplifies the labeling scheme and the interpretation of the MS/MS spectra by assigning different series of fragment ions correctly and easily and is very useful in de novo peptide sequencing. We have also successfully implemented this approach to the analysis of protein mixtures derived from the human proteome.     

Non-gel-based dual 18O labeling quantitative proteomics strategy

To improve the quantitation of target proteins in proteomic analyses, we developed a non-gel-based, dual (18)O labeling strategy. This global isotope labeling method utilizes an acylating chemical reagent with two anhydride functional groups, bicyclic anhydride diethylenetriamine-N,N,N', N' ',N' '-pentaacetic acid (DTPA) dianhydride. In the first (18)O labeling method (chemical (18)O labeling) of our dual strategy, one functional group was covalently coupled to the primary amines of the peptides and (18)O from H2(18)O was incorporated at the other functional group by hydrolysis. In the second (18)O labeling method (chemical and enzyme-catalyzed (18)O labeling), chemical (18)O labeling and enzyme-catalyzed (18)O labeling of the carboxyl- termini of the peptides were combined. The acylation reaction between DTPA and the model peptides was rapid and specific, and the DTPA-modified N-termini of the peptides promoted only y-series ions in MS/MS. The two methods of (18)O labeling were accurate in the range 0.1-10 of (16)O/(18)O peptide ratios. The deviations of the methods were <20%. In contrast to current proteolytic (18)O labeling methods, there was no (18)O to (16)O back-exchange in the first method and no isotope peaks in MS in the second method. The combination of chemical and proteolytic (18)O labeling improved the confidence of the quantitation results.     

Software for quantitative proteomic analysis using stable isotope labeling and data independent acquisition

Many software tools have been developed for analyzing stable isotope labeling (SIL)-based quantitative proteomic data using data dependent acquisition (DDA). However, programs for analyzing SIL-based quantitative proteomics data obtained with data independent acquisition (DIA) have yet to be reported. Here, we demonstrated the development of a new software for analyzing SIL data using the DIA method. Performance of the DIA on SYNAPT G2MS was evaluated using SIL-labeled complex proteome mixtures with known heavy/light ratios (H/L = 1:1, 1:5, and 1:10) and compared with the DDA on linear ion trap (LTQ)-Orbitrap MS. The DIA displays relatively high quantitation accuracy for peptides cross all intensity regions, while the DDA shows an intensity dependent distribution of H/L ratios. For the three proteome mixtures, the number of detected SIL-peptide pairs and dynamic range of protein intensities using DIA drop stepwise, whereas no significant changes in these aspects using DDA were observed. The new software was applied to investigate the proteome difference between mouse embryonic fibroblasts (MEFs) and MEF-derived induced pluripotent stem cells (iPSCs) using (16)O/(18)O labeling. Our study expanded the capacities of our UNiquant software pipeline and provided valuable insight into the performance of the two cutting-edge MS platforms for SIL-based quantitative proteomic analysis today.     

Trypsin immobilization on hairy polymer chains hybrid magnetic nanoparticles for ultra fast, highly efficient proteome digestion, facile 18O labeling and absolute protein quantification

In recent years, quantitative proteomic research attracts great attention because of the urgent needs in biological and clinical research, such as biomarker discovery and verification. Currently, mass spectrometry (MS) based bottom up strategy has become the method of choice for proteomic quantification. In this strategy, the amount of proteins is determined by quantifying the corresponding proteolytic peptides of the proteins, therefore highly efficient and complete protein digestion is crucial for achieving accurate quantification results. However, the digestion efficiency and completeness obtained using conventional free protease digestion is not satisfactory for highly complex proteomic samples. In this work, we developed a new type of immobilized trypsin using hairy noncross-linked polymer chains hybrid magnetic nanoparticle as the matrix aiming at ultra fast, highly efficient proteomic digestion and facile (18)O labeling for absolution protein quantification. The hybrid nanoparticle is synthesized by in situ growth of hairy polymer chains from the magnetic nanoparticle surface using surface initiated atom transfer radical polymerization technique. The flexible noncross-linked polymer chains not only provide large amount of binding sites but also work as scaffolds to support three-dimensional trypsin immobilization which leads to increased loading amount and improved accessibility of the immobilized trypsin. For complex proteomic samples, obviously increased digestion efficiency and completeness was demonstrated by 27.2% and 40.8% increase in the number of identified proteins and peptides as well as remarkably reduced undigested proteins residues compared with that obtained using conventional free trypsin digestion. The successful application in absolute protein quantification of enolase from Thermoanaerobacter tengcongensis protein extracts using (18)O labeling and MRM strategy further demonstrated the potential of this hybrid nanoparticle immobilized trypsin for high throughput proteome quantification.     

