Current Protocols in Protein Science最新文献

While native proteins cover diverse structural spaces and achieve various biological events, not many of them can directly serve human needs. One reason is that the native proteins usually contain idiosyncrasies evolved for their native functions but disfavoring engineering requirements. To overcome this issue, one strategy is to create de novo proteins which are designed to possess improved stability, high environmental tolerance, and enhanced engineering potential. Compared to other protein engineering strategies, in silico design of de novo proteins has significantly expanded the protein structural and sequence spaces, reduced wet lab workload, and incorporated engineered features in a guided and efficient manner. In the Baker laboratory we have been applying a design pipeline that uses the blueprint builder to design different folds of de novo proteins, and have successfully obtained libraries of de novo proteins with improved stability and engineering potential. In this article, we will use the design of de novo β-barrel proteins as an example to describe the principles and basic procedures of the blueprint builder−based design pipeline. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: The construction of blueprints

Alternate Protocol: Build blueprints based on existing protein .pdb files

Basic Protocol 2: De novo protein design pipeline using the blueprint builder

Cell-free protein synthesis is a powerful tool for engineering biology and has been utilized in many diverse applications, from biosensing and protein prototyping to biomanufacturing and the design of metabolic pathways. By exploiting host cellular machinery decoupled from cellular growth, proteins can be produced in vitro both on demand and rapidly. Eukaryotic cell-free platforms are often neglected due to perceived complexity and low yields relative to their prokaryotic counterparts, despite providing a number of advantageous properties. The yeast Pichia pastoris (also known as Komagataella phaffii) is a particularly attractive eukaryotic host from which to generate cell-free extracts, due to its ability to grow to high cell densities with high volumetric productivity, genetic tractability for strain engineering, and ability to perform post-translational modifications. Here, we describe methods for conducting cell-free protein synthesis using P. pastoris as the host, from preparing the cell lysates to protocols for both coupled and linked transcription-translation reactions. By providing these methodologies, we hope to encourage the adoption of the platform by new and experienced users alike. © 2020 The Authors.

Basic Protocol 1: Preparation of Pichia pastoris cell lysate

Basic Protocol 2: Coupled in vitro transcription and translation

Basic Protocol 3: Determining luciferase production from cell-free protein synthesis reactions

Alternate Protocol 1: Linked in vitro transcription and translation

Alternate Protocol 2: Quantifying HSA protein concentration

Support Protocol 1: Preparation of mRNA by in vitro transcription for linked transcription and translation

Glycosylated proteins, namely glycoproteins and proteoglycans (collectively called glycoconjugates), are indispensable in a variety of biological processes. The functions of many glycoconjugates are regulated by their interactions with another group of proteins known as lectins. In order to understand the biological functions of lectins and their glycosylated binding partners, one must obtain these proteins in pure form. The conventional protein purification methods often require long times, elaborate infrastructure, costly reagents, and large sample volumes. To minimize some of these problems, we recently developed and validated a new method termed capture and release (CaRe). This method is time-saving, precise, inexpensive, and it needs a relatively small sample volume. In this approach, targets (lectins and glycoproteins) are captured in solution by multivalent ligands called target capturing agents (TCAs). The captured targets are then released and separated from their TCAs to obtain purified targets. Application of the CaRe method could play an important role in discovering new lectins and glycoconjugates. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Preparation of crude extracts containing the target proteins from soybean flour

Alternate Protocol 1: Preparation of crude extracts from Jack bean meal

Alternate Protocol 2: Preparation of crude extracts from the corms of Colocasia esculenta, Xanthosoma sagittifolium, and from the bulbs of Allium sativum

Alternate Protocol 3: Preparation of Escherichia coli cell lysates containing human galectin-3

Alternate Protocol 4: Preparation of crude extracts from chicken egg whites (source of ovalbumin)

Basic Protocol 2: Preparation of 2% (v/v) red blood cell suspension

Basic Protocol 3: Detection of lectin activity of the crude extracts

Basic Protocol 4: Identification of multivalent inhibitors as target capturing agents by hemagglutination inhibition assays

Basic Protocol 5: Testing the capturing abilities of target capturing agents by precipitation/turbidity assays

Basic Protocol 6: Capturing of targets (lectins and glycoproteins) in the crude extracts by target capturing agents and separation of the target-TCA complex from other components of the crude extracts

Basic Protocol 7: Releasing the captured targets (lectins and glycoproteins) by dissolving the complex

Basic Protocol 8: Separation of the targets (lectins and glycoproteins) from their respective target capturing agents

Basic Protocol 9: Verification of the purity of the isolated targets (lectins or glycoproteins)

The development of new technologies for the efficient expression of recombinant hemoglobin (rHb) is of interest for experimental studies of protein biochemistry and the development of cell-free blood substitutes in transfusion medicine. Expression of rHb in Escherichia coli host cells has numerous advantages, but one disadvantage of using prokaryotic systems to express eukaryotic proteins is that they are incapable of performing post-translational modifications such as NH₂-terminal acetylation. One possible solution is to coexpress additional enzymes that can perform the necessary modifications in the host cells. Here, we report a new method for synthesizing human rHb with proper NH₂-terminal acetylation. Mass spectrometry experiments involving native and recombinant human Hb confirmed the efficacy of the new technique in producing correctly acetylated globin chains. Finally, functional experiments provided insights into the effects of NH₂-terminal acetylation on O₂ binding properties. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Gene synthesis and cloning the cassette to the expression plasmid

