DNA is organized by protein:DNA complexes called nucleosomes in eukaryotes. Nucleosomes are composed of 147 base pairs of DNA wrapped around a histone octamer containing two copies of each core histone protein. Histone proteins play significant roles in many nuclear processes including transcription, DNA damage repair and heterochromatin formation. Histone proteins are extensively and dynamically post-translationally modified, and these post-translational modifications (PTMs) are thought to comprise a specific combinatorial PTM profile of a histone that dictates its specific function. Abnormal regulations of PTM may lead to developmental disorders and disease development such as cancer.
Antibodies have been widely used to characterize histones and histone PTMs. However, antibody-based techniques have several limitations. Mass spectrometry (MS) has therefore emerged as the most suitable analytical tool to quantify proteomes and protein PTMs. The most commonly used strategy is still bottom-up MS, and the most widely adopted protocol includes derivatization of lysine residues in histones to allow trypsin to generate Arg-C like peptides (4–20 aa). However, samples such as primary tissues, complex model systems, and biofluids are hard to retrieve in large quantities. Because of this, it is critical to know whether the amount of sample available would lead to an exhaustive analysis if subjected to MS.
In a recent publication, Guo, et al. examined (1) the reproducibility in quantification of histone PTMs using a wide range of starting material: from 50,000 to 5,000,000 cells. They used four different cell lines: HeLa, 293T, human embryonic stem cells (hESCs), and myoblasts. Their results demonstrated that an accurate quantification of abundant histone PTMs can be efficiently obtained by using low-resolution MS and as low as 50,000 cells as starting material Low abundance histone marks showed more variability in quantification when comparing different amounts of starting material, so a larger amount of starting material (at least 500,000 cells) is recommended.
Guo, Q. et al. (2017) Assessment of Quantification Precision of Histone Post-Translational Modifications by Using an Ion Trap and down To 50,000 Cells as Starting Material. J. Proteome Res. 17, 234–42.
Recombinant erythropoietin (rhEPO) is often used as “doping agent” by athletes in endurance sports to increase blood oxygen capacity. Some strategies improve the pharmacological properties of erythropoietin (EPO) through the genetic and chemical modification of the native EPO protein. The EPO-Fcs are fusion proteins composed of monomeric or dimeric recombinant EPO and the dimeric Fc region of human IgG molecules. The Fc region includes the hinge region and the CH2 and CH3 domains. Recombinant human EPOs (rhEPO) fused to the IgG Fc domain demonstrate a prolonged half-life and enhanced erythropoietic activity in vivo compared with native or rhEPO.
Drug-testing agencies will need to obtain primary structure information and develop a reliable analytical method for the determination of EPO-Fc abuse in sport. The possibility of EPO-Fc detection using nanohigh-performance liquid chromatography−tandem mass spectrometry (HPLC−MS/MS) was already demonstrated (1). However, the prototyping peptides derived from EPO and IgG are not selective enough because both free proteins are naturally presented in human serum. In a recent publication, researchers describe the effort to identify peptides covering unknown fusion breakpoints (later referred to as “spacer” peptides; 2). The identification of “spacer” peptides will allow the confirmation of the presence of exogenous EPO-Fc in human biological fluids.
A bottom-up approach and the intact molecular weight measurement of deglycosylated protein and its IdeS proteolytic fractions was used to determine the amino acid sequence of EPO-Fc. Using multiple proteases, peptides covering unknown fusion breakpoints (spacer peptides) were identified.
Results indicated that “spacer peptides” could be used in the determination of EPO-Fc fusion proteins in biological samples using common LC−tandem MS methods.
- Reichel, C. et al. (2012) Detection of EPO-Fc fusion protein in human blood: screening and confirmation protocols for sports drug testing.
Drug Test. Anal. 4, 818−29.
- Mesonzhnik, N. et al. (2017) Characterization and Detection of Erythropoietin Fc Fusion Proteins Using Liquid Chromatography−Mass Spectrometry.
J. of Proteome Res. 17, 689-97.
Protein:protein interactions (PPIs) play a key role in regulating cellular activities including DNA replication, transcription,translation, RNA splicing, protein secretion, cell cycle control and signal transduction. A comprehensive method is needed to identify the PPIs before the significance of the protein:protein interactions can be characterized. Affinity purification−mass spectrometry (AP−MS) has become the method of choice for discovering PPIs under native conditions. This method uses affinity purification of proteins under native conditions to preserve PPIs. Using this method, the protein complexes are captured by antibodies specific for the bait proteins or for tags that were introduced on the bait proteins and pulled down onto immobilized protein A/G beads. The complexes are further digested into peptides with trypsin. The protein interactors of the bait proteins are identified by quantification of the tryptic peptides via mass spectrometry.
