生物化学

Peptergent is a novel class of amphipathic peptides that enables detergent-free extraction of membrane proteins (MPs) from lipid bilayers. This reagent self-assembles around hydrophobic transmembrane regions, forming stable, water-soluble complexes that can be isolated directly from biological membranes. Peptergent therefore bypasses the limitations imposed by traditional detergents, which often destabilize protein assemblies. Since detergents are completely avoided, MPs are directly amenable to structural and mass spectrometry (MS) analysis, thereby addressing their persistent underrepresentation in proteomic datasets and improving their accessibility in drug-screening strategies. We present here a streamlined protocol for MPs extraction with the Peptergent PDET-1, followed by exchange into His-tagged Peptidiscs for Ni-NTA-based affinity purification. The method encompasses membrane isolation, peptide preparation, protein extraction, clarification, and MPs exchange from Peptergents to Peptidiscs. This workflow yields an enriched membrane proteome compatible with downstream LC-MS/MS analysis for improved identification of multi-pass MPs.

The conventional kinesin-1 is a plus-end-directed microtubule-dependent motor protein with distinct motor head, stalk, and tail domains. Along with the motor head, which binds and walks along microtubules in an adenosine 5’-triphosphate (ATP) dependent manner, kinesin also contains a C-terminal microtubule binding tail. Motor-driven collective motility is well characterized using in vitro gliding assays, which show uninterrupted, smooth trajectories of transport. However, gliding assays driven by the full-length Drosophila kinesin-1 with both head and tail resulted in the emergence of spontaneous spatial microtubule patterns and stop-and-go motion. This was reproduced by an equimolar ratio of the active head and passive tail. Here, we describe the detailed protocol to reconstitute these microtubule gliding assays using multiple motor types: the full-length kinesin-1, the motor head or tail, mixtures of both head and tail, and a rigor mutant of the kinesin. We provide details of the approach taken to acquire the image time-series, to then quantify the spatial patterns that result from these motor combinations. Our approach provides a framework to systematically characterize the spatiotemporal effects of molecular motor-driven collective microtubule transport.

Peptidoglycan (PG), a network of glycan strands crosslinked by short peptides, is an essential and bacterial-specific structure that determines cell shape and protects cells from lysis. Understanding how bacteria assemble, maintain, and modify their PG not only addresses fundamental questions in cell biology but also provides a basis for developing strategies to treat bacterial infections. Although several in vitro methods, such as zymography, Remazol Brilliant Blue (RBB) assay, and LC-MS analyses, are available to quantify the activities of PG-modification enzymes, these approaches are not readily applicable in vivo. Here, we describe a single-particle tracking photo-activated localization microscopy (sptPALM)-based method to quantify the binding of enzymes to PG in vivo, which serves as a proxy for their enzymatic activities. Because the PG meshwork is relatively immobile, fluorescently tagged enzymes that transiently or stably bind it exhibit reduced mobility, reflected by lower diffusion coefficients. This approach provides sensitive, quantitative, and real-time insights into enzyme behavior in vivo under diverse physiological conditions or genetic backgrounds. The protocol is particularly valuable for investigating PG-modification enzymes that are essential or functionally redundant, which are often difficult to analyze using traditional genetic methods.

α-Synuclein (α-syn) aggregation has emerged as a key pathogenetic feature in several neurodegenerative disorders. The α-syn protein has various conformational strains, each with unique structural features that influence their cytotoxicity, propagation, and neuroinflammation. A post-translational modification known as O-GlcNAcylation has been found to influence the toxicity of α-syn and its propensity to aggregate. Difficulties in detecting and quantifying this modification are a major challenge to understanding its roles among the conformational forms of α-syn. We now describe a protocol for detecting O-GlcNAcylated α-syn that combines a click chemistry labeling approach and western blotting. This chemoenzymatic method involves the transfer of azido-modified galactose (GalNAz) from UDP-GalNAz to O-GlcNAcylated proteins, enabling their further functionalization with alkyne-containing polyethylene glycol of defined molecular weight. This protocol facilitates the determination of the glycosylation status of varying conformations of α-syn and their stoichiometric ratios.

Cellular function relies on a network of precisely regulated interactions among macromolecules such as proteins, peptides, carbohydrates, and nucleic acids. These molecular interactions regulate vital processes, including signaling, structural organization, and developmental patterning. Biolayer interferometry (BLI) is a label-free optical biosensing technique that enables real-time quantification of such interactions. This protocol describes how to use BLI to assess the binding affinity between a biotinylated plant peptide hormone (RALF1) and cell wall–derived oligogalacturonides (OG25–50) on the Octet RED96 platform. Streptavidin-coated biosensors are employed to immobilize the ligand, while analyte binding is monitored through wavelength shifts in the reflected light. The protocol includes detailed steps for sensor preparation, assay setup, software configuration, and kinetic data analysis. While optimized for plant peptide–matrix interactions, the method is broadly adaptable to other macromolecular systems across biological disciplines.

