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.
Galectins are soluble carbohydrate binding proteins that can bind β‐galactose‐containing glycoconjugates by means of a conserved carbohydrate recognition domain (CRD). In mammalian systems, galectins have been shown to mediate very important roles in innate and adaptive immunity as well as facilitating host‐pathogen relationships. Many of these studies have relied on purified recombinant galectins to uncover key features of galectin biology. A major limitation to this approach is that certain recombinant galectins purified using standard protocols are easily susceptible to loss of glycan‐binding activity. As a result, biochemical studies that employ recombinant galectins can be misleading if the overall activity of a galectin remains unknown in a given assay condition. This article examines fundamental considerations when purifying galectins by lactosyl‐sepharose and nickel‐NTA affinity chromatography using human galectin‐4N and ‐7 as examples, respectively. As other approaches are also commonly applied to galectin purification, we also discuss alternative strategies to galectin purification, using human galectin‐1 and ‐9 as examples. © 2021 Wiley Periodicals LLC.
This article was corrected on 20 July 2022. See the end of the full text for details.
- NSF-PAR ID:
- Publisher / Repository:
- Wiley Blackwell (John Wiley & Sons)
- Date Published:
- Journal Name:
- Current Protocols
- Medium: X
- Sponsoring Org:
- National Science Foundation
More Like this
Abstract 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 colicell 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)
Class II major histocompatibility complex peptide (MHC‐IIp) multimers are precisely engineered reagents used to detect T cells specific for antigens from pathogens, tumors, and self‐proteins. While the related Class I MHC/peptide (MHC‐Ip) multimers are usually produced from subunits expressed in
E. coli, most Class II MHC alleles cannot be produced in bacteria, and this has contributed to the perception that MHC‐IIp reagents are harder to produce. Herein, we present a robust constitutive expression system for soluble biotinylated MHC‐IIp proteins that uses stable lentiviral vector−transduced derivatives of HEK‐293T cells. The expression design includes allele‐specific peptide ligands tethered to the amino‐terminus of the MHC‐II β chain via a protease‐cleavable linker. Following cleavage of the linker, HLA‐DM is used to catalyze efficient peptide exchange, enabling high‐throughput production of many distinct MHC‐IIp complexes from a single production cell line. Peptide exchange is monitored using either of two label‐free methods, native isoelectric focusing gel electrophoresis or matrix‐assisted laser desorption/ionization time‐of‐flight (MALDI‐TOF) mass spectrometry of eluted peptides. Together, these methods produce MHC‐IIp complexes that are highly homogeneous and that form the basis for excellent MHC‐IIp multimer reagents. © 2021 Wiley Periodicals LLC.
This article was corrected on 19 July 2022. See the end of the full text for details.
Basic Protocol 1: Lentivirus production and expression line creation Support Protocol 1: Six‐well assay for estimation of production cell line yield Support Protocol 2: Universal ELISA for quantifying proteins with fused leucine zippers and His‐tags Basic Protocol 2: Cultures for production of Class II MHC proteins Basic Protocol 3: Purification of Class II MHC proteins by anti‐leucine zipper affinity chromatography Alternate Protocol 1: IMAC purification of His‐tagged Class II MHC Support Protocol 3: Protein concentration measurements and adjustments Support Protocol 4: Polishing purification by anion‐exchange chromatography Support Protocol 5: Estimating biotinylation percentage by streptavidin precipitation Basic Protocol 4: Peptide exchange Basic Protocol 5: Analysis of peptide exchange by matrix‐assisted laser desorption/ionization (MALDI) mass spectrometry Alternate Protocol 2: Native isoelectric focusing to validate MHC‐II peptide loading Basic Protocol 6: Multimerization Basic Protocol 7: Staining cells with Class II MHC tetramers
Purification of recombinant proteins is a necessary step for functional or structural studies and other applications. Immobilized metal affinity chromatography is a common recombinant protein purification method. Mass spectrometry (MS) allows for confirmation of identity of expressed proteins and unambiguous detection of enzymatic substrates and reaction products. We demonstrate the detection of enzymes purified on immobilized metal affinity surfaces by direct or ambient ionization MS, and follow their enzymatic reactions by direct electrospray ionization (ESI) or desorption electrospray ionization (DESI).