The success of AP-MS depends on the efficiency of trypsin digestion and the recovery of the tryptic peptides for MS analysis. Several different protocols have been used for trypsin digestion of protein complexes in AP-MS studies, but no systematic studies have been conducted on the impact of trypsin digestion conditions on the identification of PPIs. A recent publication used NFB/RelA and BRD4 as bait proteins and five different trypsin digestion conditions (two using “on beads” and three using “elution” digestion protocols). Although the performance of the trypsin digestion protocols changed slightly depending on the different bait proteins, antibodies and cell lines used, the authors of the paper found that elution digestion methods consistently outperformed on-beads digestion methods.
Zhang, Y. et al. (2017) Quantitative Assessment of the Effects of Trypsin Digestion Methods on Affinity Purification−Mass Spectrometry-based Protein−Protein Interaction Analysis
J of Proteome. Res. 16, 3068–82.
Bottom-up proteomics focuses on the analysis of protein mixtures after enzymatic digestion of the proteins into peptides. The resulting complex mixture of peptides is analyzed by reverse-phase liquid chromatography (RP-LC) coupled to tandem mass spectrometry (MS/MS). Identification of peptides and subsequently proteins is completed by matching peptide fragment ion spectra to theoretical spectra generated from protein databases.
Trypsin has become the gold standard for protein digestion to peptides for shotgun proteomics. Trypsin is a serine protease. It cleaves proteins into peptides with an average size of 700-1500 daltons, which is in the ideal range for MS (1). It is highly specific, cutting at the carboxyl side of arginine and lysine residues. The C-terminal arginine and lysine peptides are charged, making them detectable by MS. Trypsin is highly active and tolerant of many additives.
Even with these technical features, the use of trypsin in bottom-up proteomics may impose certain limits in the ability to grasp the full proteome, Tightly-folded proteins can resist trypsin digestion. Post-translational modifications (PTMs) present a different challenge for trypsin because glycans often limit trypsin access to cleavage sites, and acetylation makes lysine and arginine residues resistant to trypsin digestion.
To overcome these problems, the proteomics community has begun to explore alternative proteases to complement trypsin. However, protocols, as well as expected results generated when using these alternative proteases have not been systematically documented.
In a recent reference (2), optimized protocols for six alternative proteases that have already shown promise in their applicability in proteomics, namely chymotrypsin, Lys-C, Lys-N, Asp-N, Glu-C and Arg-C have been created.
Data describe the appropriate MS data analysis methods and the anticipated results in the case of the analysis of a single protein (BSA) and a more complex cellular lysate (Escherichia coli). The digestion protocol presented here is convenient and robust and can be completed in approximately in 2 days.
- Laskay, U. et al. (2013) Proteome Digestion Specificity Analysis for the Rational Design of Extended Bottom-up and middle-down proteomics experiments. J of Proteome Res. 12, 5558–69.
- Giansanti, P. et. al. (2016) Six alternative protease for mass spectrometry based proteomics beyond trypsin. Nat. Protocols 11, 993–6
Hadrosaurus skeleton vintage engraving.
Brachylophosaurus was a mid-sized member of the hadrosaurid family of dinosaurs living about 78 million years ago, and is known from several skeletons and bonebed material from the Judith River Formation of Montana and the Oldman Formation of Alberta. Recent fossil evidence indicates structures similar to blood vessels in location and morphology, have been recovered after demineralization of multiple dinosaur cortical bone fragments from multiple specimens, some of which are as old as 80 Ma. These structures were hypothesized to be either endogenous to the bone (i.e., of vascular origin) or the result of biofilm colonizing the empty network after degradation of original organic components (i.e., bacterial, slime mold or fungal in origin). Cleland et al. (1) tested the hypothesis that these structures are endogenous and thus retain proteins in common with extant archosaur blood vessels that can be detected with high-resolution mass spectrometry and confirmed by immunofluorescence. Continue reading
Here we provide two examples of “atypical” experiments that take advantage of the properties of the ProteaseMAX™ Surfactant to improve studies involving digestion of complex protein mixtures.
