Biochemistry

Immunopeptidomics enables the identification of peptides presented by major histocompatibility complex (MHC) molecules, offering insights into antigen presentation and immune recognition. Understanding these mechanisms in hypoxic conditions is crucial for deciphering immune responses within the tumor microenvironment. Current immunopeptidomics approaches do not capture hypoxia-induced changes in the repertoire of MHC-presented peptides. This protocol describes the isolation of MHC class I-bound peptides from in vitro hypoxia-treated cells, followed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis. It describes optimized steps for cell lysis, immunoaffinity purification, peptide elution, and MS-compatible preparation under controlled low-oxygen conditions. The method is compatible with various quantitative mass spectrometry approaches and can be adapted to different cell types. This workflow provides a reliable and reproducible approach to studying antigen presentation under hypoxic conditions, thereby enhancing physiological relevance and facilitating deeper immunological insights.

The antibody-uptake assay is a commonly used technique to monitor endocytosis of integral membrane proteins including transmembrane and glycosylphosphatidylinositol-anchored proteins (GPI-APs). The antibody-uptake assay typically involves incubating live cells with fluorophore-conjugated antibodies directed against the extracellular domain of the integral membrane protein of interest. Antibody uptake is then detected by flow cytometry or confocal microscopy. However, these detection modalities may be inaccessible to some labs or require extensive training to operate. Thus, we developed an easy and novel sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blot-based approach to the antibody-uptake assay that exploits the strong affinity between biotin and streptavidin. Instead of incubating cells with fluorophore-conjugated antibodies to monitor antibody uptake, our assay involves incubating cells with biotinylated antibodies, processing the cell lysates for western blot, and probing the membrane with detectably conjugated streptavidin. From preparation to quantification, this protocol requires less hands-on time than other approaches and is amenable to small-scale drug or siRNA screens. Here, we demonstrate the utility of our approach using the well-characterized misfolded GPI-AP, YFP-tagged C179A mutant of prion protein (YFP-PrP*), as our model substrate. YFP-PrP* constitutively traffics to the plasma membrane (PM), where it binds to anti-GFP antibody, and immediately undergoes endocytosis to lysosomes. To validate our protocol, we present measurements of antibody uptake under conditions known to enhance or inhibit YFP-PrP*’s traffic to the PM. Using this assay, we present new evidence that, under certain conditions, YFP-PrP* is able to undergo degradation via a pathway that does not involve exposure on the cell surface.

In neuropharmacology and drug development, in silico methods have become increasingly vital, particularly for studying receptor–ligand interactions at the molecular level. Membrane proteins such as GABA (A) receptors play a central role in neuronal signaling and are key targets for therapeutic intervention. While experimental techniques like electrophysiology and radioligand binding provide valuable functional data, they often fall short in resolving the structural complexity of membrane proteins and can be time-consuming, costly, and inaccessible in many research settings. This study presents a comprehensive computational workflow for investigating membrane protein–ligand interactions, demonstrated using the GABA (A) receptor α5β2γ2 subtype and mitragynine, an alkaloid from Mitragyna speciosa (Kratom), as a case study. The protocol includes homology modeling of the receptor based on a high-resolution template, followed by structure optimization and validation. Ligand docking is then used to predict binding sites and affinities at known modulatory interfaces. Finally, molecular dynamics (MD) simulations assess the stability and conformational dynamics of receptor–ligand complexes over time. Overall, this workflow offers a robust, reproducible approach for structural analysis of membrane protein–ligand interactions, supporting early-stage drug discovery and mechanistic studies across diverse membrane protein targets.

