To establish a standardized technical framework for large‐scale production of recombinant collagen, we have employed Pichia pastoris GS115 as a host expression system and optimized the fermentation and purification processes for a transdermal peptide–type III collagen fusion protein (transdermal peptide–hCOL3A) at the 15‐L bioreactor scale. A recombinant vector encoding the fusion gene was constructed through codon optimization and transformed into the GS115 strain. The seed culture was prepared via shake‐flask cultivation in BMGY medium. High‐cell‐density fed‐batch fermentation was conducted in a 15‐L fermenter with an inoculum size ranging from 3% to 10%, achieving a final recombinant protein yield of 1.9 g/L. In the downstream purification, optimization of membrane‐based concentration parameters and the incorporation of strong cation‐exchange chromatography using an SP column enhanced the recovery rate of the target protein to 68%. Gel‐filtration analysis demonstrated that transdermal peptide–hCOL3A exists as a trimer, and a CCK‐8 assay revealed that it promotes proliferation of HaCaT cells in vitro , indicating biological functionality. This work successfully establishes a robust and scalable collagen production process suitable for pilot‐scale manufacturing and provides a solid technical foundation for future industrial‐scale translation. © 2025 Wiley Periodicals LLC.
High-throughput screening (HTS) of proteins is used in a wide range of applications across the biology, biotechnology, and medicine disciplines. These include yield optimization, drug or biomarker discovery, and protein engineering, among others. Factors that need to be considered in designing high-throughput protein expression and screening methods (be that for expression, activity, stability, or binding assays), include the required yield, reproducibility, solubility, stability, purity, and activity of the protein. Thus, larger culture volumes and time-consuming manual protein extraction and purification steps are normally required to produce enough protein of appropriate purity. This limits the type of assay and number of protein variants that can be simultaneously tested in an experiment. Here, we describe a HTS protocol that allows the overnight expression, export, and assay of recombinant proteins from Escherichia coli cells in the same microplate well. The protocol uses a recently described Vesicle Nucleating peptide (VNp) technology that promotes high yield vesicular export of functional proteins from E. coli into the culture medium. The resulting protein is of sufficient purity and yield that it can be used directly in plate-based enzymatic assays without additional purification. This simple single-plate protocol allows itself to a wide range of high-throughput research and development screening applications, ranging from streamlining protein production and identification of activity enhancing mutations, to ligand screening for basic research, biotechnological and drug discovery applications. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol: Expression, export, and isolation of vesicular-packaged recombinant protein Support Protocol 1: 96-well plate cold-shock transformation Support Protocol 2: In-plate affinity-tag protein purification Support Protocol 3: Example in-plate enzymatic assay.
Recombinant monoclonal antibodies (mAbs) are widely used across therapeutic areas, and their glycosylation plays a critical role in product quality, manufacturing consistency, and biosimilarity assessment. A middle-down nuclear magnetic resonance (NMR) method has been developed to profile mAb glycan distribution while preserving the covalent bond between glycan and mAb domains, e.g., the fragment crystallizable (Fc) domain. The workflow uses the immunoglobulin-degrading enzyme from Streptococcus pyogenes (IdeS) to generate Fc fragments that are purified, denatured, and dissolved in urea prior to high-resolution two-dimensional 1H-13C heteronuclear single quantum coherence (2D 1H-13C HSQC) NMR spectrum collection. The resulting anomeric peak distribution reveals major and minor glycan species, including the trimannosyl core, high-mannose variants, and branch-specific galactosylation. Compared with traditional glycan mapping, which requires enzymatic cleavage on glycan and liquid chromatography separation, middle-down NMR provides a non-invasive analysis that preserves glycan integrity and enables comprehensive, semi-quantitative monosaccharide profiling. The method requires 3 to 4 days with ∼4 to 5 hr of hands-on time and can be readily implemented in regulated environments for development and quality control. Basic biochemistry and 2D NMR skills are enough to efficiently apply this protocol in a streamlined workflow. Published 2025. This article is a U.S. Government work and is in the public domain in the USA. Basic Protocol 1: mAb-Fc sample preparation Basic Protocol 2: 2D NMR of HSQC peak profile.