Clostridium difficile spores are considered the morphotype of infection, transmission and persistence of C. difficile infections. A recent publication (1) illustrated a novel strategy using three different approaches to identify proteins of the exosporium layer of C. difficile spores and complements previous proteomic studies on the entire C. difficile spores. Continue reading
Protein phosphorylation is a very important protein post-translational modification that controls many cellular processes including metabolism, transcriptional and translation regulation, degradation of proteins, cellular signaling and communication, proliferation, differentiation, and cell survival (1). Approximately 35% of human proteins are phosphorylated. Phosphoproteins are low in abundance, and, therefore, are challenging to detect and characterize by mass spectrometry. Different enrichment systems have been developed to isolate phosphopeptides. Among these techniques, immobilized metal affinity chromatography (IMAC) using Fe3+ and Ga3+ has been widely used for the enrichment of phosphopeptides.
Typical experimental workflows are tedious and consist of numerous steps, including sample collection and cell lysis. One of the major challenges of the process is to maintain the in vivo phosphorylation state of the proteins throughout the preparation process
To evaluate the effect of sample collection protocols on the global phosphorylation status of the cell, a recent paper by Kashin et al. compared different sample workflows by metabolic labeling and quantitative mass spectrometry on Saccharomyces cerevisiae cell cultures (2).
Three different sample collection workflows were evaluated: two that used denaturating conditions and involved mixing of cell cultures with an excess of either ethanol (EtOH) at −80 °C or trichloroacetic acid (TCA), and a third under nondenaturing conditions and washing the cells in PBS.
Their data suggest that either TCA or EtOH sample collection protocols introduced lower collection bias than the PBS protocol. It was also suggested that similar studies be carried out to determine what effects sample preparation has on other post translation modifications such as acetylation or ubiquitination.
- Thingholm T.E. et al, (2009) Analytical strategies for phosphoproteomics. Proteomics 9,1451–68
- Kanshin, E. et al. (2015) Sample Collection Method Bias Effects in Quantitative Phosphoproteomics. J Proteome Res. 14, 2998-04.
Filter-aided sample preparation (FASP) method is used for the on-filter digestion of proteins prior to mass-spectrometry-based analyses (1,2). FASP was designed for the removal of detergents, and chaotropes that were used for sample preparation. In addition, FASP removes components such as salts, nucleic acids and lipids. Akylation of reduced cysteine residues is also carried out on filter, after which protein is proteolyzed by use of trypsin on filter in the optimal buffer of the enzyme. Subsequent elution and desalting of the peptide-rich solution then provides a sample ready for LC–MS/MS analysis.
Erde et al. (3) described an enhanced FASP (eFASP) workflow that included 0.2% DCA in the exchange, alkylation, and digestion buffers,thus enhancing trypsin proteolysis, resulting in increases cytosolic and membrane protein representation. DCA has been reported (4) to improve the efficiency of the denaturation, solubilization, and tryptic digestion of proteins, particularly proteolytically resistant myoglobin and integral membrane proteins, thereby enhancing the efficiency of their identification with regard to the number of identified proteins and unique peptides.
In a recent publication (5) traditional FASP and eFASP were re-evaluated by ultra-high-performance liquid chromatography coupled to a quadrupole mass filter Orbitrap analyzer (Q Exactive). The results indicate that at the protein level, both methods extracted essentially the same number of hydrophobic transmembrane containing proteins as well as proteins associated with the cytoplasm or the cytoplasmic and outer membranes.
The LC–MS/MS results indicate that FASP and eFASP showed no significant differences at the protein level. However, because of the slight differences in selectivity at the physicochemical level of peptides, these methods can be seen to be somewhat complementary for analyses of complex peptide mixtures.
- Manza, L. L. et al. (2005) Sample preparation and digestion for proteomic analyses using spin filters Proteomics 5, 1742–74.
- Wiśniewski, J. R. et al. (2009) Universal sample preparation method for proteome analysis Nat. Methods 6, 359–62.
- Erde, J. et al. (2014) Enhanced FASP (eFASP) to increase proteomic coverage and sample recovery for quantitative proteome experiments. J. Proteome Res. 13, 1885–95.
- Lin, Y. et al. (2008) Sodium-deoxycholate-assisted tryptic digestion and identification of proteolytically resistant proteins Anal. Biochem. 377, 259–66.
- Nel. A. et al. (2015) Comparative Reevaluation of FASP and Enhanced FASP methods by LC-MS/MS/ J Proteome Res. 14, 1637–42.
N-Glycosylation is a common protein post-translational modification occurring on asparagine residues of the consensus sequence asparagine-X-serine/threonine, where X may be any amino acid except proline. Protein N-glycosylation takes place in the endoplasmic reticulum (ER) as well as in the Golgi apparatus.
