The spike protein of the SARS-CoV-2 virus is a very commonly researched target in COVID-19 vaccine and therapeutic studies because it is an integral part of host cell entry through interactions between the S1 subunit of the spike protein with the ACE2 protein on the target cell surface. Viral proteins important in host cell entry are typically highly glycosylated. Looking at the sequence of the SARS-CoV-2 virus, researchers predict that the spike protein is highly glycosylated. In a recent study, researchers conducted a glycosylation analysis of SARS-CoV-2 proteins using mass spec analysis to determine the N-glycosylation profile of the subunits that make up the spike protein.
Glycans assist in protein folding and help the virus avoid immune recognition by the host. Glycosylation can also have an impact on the antigenicity of the virus, as well as potential effects on vaccine safety and efficacy. Mass spectrometry is widely used for viral characterization studies of influenza viruses. Specifically, mass spec has been used to study influenza protein glycosylation, antigen quantification, and determination of vaccine potency.
In older people, low muscle mass is strongly associated with reduced functional capacity and an increased risk of disability. Myostatin is a negative regulator of muscle growth and has become an important target for pharmaceutical companies designing therapeutics to address age-associated muscle loss.
Anti-myostatin drugs increase muscle size and strength in preclinical studies. Fortetropin is a proteo-lipid complex made from fertilized egg yolk and shows anti-myostatin activity. When Fortetropin is provided as a supplement, lowered circulating myostatin levels are observed both in rodents and in young men. Fortetropin in combination with resistance exercise also lowers myostatin and increased lean body mass.
Sometimes, when using trypsin to study a protein sequence or protein modifications, sequence coverage just isn’t quite as complete as you’d like. Looking for a protease with novel cleavage specificity or a protease that functions well in a low pH environment? Promega has a protease for that.
ProAlanase is a new site-specific endoprotease that preferentially cleaves proteins on the C-terminal side of proline and alanine amino acids. The unique cleavage specificity of ProAlanase (also known as An-PEP or EndoPro; 1–3) can help to uncover parts of the proteome not previously accessible with proteases typically used in proteomic studies.
Glycosylation is the process by which a carbohydrate is covalently attached totarget macromolecules, typically proteins. This modification serves various functions including guiding protein folding (1,2), promoting protein stability (2), and participating signaling functions (3).
SARS-CoV-2 utilizes an extensively glycosylated spike (S) protein that protrudes from the viral surface to bind to angiotensin-converting enzyme 2 (ACE2) to mediate host-cell entry. Vaccine development has been focused on this protein, which is the focus of the humoral immune response. Understanding the glycan structure of the SARS-CoV-2 virus spike (S) protein will be critical in the development of glycoprotine-based vaccine candidates.
The use of mass spectrometry for the characterization of individual or complex protein samples continues to be one of the fastest growing fields in the life science market.
Bottom-up proteomics is the traditional approach to address these questions. Optimization of each the individual steps (e.g. sample prep, digestion and instrument performance) is critical to the overall success of the entire experiment.
To address issues that may arise in your experimental design, Promega has developed unique tools and complementary webinars to help you along the way.
Here you can find a summary of individual webinars for the following topics:
Large-scale analyses of the proteome have revealed proteomic changes in response to disease, and these changes hold great promise for diagnostics and treatment of complex disease if proteomic analysis can be brought into the clinical laboratory. Successful and reliable large-scale proteomics requires sample preparation workflows that are reproducible, reliable and show little variability. To bring proteomics into the clinical laboratory, standardized procedures and workflows for sample prep and analysis are required to generate valid, actionable results on a time scale useful for the clinic.
The two most common sample types analyzed for clinical proteomics are body fluids and tissue biopsies. To process these kinds of samples, there are two initial steps: tissue solubilization, followed by proteolytic digestion. Solubilization of solid tissues is the most labor-intensive and produces the most variable results.
The introduction of pressure cycling technology (PCT) using Barocycler instrumentation has greatly improved both tissue solubilization and digestion consistency. The PCT-based sample preparation protocols generally utilize urea as a lysis buffer for protein denaturing and solubilization. Urea has several drawbacks including inhibiting trypsin activity and introducing unwanted modifications like carbamylation.
Lucas and colleagues analyzed whether replacing urea with SDC would produce similar tissue digestion profiles and improve the PCT method.
SDC allowed the use of higher temperatures compared to urea, and hence the first step (lysis, reduction, and alkylation) was performed at 56 °C. The second digestion step in the Barocycler was optimized, and the third step was eliminated. To further reduce digestion time, they capitalized on Rapid Trypsin/Lys-C. Rapid Trypsin/Lys-C maintains robust activity at 70 °C, and allowed Barocycler digestion to be performed in a single step, completing digestion in 30 cycles (approximately 30 min) rather than 105 minutes, streamlining the protocol.
The data presented an improved conventional tissue PCT approach in a Barocycler by replacing urea and proteolytic enzymes with SDC, N-propanol, and modified commercially available enzymes that have higher optimum temperatures.
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.
The art of brewing alcoholic beverages has existed for thousands of years. The process of beer brewing begins with barley grains, which are malted to allow partial germination, triggering expression of key enzymes. The germinated grains are then dried and milled. Next, starch, proteins, and other molecules are solubilized during mashing. During mashing, solubilized enzymes degrade starch to fermentable sugars, and digest proteins to produce peptides and free amino acids. Fermentable sugars and free amino acids are required for efficient yeast growth during fermentation.
After the mash, the wort is removed, and hops are added for bitterness and aroma, and the wort is boiled. After boiling, the wort is inoculated with yeast, and fermentation proceeds to produce bright beer. Typically this bright beer is then filtered, carbonated, packaged, and sold. Many proteins originating from the barley grain and the yeast are present in beer, and these have been reported to affect the quality of the final product. However, some of the biochemical details of this process remain unclear. To better understand what happens during the various steps of the brewing process, Schultz et al. used mass spectrometry proteomics to perform a global untargeted analysis of the proteins present across time during beer production and described this work in a recent paper (1). Samples analyzed included sweet wort produced by a high temperature infusion mash, hopped wort, and bright beer. Continue reading “Beer Is Complicated: Proteome Analysis via Mass Spectrometry”
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.
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