A protein standard, His‐Ubq, and two recombinant proteins, His‐SHAN and His‐CS, expressed in
were immobilized on two immobilized metal affinity systems, Cu–nitriloacetic acid (Cu‐NTA) and Ni‐NTA. The proteins were purified on surface, and released in the ESI spray solvent for direct infusion, when using the 96‐well plate form factor, or analyzed directly from immobilized metal affinity‐coated microscope slides by DESI‐MS. Enzyme activity was followed by incubating the substrates in wells or by depositing substrate on immobilized protein on coated slides for analysis. Escherichia coli Results
Small proteins (His‐Ubq) and medium proteins (His‐SAHN) could readily be detected from 96‐well plates by direct infusion ESI, or from microscope slides by DESI‐MS after purification on surface from clarified
E. colicell lysate. Protein oxidation was observed for immobilized proteins on both Cu‐NTA and Ni‐NTA; however, this did not hamper the enzymatic reactions of these proteins. Both the nucleosidase reaction products for His‐SAHN and the methylation product of His‐CS (theobromine to caffeine) were detected. Conclusions
The immobilization, purification, release and detection of His‐tagged recombinant proteins using immobilized metal affinity surfaces for direct infusion ESI‐MS or ambient DESI‐MS analyses were successfully demonstrated. Recombinant proteins were purified to allow identification directly out of clarified cell lysate. Biological activities of the recombinant proteins were preserved allowing the investigation of enzymatic activity via MS.
Protein labeling strategies have been explored for decades to study protein structure, function, and regulation. Fluorescent labeling of a protein enables the study of protein‐protein interactions through biophysical methods such as microscale thermophoresis (MST). MST measures the directed motion of a fluorescently labeled protein in response to microscopic temperature gradients, and the protein's thermal mobility can be used to determine binding affinity. However, the stoichiometry and site specificity of fluorescent labeling are hard to control, and heterogeneous labeling can generate inaccuracies in binding measurements. Here, we describe an easy‐to‐apply protocol for high‐stoichiometric, site‐specific labeling of a protein at its N‐terminus with
N‐hydroxysuccinimide (NHS) esters as a means to measure protein‐protein interaction affinity by MST. This protocol includes guidelines for NHS ester labeling, fluorescent‐labeled protein purification, and MST measurement using a labeled protein. As an example of the entire workflow, we additionally provide a protocol for labeling a ubiquitin E3 enzyme and testing ubiquitin E2‐E3 enzyme binding affinity. These methods are highly adaptable and can be extended for protein interaction studies in various biological and biochemical circumstances. © 2021 Wiley Periodicals LLC.
This article was corrected on 18 July 2022. See the end of the full text for details.
Basic Protocol 1: Labeling a protein of interest at its N‐terminus with NHS esters through stepwise reaction Alternate Protocol: Labeling a protein of interest at its N‐terminus with NHS esters through a one‐pot reaction Basic Protocol 2: Purifying the N‐terminal fluorescent‐labeled protein and determining its concentration and labeling efficiency Basic Protocol 3: Using MST to determine the binding affinity of an N‐terminal fluorescent‐labeled protein to a binding partner. Basic Protocol 4: NHS ester labeling of ubiquitin E3 ligase WWP2 and measurement of the binding affinity between WWP2 and an E2 conjugating enzyme by the MST binding assay
Fungi infect over a billion people worldwide and contribute substantially to human morbidity and mortality despite all available therapies. New antifungal drugs are urgently needed. Decades of study have revealed numerous protein targets of potential therapeutic interest for which potent, fungal‐selective ligands remain to be discovered and developed. To measure the binding of diverse small molecule ligands to their larger protein targets, fluorescence polarization (FP) can provide a robust, inexpensive approach. The protocols in this article provide detailed guidance for developing FP‐based assays capable of measuring binding affinity in whole cell lysates without the need for purification of the target protein. Applications include screening of libraries to identify novel ligands and the definition of structure‐activity relationships to aid development of compounds with improved target affinity and fungal selectivity. © 2021 Wiley Periodicals LLC.
This article was corrected on 18 July 2022. See the end of the full text for details.
Basic Protocol 1: Use of saturation binding curves to optimize tracer and lysate protein concentrations Basic Protocol 2: Establishment of competition binding experiments Support Protocol 1: Preparation of fungal cell lysates Support Protocol 2: Preparation of human HepG2 cell lysate