Yanbo Pan

Global Screening of CK2 Kinase Substrates by an Integrated Phosphoproteomics Workflow

Due to its constitutive activity and ubiquitous distribution, CK2 is the most pleiotropic kinase among the individual members of the protein kinase superfamily. Identification of CK2 substrates is vital to decipher its role in biological processes. However, only a limited number of CK2 substrates were identified so far. In this study, we developed an integrated phosphoproteomics workflow to identify the CK2 substrates in large scale. First, in vitro kinase reactions with immobilized proteomes were combined with quantitative phosphoproteomics to identify in vitro CK2 phosphorylation sites, which leaded to identification of 988 sites from 581 protein substrates. To reduce false positives, we proposed an approach by comparing these in vitro sites with the public databases that collect in vivo phosphorylation sites. After the removal of the sites that were excluded in the databases, 605 high confident CK2 sites corresponding to 356 proteins were retained. The CK2 substrates identified in this study were based on the discovery mode, in which an unbiased overview of CK2 substrates was provided. Our result revealed that CK2 substrates were significantly enriched in the spliceosomal proteins, indicating CK2 might regulate the functions of spliceosome.

Depletion of acidic phosphopeptides by SAX to improve the coverage for the detection of basophilic kinase substrates.

The Ser/Thr protein kinases fall into three major subgroups, pro-directed, basophilic, and acidophilic, on the basis of the types of substrate sequences that they preferred. Despite many phosphoproteomics efforts that have been taken for global profiling of phosphopeptides, methodologies focusing on analyzing a particular type of kinase substrates have seldom been reported. Selective enrichment of phosphopeptides from basophilic kinase substrates is difficult because basophilic motifs are cleaved by trypsin during digestion. In this study, we develop a negative enrichment strategy to enhance the identification of basophilic kinase substrates. This method is based on an observation that high pH strong anion exchange (SAX) chromatography can separate tryptic phosphopeptides according to the number of acidic amino acidic residues that they have. Thus, SAX was applied to deplete acidic phosphopeptides from the phosphopeptide mixture, which improved the coverage for the detection of basophilic kinase substrates. The SAX depletion approach was further combined with online SCX-RP separation for large-scale analysis of mouse liver phosphoproteome, which resulted in the identification of 6944 phosphorylated sites. It was found that motifs associated with basophilic kinases prevail for these identified phosphorylated sites.

Protein digestion priority is independent of protein abundances

the expense of percentage time in catastrophe (Fig. 1e), apparent at the 3.3 nM concentration and above. At 1.0 Hz, percentage time in growth increases with Taxol concentration at the expense of percentage time in catastrophe but not pause (Fig. 1e), becoming statistically significant at the 10.0 nM level. The same treatment (indeed, the same data) acquired at different rates leads to opposing conclusions owing to artifacts introduced at slow acquisition rates. Alternative microtubule tracking methods such as laminar wholemicrotubule tracking show results agreeing with those from 4.0-Hz data3,4. These techniques capture the microtubule end position during all phases but are limited to isolated plus-ends near the cell periphery5,6. To avoid the introduction of artifacts in plus-end fluorescent-comet tracking, acquisitions of image stacks for microtubule plus-end dynamics analysis should be performed at the highest frame rate possible.

Integration of Cell Lysis, Protein Extraction, and Digestion into One Step for Ultrafast Sample Preparation for Phosphoproteome Analysis

Conventional sample preparation protocols for phosphoproteome analysis require multiple time-consuming and labor-intensive steps, including cell lysis, protein extraction, protein digestion, and phosphopeptide enrichment. In this study, we found that the presence of a large amount of trypsin in the sample did not interfere with phosphopeptide enrichment and subsequent LC-MS/MS analysis. Taking advantage of fast digestion achieved with high trypsin-to-protein ratio, we developed a novel concurrent lysis-digestion method for phosphoproteome analysis. In this method, the harvested cells were first placed in a lysis buffer containing a huge amount of trypsin. After ultrasonication, the cells were lysed and the proteins were efficiently digested into peptides within one step. Thereafter, tryptic digest was subjected to phosphopeptide enrichment, in which unphosphorylated peptides, trypsin, and other components incompatible with LC-MS/MS analysis were removed. Compared with conventional methods, better phosphoproteome coverage was achieved in this new one-step method. Because protein solubilization and cell lysis were facilitated by fast protein digestion, the complete transformation of cell pellets into the peptide mixture could be finished within 25 min, while it would take at least 16 h for conventional methods. Hence, our method, which integrated cell lysis, protein extraction, and protein digestion into one step, is rapid and convenient. It is expected to have broad applications in phosphoproteomics analysis.