Approximately half of all proteins typically expressed in a cell undergo this modification, which entails the covalent addition of sugar moieties to specific amino acids. There are many potential functions of glycosylation. For instance, physical properties include: folding, trafficking, packing, stabilization and protease protection. N-glycans present at the cell surface are directly involved in cell−cell or cell−protein interactions that trigger various biological responses.
The standard method used to profile the N-glycosylation pattern of cells is glycoprotein isolation followed by denaturation and/or tryptic digestion of the glycoproteins and an enzymatic release of the N-glycans using PNGase F followed by analysis mass spec. This method has been reported to yield high levels of high-mannose N-glycans that stem from both membrane proteins as well as proteins from the ER.(1,2)
For those researchers interested in characterizing only cell surface glycans (i.e., complex N-glycans) a recent reference has developed a model system using HEK-292 cells that demonstrates a reproducible, sensitive, and fast method to profile surface N-glycosylation from living cells (3). The method involves standard centrifugation followed by enzymatic release of cell surface N-glycans. When compared to the standard methods the detection and quantification of complex-type N-glycans by increased their relative amount from 14 to 85%.
- North, S. J. et al. (2012) Glycomic analysis of human mast cells, eosinophils and basophils. Glycobiology. 2012, 22, 12–22.
- Reinke, S. O. et al. (2011) Analysis of cell surface N-glycosylation of the human embryonic
kidney 293T cell line. J. Carbohydr. Chem. 30, 218–232.
- Hamouda, H. et al. (2014) Rapid Analysis of Cell Surface N‑Glycosylation from Living Cells Using Mass Spectrometry. J of Proteome Res. 13, 6144–51.
Therapeutic monoclonal antibodies (mAbs) represent the majority of therapeutics biologics now on the market, with more than 20 mAbs approved as drugs (1–3). During preclinical development of therapeutic antibodies, multiple variants of each antibody are assessed for pharmacokinetic (PK) characteristics across model systems such as rodents, beagles and primates. Ligand-binding assays (LBA) are the standard technology used to perform the PK studies for mAb candidates (4). Ligand-binding assays (LBAs) are methods used to detect and measure a macromolecular interaction between a ligand and a binding molecule. In LBAs, a therapeutic monoclonal antibody is considered to be the ligand, or analyte of interest, while the binding molecule is usually a target protein.
LBAs have certain well-documented limitations (5). Specific assay reagents are often not available early in a program. Interferences from endogenous proteins, antidrug antibodies, and soluble target ligands are potential complicating factors.
Liquid chromatography coupled to tandem mass spectrometry (LC–MS/MS)-based methods represent a viable and complementary addition to LBA for mAb quantification in biological matrixes. LC–MS/MS provides specificity, sensitivity, and multiplexing capability.
A recent reference (6) illustrates an automated method to perform LC–MS/MS-based quantitation, with IgG1 conserved peptides, a heavy isotope labeled mAb internal standard,and anti-human Fc enrichment. The method was applied to the pharmacokinetic study of a mAb dosed in cynomolgus monkey, and the results were compared with the immunoassay data. The interesting finding of the difference between ELISA and LC–MRM-MS data indicated that those two methods can provide complementary information regarding the drug’s PK profile.
- Mao, T. et al. (2013) Top-Down Structural Analysis of an Intact Monoclonal Antibody by Electron Capture Dissociation-Fourier Transform Ion Cyclotron Resonance-Mass Spectrometry. Anal.Chem. 85, 4239–46.
- Weiner, L. M. et al. (2010) Monoclonal antibodies: versatile platforms for cancer immunotherapy. Nat. Rev. Immunol. 10, 317–27.
- Nelson, A. et al. (2010) Development trends for human monoclonal antibody therapeutics. Nat. Rev. Drug Discovery. 9, 767–74.
- DeSilva, B. et al. (2003) Recommendations for the Bioanalytical Method Validation of Ligand-Binding Assays to Support Pharmacokinetic Assessments of Macromolecules. Pharm. Res. 20, 1885–00.
- Ezan, E.et al. (2009) Critical comparison of MS and immunoassays for the bioanalysis of therapeutic antibodies. Bioanalysis 1, 1375–88.
- Zhang, Q. et al. (2014) Generic Automated Method for Liquid Chromatography–Multiple Reaction Monitoring Mass Spectrometry Based Monoclonal Antibody Quantitation for Preclinical Pharmacokinetic Studies. Anal.Chem. 86, 8776–84.