You have identified and cloned your protein of interest, but you want to explore its function. A protein fusion tag might help with your investigation. However, choosing a tag for your protein depends on what experiments you are planning. Do you want to purify the protein? Would you like to identify interacting proteins by performing pull-down assays? Are you interested in examining the endogenous biology of the protein? Here we cover the advantages and disadvantages of some protein tags to help you select the one that best suits your needs.
The most commonly used protein tags fall under the category of affinity tags. This means that the tag binds to another molecule or metal ion, making it easy to purify or pull down your protein of interest. In all cases, the tag will be fused to your protein of interest at either the amino (N) or carboxy (C) terminus by cloning into an expression vector. This protein fusion can then be expressed in cells or cell-free systems, depending on the promoter the vector contains. Continue reading “Choosing a Tag for Your Protein”
It’s time to analyze your protein and you are trying to decide where to begin. You are asking questions like: Which protease do I choose? How much enzyme should I use in my digest? How long should I perform my digest?
Unfortunately, there is no one-size fits all answer to this type of question other than… “well it depends.” All protease digests will be a balance between denaturing the protein sample to allow access to cleavage sites, optimizing conditions for the protease to function, and compatibility with your workflow and downstream applications. We provide general guidelines that work for most samples, but frequently you will need to optimize the conditions need for your specific sample and application.
Asp-N is a endoproteinase hydrolyzes peptide bonds on the N-terminal side of aspartic residues. The native form is isolated from Pseudomonas fragi. The majority of vendors currently provide a commercial product that consists of 2µg of lyophilized material in a flat bottom vial, and sold for $175–200 US. Formatting such a small amount of material in flat bottom vial can lead to inconsistent resuspension of the protease. Inconsistent working concentrations will lead to non-reproducible data. The current high price also prohibits large-scale use.
The new recombinant Asp-N protease is cloned from Stenotrophomonas maltophilia and expressed in E. coli. Recombinant Asp-N has similar amino acid cleavage specificity as compared to native Asp-N. Digestion of a yeast extract with native and recombinant Asp-N produces very similar results. Providing 10µg lyophilized material in V-shaped vial with a visible cake enables more consistent re-suspension resulting in reproducible data. Due to improved yields the list price is now approximately 40% less when compared to native enzyme. Learn more about this new recombinant Asp-N protease.
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.
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.
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.
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.
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 “Dino Protein: New Methods for Old (Very) Samples”
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.
Example 1 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 “ProteaseMAX: A Surfactant for the Most Complex Mixtures”
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.