Isobaric peptide termini labeling utilizing site-specific N-terminal succinylation

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

Identification and C-terminal characterization of proteins from two-dimensional polyacrylamide gels by a combination of isotopic labeling and nanoelectrospray Fourier transform ion cyclotron resonance mass spectrometry

We propose a novel method for the identification and C-terminal characterization of proteins separated by two-dimensional polyacrylamide gel electrophoresis (2D-PAGE). Proteins were digested in a gel in a buffer solution containing 50% 18O-labeled water, and mixtures of 18O/16O-labeled peptides were analyzed by nanoelectrospray Fourier transform ion cyclotron resonance mass spectrometry (FT-ICR MS). This method was evaluated using horse skeletal muscle myoglobin as the model protein in SDS gel. The high resolution of FT-ICR MS minimized the overlapping of peptide peaks and facilitated identification of the C-terminal peptide, which was done by observing the undisrupted isotope peak pattern. As well, with its low ppm-level high mass accuracy, it can rapidly and reliably identify the in-gel-separated protein and determine its C-terminal by peptide mass fingerprinting alone. Therefore, this method should be applicable to routine and high-throughput proteome studies. Here, the method was applied to the analysis of rat liver proteins separated by 2D-PAGE. The C-termini of eight proteins were successfully identified out of 10 randomly picked Coomassie brilliant blue-stained spots. The feasibility and limitations of this approach are reported in this paper.     

Single peptide-based protein identification in human proteome through MALDI-TOF MS coupled with amino acids coded mass tagging

Identification of proteins with low sequence coverage using mass spectrometry (MS) requires tandem MS/MS peptide sequencing. It is very challenging to obtain a complete or to interpret an incomplete tandem MS/MS spectrum from fragmentation of a weak peptide ion signal for sequence assignment. Here, we have developed an effective and high-throughput MALDI-TOF-based method for the identification of membrane and other low-abundance proteins with a simple, one-dimensional separation step. In this approach, several stable isotope-labeled amino acid precursors were selected to mass-tag, in parallel, the human proteome of human skin fibroblast cells in a residue-specific manner during in vivo cell culturing. These labeled residues can be recognized by their characteristic isotope patterns in MALDI-TOF MS spectra. The isotope pattern of particular peptides induced by the different labeled precursors provides information about their amino acid compositions. The specificity of peptide signals in a peptide mass mapping is thus greatly enhanced, resolving a high degree of mass degeneracy of proteolytic peptides derived from the complex human proteome. Further, false positive matches in database searching can be eliminated. More importantly, proteins can be accurately identified through a single peptide with its m/z value and partial amino acid composition. With the increased solubility of hydrophobic proteins in SDS, we have demonstrated that our approach is effective for the identification of membrane and low-abundant proteins with low sequence coverage and weak signal intensity, which are often difficult for obtaining informative fragment patterns in tandem MS/MS peptide sequencing analysis.     

Combinatorial peptide ligand library treatment followed by a dual-enzyme, dual-activation approach on a nanoflow liquid chromatography/orbitrap/electron transfer dissociation system for comprehensive analysis of swine plasma proteome