Chromatin remodeling is a process of regulating gene expression levels by altering chromatin structure, facilitated by chromatin remodeling factors. ISWI (Imitation SWItch), a type of chromatin remodeling factor, plays a crucial role in epigenetic regulatory processes, such as cell differentiation and development. It primarily achieves remodeling effects by promoting the direct sliding of nucleosomes and their relocation on DNA. To study the function and structural characteristics of ISWI, this basic protocol describes a simple and rapid procedure for the large-scale expression and purification of soluble CtISWI77-1038 and CtATPase77-722 proteins in Escherichia coli (E. coli). The procedure provides high yields (∼20 mg) and is followed by in vitro measurement of ATP hydrolysis activity. © 2025 Wiley Periodicals LLC. Basic Protocol: Expression, purification, and ATP hydrolysis activity measurement of CtISWI and CtATPase.
Neutrophil extracellular traps (NETs) are pivotal in the immune response, trapping and neutralizing pathogens. The process of NET formation, or NETosis, is a critical innate immune response mechanism implicated in diverse physiological and pathological processes, including infections, chronic inflammation, and cancer. Accurate imaging of NETosis is essential for unraveling its mechanisms and therapeutic implications, although limitations of specificity and flexibility remain. In this article, we present modified nanobodies targeting Ly6C/G and histones for imaging of NETosis. Single-domain antibodies (VHHs), also known as nanobodies, targeting Ly6C/G specifically bind to neutrophils, and anti-histone nanobodies selectively target NET-associated histones, enabling NETosis monitoring. These nanobodies are engineered to optimize fluorescent probe labeling and can enable high-resolution visualization of NETosis through whole-body fluorescence tomography and/or intravital microscopy. Our approach offers an alternative for expanding the existing NETosis imaging toolkit. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Cloning, cytoplasmic expression, and purification of proteins Basic Protocol 2: Fluorophore binding assay Support Protocol 1: Analysis by gel staining and immunoblotting Support Protocol 2: In-gel tryptic digestion and sample preparation for LC-MS/MS analysis Support Protocol 3: SpyCatcher003/SpyTag003 binding assay.
Since the birth of biochemistry, researchers have investigated the structure-function relationship of a wide variety of proteins. However, until recently, when X-ray free-electron lasers (XFELs) became available, it was not possible to visualize the motion of proteins from moment to moment with excellent temporal and spatial resolution. Here, we introduce practical methods to visualize protein motions at room temperature using serial femtosecond crystallography (SFX) using XFELs. With the development of this technology, it will be possible to visualize the entire reaction mechanism of many proteins in the future. We first outline a streamlined microcrystallization workflow for hen egg-white lysozyme, enabling rapid detector calibration and data-collection optimization. Next, we present a rotational seeding approach refined on copper-containing nitrite reductase that yields homogeneous microcrystals suitable for high-resolution SFX and readily adaptable to other challenging targets. Finally, we describe a time-resolved strategy combining microcrystals of fungal nitric-oxide reductase with photolabile caged substrates and synchronized UV triggering, capturing catalytic intermediates on the millisecond timescale. Together, these procedures enable investigators to progress from preparing samples to capturing dynamic structural snapshots. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Microcrystallization of lysozyme Basic Protocol 2: Microcrystallization of copper-containing nitrite reductase Basic Protocol 3: Time-resolved serial femtosecond crystallography.
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 Kd , 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.
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.
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.
Histones are the major proteinaceous component of chromatin in eukaryotic cells and an important part of the epigenome, affecting most DNA‐related events, including transcription, DNA replication, and chromosome segregation. The properties of histones are greatly influenced by their post‐translational modifications (PTMs), over 200 of which are known today. Given this large number, researchers need sophisticated methods to study histone PTMs comprehensively. In particular, mass spectrometry (MS)−based approaches have gained popularity, allowing for the quantification of dozens of histone PTMs at once. Using these approaches, even the study of co‐occurring PTMs and the discovery of novel PTMs become feasible. The success of MS‐based approaches relies substantially on obtaining pure and well‐preserved histones for analysis, which can be difficult depending on the source material. Caenorhabditis elegans has been a popular model organism to study the epigenome, but isolation of pure histones from these animals has been challenging. Here, we address this issue, presenting a method for efficient isolation of pure histone proteins from C. elegans at good yield. Further, we describe an MS pipeline optimized for accurate relative quantification of histone PTMs from C. elegans. We alkylate and tryptically digest the histones, analyze them by bottom‐up MS, and then evaluate the resulting data by a C. elegans−adapted version of the software EpiProfile 2.0. Finally, we show the utility of this pipeline by determining differences in histone PTMs between C. elegans strains that age at different rates and thereby achieve very different lifespans. © 2020 The Authors.
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 (τc ) 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.
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.