Insects rely on chemosensory proteins, including gustatory receptors, to detect chemical cues that regulate feeding, mating, and oviposition behaviours. Conventional approaches for studying these proteins are limited by the scarcity of experimentally resolved structures, especially in non-model pest species. Here, we present a reproducible computational protocol for the identification, functional annotation, and structural modelling of insect chemosensory proteins, demonstrated using gustatory receptors from the red palm weevil (Rhynchophorus ferrugineus) as an example. The protocol integrates publicly available sequence data with OmicsBox for functional annotation and ColabFold for high-confidence structure prediction, providing a step-by-step framework that can be applied to genome-derived or transcriptomic datasets. The workflow is designed for broad applicability across insect species and generates structurally reliable protein models suitable for downstream applications such as ligand docking or molecular dynamics simulations. By bridging functional annotation with structural characterisation, this protocol enables reproducible studies of chemosensory proteins in agricultural and ecological contexts and supports the development of novel pest management strategies.

Nowadays, recombinant proteins are the focus of various research fields, and their use ranges from therapeutic investigations to cellular model systems for the development of therapeutic approaches. Cell systems used for the expression of recombinant proteins should be comparable in terms of yield and expression efficiency. In many research fields, it is desirable to obtain high protein concentrations. A method that combines an easy workflow with rapid results and affordable costs remains missing, and a standardized approach to determining protein concentration in transgenic cell lines is essential for more reliable data analysis. Our protocol demonstrates the cluster fluorescence-linked immunosorbent assay (FLISA), a technique that allows the exact quantification of comparable protein expression amounts. Moreover, it enables the detection of clustered or bound subunits of a protein without necessitating ultracentrifugation. In the present protocol, we demonstrate the utilization of two transgene cell lines, each expressing distinct recombinant proteins, to provide comparability of protein yields and detectable subunit clustering.

Cellular phenomena such as signal integration and transmission are based on the correct spatiotemporal organization of biomolecules within the cell. Therefore, the targeted manipulation of such processes requires tools that can precisely induce the localizations and interactions of the key players relevant to these processes with high temporal resolution. Chemically induced dimerization (CID) techniques offer a powerful means to manipulate protein function with high temporal resolution and subcellular specificity, enabling direct control over cellular behavior. Here, we present the detailed synthesis and application of dual SLIPT and dual SLIPT^NVOC, which expand the SLIPT (self-localizing ligand-induced protein translocation) platform by incorporating a dual-ligand CID system. Dual SLIPT and dual SLIPT^NVOC independently sort into the inner leaflet of the plasma membrane via a lipid-like anchoring motif, where they present the two headgroup moieties trimethoprim (TMP) and HaloTag ligand (HTL), which recruit and dimerize any two ^iK6eDHFR- and HOB-tagged proteins of interest (POIs). Dual-SLIPT^NVOC furthermore enables this protein dimerization of POIs at the inner leaflet of the plasma membrane in a pre-determined order and light-controlled manner. In this protocol, we detail the synthetic strategy to access dual SLIPT and dual SLIPT^NVOC, while also providing the underlying rationale for key design and synthetic decisions, with the aim of offering a streamlined, accessible, and broadly implementable methodology. In addition to the detailed synthesis, we present representative applications and typical experimental outcomes and recommend strategies for data analysis to support effective use of the system. Notably, dual SLIPT and dual SLIPT^NVOC represent the first CID systems to emulate endogenous lipidation-driven membrane targeting, while retaining hallmark advantages of CID technology—the precision over POI identity, recruitment sequence, high spatiotemporal control, and “plug-and-play” flexibility.

Protein isolation combined with two-dimensional electrophoresis (2-DE) is a powerful technique for analyzing complex protein mixtures, enabling the simultaneous separation of thousands of proteins. This method involves two distinct steps: isoelectric focusing (IEF), which separates proteins based on their isoelectric points (pI), and sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), which separates proteins by their relative molecular weights. However, the success of 2-DE is highly dependent on the quality of the starting material. Isolating proteins from plant mature roots is challenging due to interfering compounds and a thick, lignin-rich cell wall. Bacterial proteins and metabolites further complicate extraction in legumes, which form symbiotic relationships with bacteria. Endogenous proteases can degrade proteins, and microbial contaminants may co-purify with plant proteins. Therefore, comparing extraction methods is essential to minimize contaminants, maximize yield, and preserve protein integrity. In this study, we compare two protein isolation techniques for lupine roots and optimize a protein precipitation protocol to enhance the yield for downstream proteomic analyses. The effectiveness of each method was evaluated based on the quality and resolution of 2-DE gel images. The optimized protocol provides a reliable platform for comparative proteomics and functional studies of lupine root responses to stress, e.g., drought or salinity, and symbiotic interactions with bacteria.