The plasma proteome holds enormous clinical potential, yet an in-depth analysis of the plasma proteome remains a daunting challenge due to its high complexity and the extremely wide dynamic range in protein concentrations. Furthermore, existing antibody-based approaches for depleting high-abundance proteins are not adaptable to the analysis of the animal plasma proteome, which is often essential for experimental pathology/pharmacology. Here we describe a highly comprehensive method for the investigation of the animal plasma proteome which employs an optimized combinatorial peptide ligand library (CPLL) treatment to reduce the protein concentration dynamic range and a dual-enzyme, dual-activation strategy to achieve high proteomic coverage. The CPLL treatment enriched the lower abundance proteins by >100-fold when the samples were loaded in moderately denaturing conditions with multiple loading-washing cycles. The native and the CPLL-treated plasma were digested in parallel by two enzymes (trypsin and GluC) carrying orthogonal specificities. By performing this differential proteolysis, the proteome coverage is improved where peptides produced by only one enzyme are poorly detectable. Digests were fractionated with high-resolution strong cation exchange chromatography and then resolved on a long, heated nano liquid chromatography column. MS analysis was performed on a linear triple quadrupole/orbitrap with two complementary activation methods (collisionally induced dissociation (CID) and electron transfer dissociation). We applied this optimized strategy to investigate the plasma proteome from swine, a prominent animal model for cardiovascular diseases (CVDs). This large-scale analysis results in identification of a total of 3421 unique proteins, spanning a concentration range of 9-10 orders of magnitude. The proteins were identified under a set of commonly accepted criteria, including a precursor mass error of <15 ppm, Xcorr cutoffs, and ≥2 unique peptides at a peptide probability of ≥95% and a protein probability of ≥99%, and the peptide false-positive rate of the data set was 1.8% as estimated by searching the reversed database. CPLL treatment resulted in 55% more identified proteins over those from native plasma; moreover, compared with using only trypsin and CID, the dual-enzyme/activation approach enabled the identification of 2.6-fold more proteins and substantially higher sequence coverage for most individual proteins. Further analysis revealed 657 proteins as significantly associated with CVDs (p < 0.05), which constitute five CVD-related pathways. This study represents the first in-depth investigation of a nonhuman plasma proteome, and the strategy developed here is adaptable to the comprehensive analysis of other highly complex proteomes.     

Identification and quantitative studies of protein immobilization sites by stable isotope labeling and mass spectrometry

A method was developed for characterizing immobilization sites on a protein based on stable isotope labeling and MALDI-TOF mass spectrometry. The model for this work was human serum albumin (HSA) immobilized onto silica by the Schiff base method. The immobilized HSA was digested by various proteolytic enzymes in the presence of normal water, while soluble HSA was digested in (18)O-enriched water for use as an internal standard. These two digests were mixed and analyzed, with the (18)O/(16)O ratio for each detected peptide then being measured. Several peptides in the tryptic, Lys-C, and Glu-C digests gave significantly higher (18)O/(16)O ratios than other peptides in the same digests, implying their involvement in immobilization. Analysis of these results led to identification of the N-terminus and several lysines as likely immobilization sites for HSA (e.g., K4, K41, K190, K225, K313, and K317). It was also possible from these results to quantitatively rank these sites in terms of the relative degree to which each might take part in immobilization. This method is not limited to HSA and silica but can be used with other proteins and supports.     

Identification of cross-linked peptides for protein interaction studies using mass spectrometry and 18O labeling

A new method is presented to screen proteolytic mass maps of cross-linked protein complexes for the presence of cross-linked peptides and for the verification of proposed structures. On the basis of the incorporation of 18O from isotopically enriched water into the C-termini of proteolytic peptides, cross-linked peptides are readily distinguished in mass spectra by a characteristic 8 amu shift. This is due to the incorporation of two 18O atoms in each C-terminus, so that normal and surface-labeled peptides shift 4 amu and cross-linked peptides containing two C-termini will shift 8 amu compared with their unlabeled counterparts. The method is fast, sensitive, and reliable and can be combined with any available cross-linking reagent and a wide range of proteolytic agents. As proof of principle, we successfully applied the method to a complex of two DNA repair proteins (Rad18-Rad6) and identified the interaction domain.     