Unsaturated fatty acids (UFAs) play key roles in essential cellular functions such as membrane dynamics, metabolism, and animal development. Disruptions in UFA metabolism are linked to metabolic, cardiovascular, and neurodegenerative disorders. Cellular UFAs composition and quantification are normally determined using methods such as gas chromatography and/or mass spectrometry, which require extraction procedures and prevent analysis of live specimens. Here, we describe a protocol that employs uniform ¹³C isotope labeling and high-resolution 2D solution-state nuclear magnetic resonance (NMR) spectroscopy to analyze lipid composition and fatty acid unsaturation directly in the model organism Caenorhabditis elegans. The approach enables in vivo assessment of lipid storage compositions with sufficient resolution and sensitivity to distinguish wild-type animals from those with altered fatty acid desaturation. Complementary analysis of total lipid extracts provides information regarding lipid molecules that are not detected in vivo, such as phospholipid molecules organized in biological membranes. Overall, this non-destructive NMR-based method offers a powerful tool for investigating lipid metabolism in C. elegans and other small model systems that can be isotopically enriched.

Nanobodies are recombinant single-domain antibodies (VHHs) derived from the heavy chain–only subset of camelid immunoglobulins that can be reverse-engineered into bivalent antibodies by fusion to immunoglobulin Fc constant regions. Mammalian cells are the system of choice to produce VHH-Fcs to ensure authentic folding and post-translation glycosylation of the expressed VHH-Fcs. In a recent project to find neutralising VHH-Fc binders to the spike proteins of SARS-CoV-2 viruses, we identified a need for rapid expression and purification of multiple VHH-Fc fusions from nanobodies selected by phage display. Here, we present a protocol for the construction of expression vectors by parallel ligase-independent cloning, transient small-scale expression in mammalian cells (4 mL culture volume), screening antigen-binding activity, and midi-scale purification (30 mL culture volume) for downstream activity assays. The workflow is completely transferable between different vector formats, of which three are described herein: Fc fusion dimers, monomeric CD4 fusions, and His-tagged monomers.

Bottom-up proteomics workflows encompass several key stages, including sample preparation, data acquisition, and data analysis. Of these, sample preparation is the initial and critical stage, as it significantly influences the depth, reproducibility, and reliability of subsequent mass spectrometry–based analyses. While several main digestion strategies exist, including in-gel, in-solution, and filter-aided methods, each presents distinct trade-offs in terms of throughput, contamination removal, and applicability to complex biological matrices. The Suspension Trapping (S-Trap) method offers a compelling alternative by efficiently capturing and digesting proteins while removing interferents like sodium dodecyl sulfate (SDS), which can compromise downstream LC–MS/MS performance. This protocol details a S-Trap workflow optimized for biofluid proteomics, specifically plasma, serum, and cerebrospinal fluid (CSF). We describe two complementary formats: a manual tube-based procedure for individual or small-batch samples and a 96-well-plate-based system enabling high-throughput processing. The protocol integrates optional high-abundance protein depletion to enhance coverage of low-abundance analytes and includes steps for reduction, alkylation, digestion, and peptide elution for low total protein content samples, such as plasma, serum, and cerebrospinal fluid. By providing a detailed protocol, this work aims to improve the consistency and accessibility of S-Trap-based sample preparation, facilitating robust and reproducible discoveries in bottom-up proteomics.

Manganese (Mn) is an essential trace element whose intracellular homeostasis is tightly controlled by specialized membrane transporters. Dysregulation of Mn transport leads to pathological Mn accumulation and severe human disease; however, efficient and quantitative cell-based methods for assessing Mn²⁺ transporter activity remain limited. Here, we present an optimized cellular Fura-2 manganese extraction assay (CFMEA) that enables robust quantification of cellular Mn content and provides a normalized framework for assessing relative Mn²⁺ transport activity in a high-throughput format. This protocol integrates Fura-2-based fluorescence detection of Mn²⁺ at the Ca²⁺ isosbestic excitation wavelength with dsDNA quantification to normalize dsDNA levels in cell extracts and immunoblotting to account for transporter protein expression levels. Cells expressing Mn²⁺ transporters are exposed to MnCl₂ in 96-well plates, washed to remove extracellular Mn²⁺, and lysed in a Fura-2-containing extraction buffer. Fluorescence quenched by Mn²⁺ is quantified and converted to cellular Mn content using a cell-free Mn-Fura-2 standard curve and then normalized to dsDNA content and protein abundance to determine relative transporter activity. This workflow provides a relatively sensitive, reproducible, and low-cost approach for comparative analysis of Mn²⁺ transporters and their variants across multiple cell types. The protocol is demonstrated using the Mn²⁺ efflux transporter SLC30A10 in HEK293T cells and is readily adaptable for studying other Mn²⁺ transport pathways.

Membrane transporters mediate the selective movement of ions and molecules across biological membranes and are essential for cellular homeostasis. However, their functional characterization in living cells is often complicated by the complexity of the native membrane environment. Reconstitution into model membrane systems provides a powerful alternative by enabling precise control over lipid composition and experimental conditions. Giant unilamellar vesicles (GUVs) are particularly well suited for transporter studies, as their cell-sized dimensions allow direct microscopic observation and fluorescence-based measurements of protein activity. Here, we describe a two-step reconstitution protocol in which transport proteins are first incorporated into large unilamellar vesicles and then used to generate protein-containing giant unilamellar vesicles (proteo-GUVs) via the poly(vinyl alcohol) swelling method. This two-step approach enhances protein incorporation efficiency and preserves transporter functionality. The method is exemplified using the P3-type ATPase Arabidopsis thaliana plasma membrane H⁺-ATPase isoform 2 (AHA2). We further describe a fluorescence-based assay to assess proton transport activity in proteo-GUVs. Our approach provides a versatile and controlled platform for biochemical, biophysical, and single-molecule analysis of membrane transporters.