生物化学

De novo synthesis of purine nucleotide is essential for the production of genetic materials and cellular chemical energy. PRPP amidotransferase (PPAT) is the rate-limiting enzyme in de novo purine synthesis, thereby playing a crucial regulatory role in this pathway. Recent studies suggest that metabolic enzymes, including PPAT, form condensates through phase separation to regulate cellular metabolism in response to environmental changes. However, due to the lack of methods for purifying eukaryotic PPAT, the biophysical properties of the enzyme have remained unknown. Here, I describe a protocol for purifying budding yeast PPAT tagged with green fluorescent protein from yeast cells, as well as an in vitro assay to examine condensation of the fluorescent PPAT by microscopy. These techniques enabled us to elucidate the mechanism controlling PPAT condensation and may also be applicable to the purification and condensation assay of other enzymes.

Phosphatidylinositol 4,5-bisphosphate [PI(4,5)P₂] is a phospholipid enriched on the cytoplasmic leaflet of the plasma membrane, where it plays important roles in membrane trafficking and cytoskeletal dynamics through proteins that directly bind to it. PI(4,5)P₂ can be metabolized to other phosphorylated forms of phosphatidylinositol to regulate numerous processes such as cell growth and development. PI(4,5)P₂ can also be hydrolyzed to generate the second messengers diacylglycerol (DAG) and inositol triphosphate (IP₃). Altered metabolism or mislocalization of PI(4,5)P₂ can perturb one or more of its functions and contribute to disease states. Here, we present a protocol to visualize and quantify the localization of PI(4,5)P₂ in live cells. The protocol uses a highly specific PI(4,5)P₂ protein binding domain coupled to enhanced green fluorescence protein (PH-PLCD1-GFP), enabling localization and quantification of cytosol-facing PI(4,5)P₂ to be determined. Localization and quantification of the PH-PLCD1-GFP, PI(4,5)P₂ specific probe, is enabled by fluorescence imaging and confocal microscopy. This approach can be used to study the dynamics of PI(4,5)P₂ localization temporally in live cells under both physiological and pathological conditions.

We have observed that some proinsulin molecules in pancreatic islets and beta cell lines have incomplete or improper intramolecular disulfide bonds. These misfolded monomers can form intermolecular disulfide-linked complexes. Accurately quantifying the fraction of proinsulin molecules contained in these complexes is challenging. By proinsulin immunoblotting after nonreducing SDS-PAGE, the signal for disulfide-linked complexes can exceed the total proinsulin signal detected after reducing SDS-PAGE (i.e., overestimating the abundance of misfolded species due to antibody affinity differences). However, after modification of the SDS-PAGE and electrotransfer protocol, we have been able to more accurately estimate the fraction of proinsulin monomers folded to the native state, as well as misfolded proinsulin monomers and disulfide-linked complexes. Our improved technique offers the ability to obtain a more precise assessment of proinsulin misfolding under different environmental conditions in beta cells and normal islets and in diabetes.

Protein purification is a critical step in both life sciences and biomanufacturing. Traditional affinity chromatography (AC) methods, including His-tag-based purification, provide high-purity proteins but are limited by the high cost of resins and the need for additional tag-removal steps. In this protocol, we present a reusable SpyDock-modified epoxy resin coupled with a pH-inducible self-cleaving intein for direct purification of proteins with authentic N-termini. This method enables efficient protein purification from cell lysates, achieving high purity (>90%) and yields comparable to the His-tag approach, without requiring tag removal. The SpyDock-modified resin protocol is robust, easy to implement, and cost-effective, making it suitable for both research and large-scale industrial applications.

Expansion microscopy (ExM) is an imaging technique that enables super-resolution imaging of biological specimens using conventional confocal microscopy. This process entails the isotropic physical expansion of a (biomolecular) sample that has been cross-linked to a swellable polymer. The grafting of biomolecules (and the subsequent fluorescent readout) is accomplished by introducing an acryloyl group to the amine groups of lysine residues within the proteins, enabling subsequent imaging. However, visualizing actin filaments with high spatial resolution using ExM remains challenging. Herein, we report the construction of a phalloidin conjugate containing actin stains and their application in ExM. This protocol highlights the efficacy of trifunctional linker (TRITON/Actin-ExM) for F-actin imaging, demonstrating that TRITON-labeled actin allows for efficient anchoring and signal retention, enabling robust visualization of actin filaments in expansion microscopy.

Biomolecular condensates are macromolecular assemblies constituted of proteins that possess intrinsically disordered regions and RNA-binding ability together with nucleic acids. These compartments formed via liquid-liquid phase separation (LLPS) provide spatiotemporal control of crucial cellular processes such as RNA metabolism. The liquid-like state is dynamic and reversible, containing highly diffusible molecules, whereas gel, glass, and solid phases might not be reversible due to the strong intermolecular crosslinks. Neurodegeneration-associated proteins such as the prion protein (PrP) and Tau form liquid-like condensates that transition to gel- or solid-like structures upon genetic mutations and/or persistent cellular stress. Mounting evidence suggests that progression to a less dynamic state underlies the formation of neurotoxic aggregates. Understanding the dynamics of proteins and biomolecules in condensates by measuring their movement in different timescales is indispensable to characterize their material state and assess the kinetics of LLPS. Herein, we describe protein expression in E. coli and purification of full-length mouse recombinant PrP, our in vitro experimental system. Then, we describe a systematic method to analyze the dynamics of protein condensates by X-ray photon correlation spectroscopy (XPCS). We also present fluorescence recovery after photobleaching (FRAP)-optimized protocols to characterize condensates, including in cells. Next, we detail strategies for using fluorescence microscopy to give insights into the folding state of proteins in condensates. Phase-separated systems display non-equilibrium behavior with length scales ranging from nanometers to microns and timescales from microseconds to minutes. XPCS experiments provide unique insights into biomolecular dynamics and condensate fluidity. Using the combination of the three strategies detailed herein enables robust characterization of the biophysical properties and the nature of protein phase-separated states.