Inverse 18O labeling mass spectrometry for the rapid identification of marker/target proteins

Systematic analysis of proteins is essential in understanding human diseases and their clinical treatments. To achieve the rapid and unambiguous identification of marker or target proteins, a new procedure termed "inverse labeling" is proposed. With this procedure, to evaluate protein expression of a diseased or a drug-treated sample in comparison with a control sample, two converse labeling experiments are performed in parallel. The perturbed sample (by disease or by drug treatment) is labeled in one experiment, whereas the control is labeled in the second experiment. When mixed and analyzed with its unlabeled counterpart for differential comparison using mass spectrometry, a characteristic inverse labeling pattern of mass shift will be observed between the two parallel analyses for proteins that are differentially expressed. In this study, protein labeling is achieved through 18O incorporation into peptides by proteolysis performed in [18O]water. Once the peptides are identified with the characteristic inverse labeling pattern of 18O/16O ion intensity shift, MS data of peptide fingerprints or peptide sequence information can be used to search a protein database for protein identification. The methodology has been applied successfully to two model systems in this study. It permits quick focus on the signals of differentially expressed proteins. It eliminates the detection ambiguities caused by the dynamic range of detection on proteins of extreme changes in expression. It enables the detection of protein modifications responding to perturbation. This strategy can also be extended to other protein-labeling methods, such as chemical or metabolic labeling, to realize the same benefits.     

Mapping of protein disulfide bonds using negative ion fragmentation with a broadband precursor selection

Fast mapping of disulfide bonds in proteins containing multiple cysteine residues is often required in order to assess the integrity of the tertiary structure of proteins prone to degradation and misfolding or to detect distinct intermediate states generated in the course of oxidative folding. A new method of rapid detection and identification of disulfide-linked peptides in complex proteolytic mixtures utilizes the tendency of collision-activated peptide ions to lose preferentially side chains of select amino acids in the negative ion mode. Cleavages of cysteine side chains result in efficient dissociation of disulfide bonds and produce characteristic signatures in the fragment ion mass spectra. While cleavages of other side chains result in insignificant loss of mass and full retention of the peptide ion charge, dissociation of external disulfide bonds results in physical separation of two peptides and, therefore, significant changes of both mass and charge of fragment ions relative to the precursor ion. This feature allows the fragment ions generated by dissociation of external disulfide bonds to be easily detected and identified even if multiple precursor ions are activated simultaneously. Such broadband selection of precursor ions for consecutive activation is achieved by lowering the dc/rf amplitude ratio in the first quadrupole filter of a hybrid quadrupole time-of-flight mass spectrometer. The feasibility of the new method is demonstrated by partial mapping of disulfide bridges within a 37-kDa protein containing 16 cysteine residues and complete disulfide mapping within a lysozyme (14.5 kDa) containing 8 cysteine residues. In addition to detecting peptide pairs connected by a single external disulfide, the new method is also shown to be capable of identifying peptides containing both external and internal disulfide bonds. The two major factors determining the efficiency of disulfide mapping using the new methodology are the effectiveness of proteolysis and the ability of the resulting proteolytic fragments to form multiply charged negative ions.     

Determination of fractional synthesis rates of mouse hepatic proteins via metabolic 13C-labeling, MALDI-TOF MS and analysis of relative isotopologue abundances using average masses

Proteins of a liver extract taken from a metabolically (13)C-labeled mouse were separated by 2D-PAGE and identified after tryptic digestion by MALDI-TOF MS peptide mass fingerprinting. (13)C-Labeling of proteins was achieved by an infusion of U-(13)C-glucose, which is metabolized to labeled nonessential amino acids. The labeling was analyzed using the relative isotopologue abundances of the measured isotope pattern of tryptic peptides and quantified by their increase in the average molecular mass (DeltaAVM). Fractional synthesis rates (FSR) of proteins were determined from corresponding peptides using measured DeltaAVM values as well as DeltaAVM values deduced from tRNA-precursor amino acid labeling, which in turn was derived from proteins showing high (13)C enrichments. The 8-h FSR values of 43 proteins were determined to range from 0 +/- 0.6 to 95 +/- 1%/8 h, with typical errors given as SEM values, which depend on the number of peptides of a specific protein usable for calculation. The method demonstrates that FSR values as an indicator for protein turnover in the liver proteome can be estimated within narrow error margins, providing baseline values from which treatment-dependent deviations could be detected with high statistical certainty.     