Artificial metalloenzymes (ArMs) hold great promise for expanding the toolbox of non-natural transformations usable in living systems, such as cells, plants, and animals. However, their practical application remains challenging, primarily due to their unsatisfactory stability and inefficient intracellular assembly. We recently reported a new strategy, called artificial metalloenzymes in artificial sanctuaries (ArMAS) through liquid–liquid phase separation (LLPS), to enhance the performance of ArMs in cells by placing them in more friendly artificial microenvironments. Here, this protocol describes the detailed method for using this ArMAS–LLPS strategy, a robust way to create artificial compartments using an ArM protein scaffold through LLPS and construct ArMs within using self-labeling cofactor anchoring reactions. In detail, in Escherichia coli, membraneless protein condensates are formed by expressing a self-labeling fusion protein, HaloTag-SNAPTag (HS) and act as intracellular sanctuaries. Simultaneously, the HS scaffolds enable site-specific, bioorthogonal conjugation with synthetic metal cofactors, facilitating efficient ArM formation within the LLPS domains. This strategy can significantly enhance the intracellular catalytic activity and stability of the named HS-based ArMs, allowing whole-cell catalysis to be performed to enable abiotic transformations both in vitro and in vivo. The protocol provides a proof-of-concept approach for researchers aiming to develop stable ArM-based whole-cell catalytic systems for synthetic biology and therapeutic applications.

Liquid–liquid phase separation (LLPS) underlies the spatial organization of the nucleolus, a membraneless organelle responsible for ribosomal RNA (rRNA) transcription and ribosome subunit assembly. One of the key proteins involved in the formation of the fibrillar center of the nucleolus is the treacle, an intrinsically disordered protein that contains low-complexity repeats enriched in charged amino acid residues. In this work, we present a detailed protocol for the bacterial expression and purification of a recombinant fragment of treacle comprising two tandem low-complexity repeat (LCR) modules, with a total length of 136 amino acids. This fragment is intended for subsequent in vitro investigation of its ability to undergo LLPS. The described method enables the production of a soluble, biochemically pure protein preparation suitable for studying the mechanisms of spontaneous condensate formation in a cell-free system. This approach allows for the controlled modeling and quantitative evaluation of the contribution of low-complexity sequences to the phase behavior of treacle, independently of its interactions with cellular partners in vivo.

N-glycosylation is a ubiquitous post-translational modification (PTM) that regulates protein folding, stability, and biological function. Accurate identification and validation of N-glycosylation are therefore critical for understanding how glycosylation modulates protein activity. Here, we present a robust workflow for analyzing protein N-glycosylation in both animal and plant systems using peptide-N⁴-(N-acetyl-β-glucosaminyl) asparagine-amidase A and F (PNGase A and PNGase F). After enzymatic cleavage of the asparagine-linked N-glycans, samples are analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and western blotting (WB) to detect shifts in apparent molecular weight (MW) indicative of deglycosylation. Key steps include denaturing the protein to expose glycosylation sites, optimizing buffer conditions for PNGase F and A treatment, and comparing glycosylated vs. deglycosylated forms by electrophoretic mobility. A troubleshooting guide addresses common challenges, including incomplete deglycosylation and low transfer efficiency during WB, offering practical solutions to ensure reliable results. This protocol provides researchers with a standardized, cost-effective framework for investigating protein N-glycosylation in diverse systems, from cell lysates to purified proteins, in both animal and plant models.