Biochemistry

Three-dimensional immunohistochemistry (3D-IHC) shows the organization of molecular assemblies in the context of tissue architecture. Deep and rapid antibody penetration into 3D tissues and highly sensitive detection are crucial for high-throughput analysis of 3D-IHC imaging. Here, we provide a detailed protocol for a nanobody (nAb)-based 3D-IHC technique, namely POD-nAb/FT-GO 3D-IHC, for high-speed and high-sensitivity detection of targets within 1-mm-thick mouse brain tissues. Peroxidase-fused nAb (POD-nAb) is a genetically encoded recombinant antibody, which consists of a camelid nAb and a variant of horseradish peroxidase, and fluorochromized tyramide-glucose oxidase (FT-GO) is a fluorescent tyramide signal amplification (TSA) system. POD-nAb/FT-GO 3D-IHC incorporates three main components: 1) tissue permeabilization, 2) POD-nAb binding, and 3) 3D-TSA reaction with FT-GO. POD-nAbs enhance signal penetration depth and allow for highly sensitive detection when combined with FT-GO signal amplification. By using the 3D-IHC protocol provided herein, we can visualize target molecules in mouse brain tissues of 1-mm thickness with drastic signal enhancement within three days. This protocol for POD-nAb/FT-GO 3D-IHC could facilitate structural and molecular interrogation of 3D tissues.

The activity of chloroplast ATP synthase (CF_oCF₁) is precisely regulated through a thioredoxin (Trx)-mediated dithiol/disulfide reaction in response to varying light conditions. This regulatory mechanism is further controlled by ΔpH formation across the thylakoid membrane. To better understand this complicating regulatory function of CF_oCF₁, a method is required to evaluate the extent of CF_oCF₁ reduction by Trx under controlled ΔpH conditions and to directly evaluate the redox state of CF_oCF₁. In this study, we present a simple in vitro procedure to assess the CF_oCF₁ reduction system using spinach thylakoids. The method consists of three key steps: (A) simple preparation of intact thylakoids from spinach leaves; (B) reduction of CF_oCF₁ on the thylakoid membrane using recombinant Trx under light irradiation; and (C) in situ determination of the redox state of CF_oCF₁ by labeling thiol groups with a maleimide reagent followed by protein detection using western blotting. The redox state of CF_oCF₁ was determined by mobility shifts on non-reducing SDS-PAGE. This protocol provides a refined strategy for elucidating the regulatory mechanism controlling energy conversion by CF_oCF₁ under fluctuating photosynthetic conditions.

Antibody therapeutics have demonstrated transformative impacts on improving the quality of life of millions of patients, whereas advances in antibody discovery technologies have imposed a significant production challenge for the generation of a large diversity of therapeutic antibody candidates. A demand for the rapid production of dozens of purified antibodies in 10-mg quantities is entailed for functional screening and molecular assessment studies. Here, we present a robust semi-automated production protocol that bridges the gap between miniaturized high-throughput screenings and conventional custom-scale workflows. This methodology and workflow utilize a simple high-titer transient Chinese hamster ovary (CHO) cell host–CHO4Tx^® expression system, a procedure of magnetic protein-A bead in-culture antibody capturing, and a semi-automated purification process with the GenScript AmMag^TM SA Plus system. This production protocol has been proven to be robust and valuable for the routine production of dozens of antibody constructs per week in sufficient quality and quantity for cell-based and biophysical studies.

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.