Basic Protocol 2: Selection of E. coli expression strains for coexpression

Basic Protocol 3: Large-scale recombinant hemoglobin expression and purification

Support Protocol 1: Measuring O₂ equilibration curves

Support Protocol 2: Mass spectrometry to confirm NH₂-terminal acetylation

The small ubiquitin-like modifier (SUMO) is an important post-translational modifier that regulates various cellular processes. Extensive investigations have been made to comprehend the enzymatic process and consequence of SUMOylation. In vitro SUMOylation assays are invaluable for understanding the fundamental mechanisms of SUMOylation. A majority of these assays monitor changes in the size of the substrate upon SUMO conjugation. Current methods typically detect the size difference through SDS-PAGE and western blots, which makes these methods cumbersome, error-prone, and time-consuming. Here, we describe a fluorescence-based assay for real-time detection of SUMOylation. In the method, a fluorophore-tagged substrate is used in the SUMOylation reaction. Upon SUMOylation, the size and correlation time (τ_c) of the substrate increases, and so does its anisotropy. The rate of change in anisotropy with time reflects the efficiency of the SUMOylation machinery. The real-time SUMOylation assay protocol is elegant, time-saving, and less prone to errors. © 2020 Wiley Periodicals LLC.

Basic Protocol: Fluorescent anisotropy-based in vitro SUMOylation assay

Sedimentation velocity analytical ultracentrifugation is a powerful classical method to study protein self-association processes in solution based on the size-dependent macromolecular migration in the centrifugal field. This technique can elucidate the assembly scheme, measure affinities ranging from picomolar to millimolar K_d, and in favorable cases provide information on oligomer lifetimes and hydrodynamic shape. The present step-by-step protocols detail the essential steps of instrument calibration, experimental setup, and data analysis. Using a widely available commercial protein as a model system, the protocols invite replication and comparison with our results. A commentary discusses principles for modifications in the protocols that may be necessary to optimize application of sedimentation velocity analysis to other self-associating proteins. ©2020 Wiley Periodicals LLC.

Basic Protocol 1: Measurement of external calibration factors

Basic Protocol 2: Sedimentation velocity experiment for protein self-association

Basic Protocol 3: Sedimentation coefficient distribution analysis in SEDFIT and isotherm analysis in SEDPHAT

Peripheral membrane proteins participate in numerous biological pathways. Thus, methods to analyze their membrane-binding characteristics have become important. In this report, we detail protocols for the synthesis and utilization of a photoactivable fluorescent lipid as a reporter to monitor membrane binding of proteins. The assay, referred to as proximity-based labeling of membrane-associated proteins (PLiMAP), is based on UV activation of a fluorescent lipid reporter, which in turn crosslinks with proteins bound to membranes and renders them fluorescent. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Synthesis of BODIPY-diazirine phosphatidylethanolamine (BDPE)

Basic Protocol 2: Preparation of BDPE-containing liposomes

Basic Protocol 3: Performing PLiMAP with a candidate protein

Basic Protocol 4: Quantitation of liposome-binding properties of the candidate protein from analyzing in-gel fluorescence

Support Protocol: Purification of GST-2×P4M domain of SidM protein

Although various affinity chromatography and photoaffinity labeling methods have been developed for target protein identification of bioactive molecules, it is often difficult to detect proteins that bind the ligand with weak transient affinity using these techniques. We have developed single electron transfer–mediated tyrosine labeling using ruthenium photocatalysts. Proximity labeling using 1-methyl-4-aryl-urazole (MAUra) labels proteins in close proximity to the photocatalyst with high efficiency and selectivity. Performing this labeling reaction on affinity beads makes it possible to label proteins that bind the ligand with weak transient affinity. In this article, novel protocols are described for target protein identification using photocatalyst proximity labeling on ruthenium photocatalyst-functionalized magnetic affinity beads. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Synthesis of ruthenium photocatalyst

Basic Protocol 2: Synthesis of azide- or desthiobiotin-conjugated labeling reagents

Basic Protocol 3: Preparation of photocatalyst and ligand-functionalized affinity beads

Basic Protocol 4: Target protein labeling in cell lysate

Basic Protocol 5: Enrichment of labeled proteins with MAUra-DTB for LC-MS/MS analysis

Basic Protocol 6: 2D-DIGE analysis of fluorescence-labeled proteins

Cilia and flagella play essential roles in environmental sensing, cell locomotion, and development. These organelles possess a central microtubule–based structure known as the axoneme, which serves as a scaffold and is crucial for the function of cilia. Despite their key roles, the biochemical and biophysical properties of the ciliary proteins are poorly understood. To address this issue, we have developed a novel method to purify functional tubulins from different parts of the axoneme, namely the central pair and B-tubule. We use the biflagellate green alga Chlamydomonas reinhardtii, a model organism for studying cilia due to the conserved structure of this organelle, availability of genetic tools and a large collection of mutant strains. Our method yields highly purified functional axonemal tubulins in sufficient quantities to be used for in vitro biochemical and biophysical studies, such as microtubule dynamic assays. It takes 7 to 8 days to grow enough cells; the isolation of the flagella and the purification of the axonemal tubulins require an additional two full days.© 2020 Wiley Periodicals LLC.

Basic Protocol 1: Growth and harvest of large volume of cell culture

Support Protocol: Assembly of homemade concentration apparatus

Basic Protocol 2: Isolation of flagella

Basic Protocol 3: Tubulin extraction and purification