Characterization of the higher-order structure (HOS) of protein therapeutics, and in particular of monoclonal antibodies, by 2D 1 H-13 C methyl correlated NMR has been demonstrated as precise and robust. Such characterization can be greatly enhanced when collections of spectra are analyzed using multivariate approaches such as principal component analysis (PCA), allowing for the detection and identification of small structural differences in drug substance that may otherwise fall below the limit of detection of conventional spectral analysis. A major limitation to this approach is the presence of aliphatic signals from formulation or excipient components, which result in spectral interference with the protein signal of interest; however, the recently described Selective Excipient Reduction and Removal (SIERRA) filter greatly reduces this issue. Here we will outline how basic 2D 1 H-13 C methyl-correlated NMR may be combined with the SIERRA approach to collect 'clean' NMR spectra of formulated monoclonal antibody therapeutics (i.e., drug substance spectra free of interfering component signals), and how series of such spectra may be used for HOS characterization by direct PCA of the series spectral matrix. © 2020 U.S. Government. Basic Protocol 1: NMR data acquisition Basic Protocol 2: Full spectral matrix data processing and analysis Support Protocol: Data visualization and cluster analysis.
Heterologous expression of exogenous proteases in Escherichia coli often results in the formation of insoluble inclusion bodies. When sequestered into inclusion bodies, the functionality of the proteases is minimized. To be characterized structurally and functionally, however, proteases must be obtained in their native conformation. HIV protease is readily expressed as inclusion bodies, but must be recovered from the inclusion bodies. This protocol describes an efficient method for recovering HIV protease from inclusion bodies, as well as refolding and purifying the protein. HIV protease-containing inclusion bodies are treated with 8 M urea and purified via cation-exchange chromatography. Subsequent refolding by buffer exchange via dialysis and further purification by anion-exchange chromatography produces highly pure HIV protease that is functionally active. © 2020 by John Wiley & Sons, Inc. Basic Protocol: Recovery, refolding, and purification of HIV protease from inclusion bodies Support Protocol 1: Expression and extraction of inclusion bodies containing HIV protease expressed in Escherichia coli Support Protocol 2: Determination of the active site concentration of HIV protease via isothermal titration calorimetry.
Pichia pastoris is a eukaryotic microorganism reputed for its ability to mass-produce recombinant proteins, including integral membrane proteins, for various applications. This article details a series of protocols that progress towards the production of integral membrane proteins, their extraction and purification in the presence of detergents, and their eventual reconstitution in lipid nanoparticles. These basic procedures can be further optimized to provide integral membrane protein samples that are compatible with a number of structural and/or functional investigations at the molecular level. Each protocol provides general guidelines, technical hints, and specific recommendations, and is illustrated with case studies corresponding to several representative mammalian proteins. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Production of membrane proteins in a P. pastoris recombinant clone using methanol induction Basic Protocol 2: Preparation of whole-membrane fractions Alternate Protocol 1: Preparation of yeast protoplasts Basic Protocol 3: Extraction of membrane proteins from whole-membrane fractions Basic Protocol 4: Purification of membrane proteins Alternate Protocol 2: Purification of membrane proteins from yeast protoplasts Alternate Protocol 3: Simultaneous protoplast preparation and membrane solubilization for purification of membrane proteins Basic Protocol 5: Reconstitution of detergent-purified membrane proteins in lipid nanoparticles.
Lectin is a biomolecule that recognizes a specific part of glycans and, thus, has been used widely as a probe for glycoprotein analysis. Owing to the wide repertoire in nature combined with the recent two decades of advances in microarray technology, the multiplexed use of lectins has been widely used for glycan profiling of endogenous proteins. Because protein glycosylation is recognized as being biologically important and is expected to be a reliable disease marker, lectin microarray analysis with highly sensitive detection has been used to discover disease-relevant glycosylation alterations. However, the conventional system is limited to research purposes; thus, its implementation in clinical settings is warranted. Here, we provide an automatic glycan profiling method using GlycoBIST. A unique array format is used for 10-plexed lectin-glycoprotein interaction analysis on 1-mm-sized beads, which are arranged vertically in a capillary-shaped plastic tip. Using a one-boxed autopipetting machine, the whole process (including interaction, washing, and detection) is performed automatically and serially, resulting in reproducible measurements. In this article, a typical method for glycan profiling of a purified glycoprotein and the fabrication of GlycoBIST tips is explained. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Fabrication of a GlycoBIST tip Basic Protocol 2: Automatic profiling of a target glycoprotein using GlycoBIST.