MALDI analysis of Bacilli in spore mixtures by applying a quadrupole ion trap time-of-flight tandem mass spectrometer

A novel ion trap time-of-flight hybrid mass spectrometer (qIT-TOF MS) has been applied for peptide sequencing in proteolytic digests generated from spore mixtures of Bacilli. The method of on-probe solubilization and in situ proteolytic digestion of small, acid-soluble spore proteins has been recently developed in our laboratory, and microorganism identification in less than 20 min was accomplished. In this study, tryptic peptides were generated in situ from complex spore mixtures of B. subtilis 168, B. globigii, B. thuringiensis subs. Kurstaki, and B. cereus T, respectively. MALDI analysis of bacterial peptides generated was performed with an average mass resolving power of 6200 and a mass accuracy of up to 10 ppm using a trap-TOF tandem configuration. Precursor ions of interest were usually selected and stored in the quadrupole ion trap with their complete isotope distribution by choosing a window of +/- 2 Da. Sequence-specific information on isolated protonated peptides was gained via tandem MS experiments with an average mass resolving power of 4450 for product ion analysis, and protein and bacterial sources were identified by database searching.     

Site-specific mass tagging with stable isotopes in proteins for accurate and efficient protein identification

Proteolytic peptide mass mapping as measured by mass spectrometry provides a major approach for the identification of proteins. A protein is usually identified by the best match between the measured and calculated m/z values of the proteolytic peptides. A unique identification is, however, heavily dependent upon the mass accuracy and sequence coverage of the fragment ions generated by peptide ionization. Without ultrahigh instrumental accuracy, it is possible to increase the specificity of the assignments of particular proteolytic peptides by the incorporation of selected amino acid residue(s) enriched with stable isotope(s) into the protein sequence. Here we report this novel method of generating residue-specific mass-tagged proteolytic peptides for accurate and efficient protein identification. Selected amino acids are labeled with 13C/15N/2H and incorporated into proteins in a sequence-specific manner during cell culturing. Each of these labeled amino acids carries a defined mass change encoded in its monoisotopic distribution pattern. Through their characteristic patterns, the peptides with mass tags can then be readily distinguished from other peptides in mass spectra. This method of identifying unique proteins can also be extended to protein complexes and will significantly increase data search specificity, efficiency, and accuracy for protein identifications.     

N-terminal isotope tagging strategy for quantitative proteomics: results-driven analysis of protein abundance changes

Comparing the relative abundance of each protein present in two or more complex samples can be accomplished using isotope-coded tags incorporated at the peptide level. Here we describe a chemical labeling strategy for the incorporation of a single isotope label per peptide, which is completely sequence-independent so that it potentially labels every peptide from a protein including those containing posttranslational modifications. It is based on a gentle chemical labeling strategy that specifically labels the N-terminus of all peptides in a digested sample with either a d5- or d0-propionyl group. Lysine side chains are blocked by guanidination prior to N-terminal labeling to prevent the incorporation of multiple labels. In this paper, we describe the optimization of this N-terminal isotopic tagging strategy and validate its use for peptide-based protein abundance measurements with a 10-protein standard mixture. Using a results-driven strategy, which targets proteins for identification based on MALDI TOF-MS analysis of isotopically labeled peptide pairs, we also show that this labeling strategy can detect a small number of differentially expressed proteins in a mixture as complex as a yeast cell lysate. Only peptides that show a difference in relative abundance are targeted for identification by tandem MS. Despite the fact that many peptides are quantitated, only those few showing a difference in abundance are targeted for protein identification. Proteins are identified by either targeted LC-ES MS/MS or MALDI TOF/TOF. Identifications can be accomplished equally well by either technique on the basis of multiple peptides. This increases the confidence level for both identification and quantitation. The merits of ES MS/MS or MALDI MS/MS for protein identification in a results-driven strategy are discussed.     

Phosphoprotein isotope-coded solid-phase tag approach for enrichment and quantitative analysis of phosphopeptides from complex mixtures