Quantitative proteomics reveals the kinetics of trypsin-catalyzed protein digestion

AbstractTrypsin is the popular protease to digest proteins into peptides in shotgun proteomics, but few studies have attempted to systematically investigate the kinetics of trypsin-catalyzed protein digestion in proteome samples. In this study, we applied quantitative proteomics via triplex stable isotope dimethyl labeling to investigate the kinetics of trypsin-catalyzed cleavage. It was found that trypsin cleaves the C-terminal to lysine (K) and arginine (R) residues with higher rates for R. And the cleavage sites surrounded by neutral residues could be quickly cut, while those with neighboring charged residues (D/E/K/R) or proline residue (P) could be slowly cut. In a proteome sample, a huge number of proteins with different physical chemical properties coexists. If any type of protein could be preferably digested, then limited digestion could be applied to reduce the sample complexity. However, we found that protein abundance and other physicochemical properties, such as molecular weight (Mw), grand average of hydropathicity (GRAVY), aliphatic index, and isoelectric point (pI) have no notable correlation with digestion priority of proteins. Graphical AbstractSequence logos of four cleavage site types with different kinetics (very fast, fast, slow, and very slow sites)

N‐Terminal Labeling of Peptides by Trypsin‐Catalyzed Ligation for Quantitative Proteomics

Labeling tryptic peptides with stable isotopes is one of the most important methods for quantitative proteomics, and many ingenious chemical labeling strategies have been developed. Incorporation of one isotopically labeled tag onto a peptide terminus represents an ideal labeling approach, as it would simplify the interpretation of mass spectra. Moreover, the absence of labels on the side chains would facilitate the quantification of post-translational modifications (PTMs). However, to date all the reported chemical labeling strategies, including dimethyl labeling, iTRAQ (isobaric tags for absolute and relative quantification), and ICAT (isotope-coded affinity tags), result in the modification of side chains. One promising method to achieve the incorporation of a single tag is enzymatic labeling. Proteolytic O labeling can specifically label tryptic peptide termini without modification of side chains, but the small change in mass and O/O back-exchange, i.e. the exchange of O with O after the labeling process hinder its wide application in quantitative proteomics. Herein, we report a novel enzymatic labeling approach, in which trypsin is used as a ligase to specifically incorporate amino acids labeled with stable isotopes onto the N termini of peptides for quantitative analysis. Trypsin is a serine protease that specifically hydrolyzes peptide bonds in proteins after arginine and lysine residues. However, trypsin also catalyses peptide synthesis in organic solvents. Therefore, it is possible to covalently link isotopically labeled amino acids to tryptic peptides by using trypsin as a ligase. In this study, arginine, the prototype substrate for trypsin, was used as an acyl moiety donor. The primary amine group of arginine was protected with a benzoyl group (Bz) to prevent the formation of dipeptides or oligopeptides, and the carboxy group was esterified with ethanol to activate the acyl donor (see the Supporting Information, Figure S1). The final product, Na-benzoyl-l-arginine ethyl ester (Bz–R–OEt), was used as a substrate for the trypsin-catalyzed ligation. The quantitative proteomics workflow based on the trypsincatalyzed N-terminal labeling is shown in Figure 1a. Trypsin was first used as a protease to digest proteins in an aqueous solution (1m urea/50 mm Tris-HCl, pH 8.0). Then, the generated tryptic peptides were lyophilized and transferred to an ethanol solution containing 4% aqueous buffer (0.1m TrisHCl, pH 8.0). Next, trypsin immobilized on magnetic nanoparticles (IM-trypsin) and Bz–R–OEt were added to the ethanol solution for the N-terminal labeling of peptides with Bz–R based on a kinetically controlled mechanism (see the Supporting Information). Finally, quantification of the proteins was achieved by the differential labeling of two samples by using Bz–R–OEt (light label) and Bz–(C6)R–OEt (bearing six C atoms; heavy label), the incorporation of which are indicated by mass shifts of 260 and 266 Da, respectively. We first validated the protease and ligase activities of trypsin by using the synthetic peptide VGKANEELAGVVAEVQK (Figure 1b; m/z= 1740.86), which contains one trypsin cleavage site. In aqueous solution, treatment with IMtrypsin generated a shorter peptide ANEELAGVVAEVQK (Figure 1b, m/z= 1456.82). Treatment of the obtained peptide with an ethanol solution containing Bz–R–OEt and IMtrypsin gave an N-terminus labeled peptide Bz–RANEELAGVVAEVQK (Figure 1b, m/z= 1716.9). Similar results were also obtained for three other synthetic peptides that contained one trypsin cleavage site (Supporting Information, Figure S2). These examples clearly illustrate that trypsin functions as a protease in an aqueous solution whereas it acts as an N-terminus ligase in an ethanol solution. Thus, these results imply that this enzymatic labeling approach can be used to label peptides generated by trypsin digestion of a proteome sample. In this study, free trypsin was used for the digestion of proteins, as is conventional in proteomics analysis, whereas IM-trypsin (trypsin from the same source) was used for ligation because it is more tolerant to organic solvents and it is readily removed using magnetism. We tested whether the labeling of side-chain primary amino groups would also be facilitated by trypsin ligase. A synthetic peptide VIFIEHAKRKG, containing two sidechain amino groups and one terminal primary amino group, was labeled by trypsin. The Arg tag was incorporated only onto the terminal primary amino group and not onto the sidechain amino groups (see the Supporting Information, Figure S3a). These results are markedly different from those of other amine labeling approaches. For example, for labeling [*] Y. Pan, Prof. Dr. M. Ye, Dr. L. Zhao, K. Cheng, M. Dong, C. Song, H. Qin, Dr. F. Wang, Prof. Dr. H. Zou CAS Key Lab of Separation Sciences for Analytical Chemistry National Chromatographic R&A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences Dalian 116023 (China) E-mail: mingliang@dicp.ac.cn hanfazou@dicp.ac.cn