Membranes are very complex and dynamic structures that are essential for plant cellular functions and whose lipidic composition can be influenced by numerous factors. Anionic phospholipids, which include phosphatidylserine, phosphatidic acid, phosphatidylinositol, and phosphoinositides are key components of these membranes as they are involved in plant cell signaling and as even slight modifications in their quantities may largely impact the cell metabolism. However, the presence of these compounds in low amounts, as well as their poor stability during analysis by mass spectrometry, make their study very complicated. In addition, the precise quantification of all anionic phospholipid species is not possible by lipid separation using thin-layer chromatography followed by the analysis of their fatty acyl chains by gas chromatography. Here, we describe a straightforward strategy for the extraction and semi-quantification of all anionic phospholipid species from plant samples. Our method is based on the derivatization of the anionic phospholipids, and more especially on their methylation using trimethylsilyldiazomethane, followed by analysis by high-performance liquid chromatography coupled with a triple quadrupole mass spectrometer. This approach allows largely improving the sensitivity of the analysis of anionic phospholipids from plant samples, which will help to gain deeper insights into the functions and dynamics of these key parts of plant cellular signaling.

Xylan is the main component of hemicellulose and consists of a complex heteropolysaccharide with a heterogeneous structure. This framework, in addition to the crystalline structure of cellulosic fibers and the rigidity of lignin, makes lignocellulosic biomass (LCB) highly recalcitrant to degradation. Xylanases are glycoside hydrolases that cleave the β-1,4-glycoside linkages in the xylan backbone and have attracted increasing attention due to their potential uses in various industrial sectors such as pulp and paper, baking, pharmaceuticals, and lignocellulosic biorefining. For decades, the measurement of xylanase activity was based on reducing sugar quantification methods like DNS or Nelson/Somogyi assays, with numerous limitations in terms of specificity and interference from other enzymatic activities. A better alternative is the colorimetric Azo-Xylan assay, which specifically measures the endo-1,4-β-D-xylanase activity. In this study, the Azo-Xylan protocol was adapted from the company Megazyme to determine the enzymatic activity of thermostable xylanases produced by microbial consortia (i.e., microbiomes), aiming to determine biochemical features such as temperature and pH optima, thermostability, and shelf life. This modified approach offers a rapid, cost-effective, and highly specific method for the determination of xylanase activity in complex mixtures, helping the development of a xylanase-based method for the hydrolysis of hard-degrading substrates in bio-based industries.

Antibody purification is a fundamental technology for therapeutic and diagnostic applications. While traditional methods like ammonium sulfate precipitation and polyethylene glycol precipitation are cost-effective, they often result in low purity and require multiple purification steps. Protein A–based affinity chromatography, the gold standard for antibody purification, provides high specificity but can be further improved to increase loading capacity and reduce costs. In this protocol, we introduce a novel approach for purifying high-quality, high-purity antibodies from complex samples using SpyFixer/Z domain–modified resin. This method utilizes Spy chemistry for site-specific immobilization of the Z domain of Protein A, significantly enhancing antibody loading capacity up to 200 mg/mL resin and ensuring stable, durable immobilization. Using this protocol, we achieved >90% purity of human immunoglobulin G (hIgG) from diverse sources, including E. coli cell lysates, human serum, and industrial fermentation broth. The SpyFixer/Z domain–modified resin protocol is simple, cost-effective, and offers a robust, scalable solution for efficient antibody purification in bioengineering applications. This immobilization scheme based on Spy chemistry can also be extended to other immunoglobulin-binding proteins, such as Protein G and Protein L, to develop high-efficiency affinity resins.

Fluorescent protein biosensors (FPBs) that turn on—go from dark to bright upon binding their ligands—enable the detection of targets in living cells with high sensitivity and spatial localization. Several approaches exist for creating turn-on FPBs, most notably the method that gave rise to the GCaMP family of genetically encoded calcium indicators. However, it remains challenging to modify these sensors to recognize new ligands. We recently developed adaptable turn-on maturation (ATOM) biosensors, in which target recognition by a small binding domain triggers chromophore maturation in the fluorescent protein to which it is attached. ATOM sensors are advantageous because they are generalizable (by virtue of the monobody and nanobody binding domains) and modular (binding domains and fluorescent proteins of various colors can be mixed and matched for multiplexed imaging), capable of detecting endogenously expressed proteins, and able to function in subcellular compartments including the cytoplasm, nucleus, endoplasmic reticulum, and mitochondria. The protocols herein detail how to design, clone, and screen new ATOM sensors for detecting targets of choice. The starting materials are the genes encoding for a monobody or nanobody and for a cyan, yellow, or red fluorescent protein. We also present general guidelines for creating ATOM sensors using binding domains other than nanobodies and monobodies.