Many cellular processes are regulated by reversible protein phosphorylation, and the ability to broadly identify and quantify phosphoproteins from proteomes would provide a basis for gaining a better understanding of these dynamic cellular processes. However, such a sensitive, efficient, and global method capable of addressing the phosphoproteome has yet to be developed. Here we describe an improved stable-isotope labeling method using a phosphoprotein isotope-coded solid-phase tag (PhIST) for isolating and measuring the relative abundances of phosphorylated peptides from complex peptide mixtures resulting from the enzymatic digestion of extracted proteins. The PhIST approach is an extension of the previously reported phosphoprotein isotope-coded affinity tag (PhIAT) approach developed by our laboratory, where phosphoseryl and phosphothreonyl residues were derivatized by hydroxide ion-mediated beta-elimination followed by the Michael addition of 1,2-ethanedithiol (EDT). Instead of using the biotin affinity tag, peptides containing the EDT moiety were captured and labeled in one step using isotope-coded solid-phase reagents containing either light (12C6, 14N) or heavy (13C6, 15N) stable isotopes. The captured peptides labeled with the isotope-coded tags were released from the solid-phase support by UV photocleavage and analyzed by capillary liquid chromatography-tandem mass spectrometry. The efficiency and sensitivity of the PhIST labeling approach for identification of phosphopeptides from mixtures were determined using casein proteins. Its utility for proteomic applications was demonstrated by the labeling of soluble phosphoproteins from a human breast cancer cell line.     

Peptide and protein quantitation by acid-catalyzed 18O-labeling of carboxyl groups

We have developed a new method that applies acidic catalysis with hydrochloric acid for (18)O-labeling of peptides at their carboxyl groups. With this method, peptides get labeled at their C-terminus, at Asp and Glu residues, and at carboxymethylated cysteine residues. Oxygen atoms at phosphate groups of phosphopeptide are not exchanged. Our elaborated labeling protocol is easy to perform, fast (5 h and 30 min), and results in 95-97 atom % incorporation of (18)O at carboxyl groups. Undesired side reactions, such as deamidation or peptide hydrolysis, occur only at a very low level under the conditions applied. In addition, data analysis can be performed automatically using common software tools, such as Mascot Distiller. We have demonstrated the capability of this method for the quantitation of peptides as well as for phosphopeptides.     

A proteome strategy for fractionating proteins and peptides using continuous free-flow electrophoresis coupled off-line to reversed-phase high-performance liquid chromatography

Extensive prefractionation is now considered to be a necessary prerequisite for the comprehensive analysis of complex proteomes where the dynamic range of protein abundances can vary from approximately 10(6) for cells to approximately 10(10) for tissues such as blood. Here, we describe a high-resolution 2D protein separation system that uses a continuous free-flow electrophoresis (FFE) device to fractionate complex protein mixtures by solution-phase isoelectric focusing (IEF) into 96 well-defined pools, each separated by approximately 0.02-0.10 pH unit depending on the gradient created, followed by rapid (approximately 6 min per analysis) reversed-phase high-performance liquid chromatography (RP-HPLC) of each FFE pool. Fractionated proteins are readily visualized in a virtual 2D format using software that plots protein loci, pI in the first dimension and relative hydrophobicity (i.e., RP-HPLC retention time) in the second dimension. By coupling a diode-array detector in line with a multiwavelength fluorescence detector, separated proteins can be monitored in the RP-HPLC eluent by both UV absorbance and intrinsic fluorescence simultaneously from a single experiment. Triplicate analyses of standard proteins using a pH 3-10 gradient conducted over a 3-day period revealed a high system reproducibility with a SD of 0.57 (0.05 pH unit) within the FFE pools and 0.003 (0.18 s) for protein retention times in the second-dimension RP-HPLC step. In addition, we demonstrate that the FFE-IEF/RP-HPLC separation strategy can also be applied to complex mixtures of low molecular weight compounds such as peptides. With the facile ability to measure the pH of the isoelectric focused pools, peptide pI values can be estimated and used to qualify peptide identifications made using either MS/MS sequencing approaches or pI discriminated peptide mass fingerprinting. The calculated peak capacity of this 2D liquid-based FFE-IEF/RP-HPLC system is 6720.     

Minimizing back exchange in 18O/16O quantitative proteomics experiments by incorporation of immobilized trypsin into the initial digestion step

Differential labeling of peptides via the use of the 18O-water proteolytic labeling method has been widely adopted for quantitative shotgun proteomics studies due to its simplicity and low reagent costs. In this report, the use of immobilized trypsin in the initial digestion step, in addition to the initial digestion step, is explored as a means to minimize postlabeling back exchange of 18O-labeled peptides into the 16O form when multidimensional peptide separation methods (here, isoelectric focusing of peptides) are incorporated into the sample workflow. Examples are shown with a mixture of standard proteins and a sample from an ongoing clinical proteomics study.