Trypsin-Catalyzed N-Terminal Labeling of Peptides with Stable Isotope-Coded Affinity Tags for Proteome Analysis

An enzymatic approach to label peptide N-termini with isotope-coded affinity tags is presented. This method exploits the high activity of trypsin for peptide synthesis in organic solvents. A cosubstrate containing a stable isotope-coded Arg residue and a biotin tag was synthesized. When the cosubstrate was incubated with tryptic peptides and trypsin in ethanol solution, the stable isotope-coded affinity tag was specifically coupled onto the N-termini of peptides via the formation of new peptide bonds. The labeled peptides were specifically enriched by avidin affinity chromatography and then were submitted to liquid chromatography-tandem mass spectrometry (LC/MS/MS) for quantification. This enrichment step effectively reduced the interference by unlabeled peptides. The excellent performance of this approach was demonstrated by labeling standard peptides as well as a mouse liver digest. In addition to one amino acid residue, a few dipeptide tags were also introduced to the N-termini of peptides successfully by this enzymatic approach. It was found that the identifications for samples labeled with these tags were highly complementary. Coupling a short sequence tag onto peptides could be an effective approach to improve the coverage for proteome analysis.

Protein Arginine Allylation and Subsequent Fluorophore Targeting

Protein allylation and fluorophore targeting: Arginine residues of the yeast nuclear ribonucleoprotein Npl3 were extensively modified by Hmt1-catalyzed allylation reaction with allyl-SAM as the allyl group donor. The allylated protein was further treated with tetrazole compounds under UV irradiation, leading to formation of protein-attached fluorescent products.

The proteomic analysis improved by cleavage kinetics-based fractionation of tryptic peptides.

Selective enrichment of specific peptides is an effective way to identify low abundance proteins. Fractionation of peptides prior to mass spectrometry is another widely used approach to reduce sample complexity in order to improve proteome coverage.In this study, we designed a multi‐stage digestion strategy to generate peptides with different trypsin cleavage kinetics. It was found that each of the collected peptide fractions yielded many new protein identifications compared to the control group due to the reduced complexity. The overlapping peptides identified between adjacent fractions were very low, indicating that each fraction had different sets of peptides. The multi‐stage digestion strategy separates tryptic peptides with different cleavage kinetics while RPLC separates peptides with different hydrophobicity. These two separation strategies were highly orthogonal, and showed an effective multidimensional separation to improve proteome coverage.

High Concentration Trypsin Assisted Fast In-Gel Digestion for Phosphoproteome Analysis

Abstract Polyacrylamide gel electrophoresis (PAGE) is a powerful protein separation technology. Combined with mass spectrometry, it could identify thousands of proteins in proteomics analysis. However, the time-consuming procedure restricts its broad applications in proteomics study. In this work, it was found that high concentration of trypsin did not compromise subsequent phosphopeptide enrichment after in-gel digestion, but could promote the in-gel digestion. Hence, a new, fast and robust digestion method was established. Firstly, 50 μg of HeLa cell protein was separated into 5 fractions by SDS-PAGE, and then the proteins were digested with high concentration trypsin for 30 min. Finally, the phosphopeptides were enriched by Ti-IMAC Tip. After liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis, about 2000 phosphorylation sites were identified in the experimental group, while less than 1500 phosphorylation sites were identified in control group. With the aid of high concentration of trypsin, in-gel digestion could be completed within only 30 min, and more phosphorylation site identifications and lower percentage of missed cleavages were acquired than those in control experiments. The experiment results demonstrated that high concentration of trypsin could not only accelerate the in-gel digestion, but also improve the phosphoproteome coverage.