Biochemistry

The plant cell wall is a dynamic and complex extracellular matrix that not only provides structural integrity and determines cell shape but also mediates intercellular communication. Among its major components, pectins play essential roles in cell adhesion, wall porosity, hydration, and flexibility. Rhamnogalacturonan-I (RG-I), a structurally diverse pectic polysaccharide, remains one of the least understood components of the plant cell wall. Its backbone is substituted with arabinan, galactan, and arabinogalactan side chains that vary in length, branching, and composition across tissues, species, and developmental stages. In addition, RG-I can undergo modifications such as backbone acetylation, further contributing to its structural complexity and functional diversity. To advance understanding of RG-I, we present a detailed method for isolating RG-I from the model plant Arabidopsis thaliana. Leveraging Arabidopsis as a model system provides major advantages owing to its well-characterized genome and powerful molecular toolkit, enabling deeper investigation into the roles of RG-I in plant development and responses to environmental stress. Our method consists of two major steps: an initial chemical extraction using oxalate, followed by endo-polygalacturonase (EPG) digestion to fragment the pectic domains. An advantage of this approach is that it produces a dry material that can be stored at room temperature without special handling and does not introduce chemicals that may interfere with downstream analyses. The purified RG-I can be used for detailed compositional and structural analyses, as well as for functional studies of enzymes involved in pectin biosynthesis, modification, and degradation. Although this protocol was developed for isolating RG-I from Arabidopsis rosette leaves, it is also applicable to other Arabidopsis organs and other plant species.

Plasma membrane–associated condensates driven by liquid–liquid phase separation represent a novel mechanism of receptor-mediated signaling transduction, serving as mesoscale platforms that concentrate signaling molecules and modulate reaction kinetics. Condensate formation is a highly dynamic process that occurs within seconds to minutes following receptor activation. Here, we present methods for de novo reconstituting liquid-like condensates on supported lipid bilayers and assessing the condensate fluidity using fluorescence recovery after photobleaching (FRAP). This protocol encompasses supported lipid bilayer preparation, condensation imaging, and FRAP analysis using total internal reflection fluorescence (TIRF) microscopy. Supported lipid bilayers provide a membrane-mimicking environment for receptor signaling cascades, offering mechanistic insights into protein–protein and lipid–protein interactions amid micron-scale condensates. The protocol can also be adapted to study condensates associated with the internal membranes of the Golgi apparatus, mitochondria, and other organelles.

The CRISPR/Cas9 system is a cornerstone technology in genome editing. Delivery of pre-assembled Cas9 ribonucleoprotein (RNP) complexes exhibits distinct advantages, including reduced off-target effects and lower immunogenicity. Conventional methods for purifying Cas9 protein typically involve multi-step chromatography and the cleavage of fusion tag, which are time-consuming and result in diminished yields. In this study, we present a simplified, one-step purification strategy for functional Streptococcus pyogenes Cas9 (SpCas9) using the ubiquitin (Ub) fusion system in Escherichia coli. The N-terminal Ub fusion not only improves protein solubility but also facilitates high-yield production of the His-Ub-Cas9 fusion protein. Importantly, the Ub tag does not require proteolytic removal during purification, allowing direct one-step purification of the fusion protein via nickel-affinity chromatography. The purified His-Ub-Cas9 retains robust DNA cleavage activity in vivo, as validated in zebrafish embryos. This protocol greatly simplifies the production of functional Cas9 protein, facilitating its broad application in genome editing.

This article presents an efficient protocol for refolding recombinant proteins that are prone to aggregation and form inclusion bodies during expression in Escherichia coli. As a model system, the homolog of CRISPR-associated effector protein CasV-M was investigated. The key element of the developed approach is refolding directly on a metal-affinity Ni-TED (N,N,N´-tris(carboxymethyl)ethylendiamine) resin using a dual-gradient system: a stepwise reduction in the concentration of the chaotropic agent combined with a simultaneous increase in the concentration of a mild nonionic detergent. This combination ensures spatial separation of protein molecules, minimizes aggregation, and promotes the recovery of the native conformation. The resulting method appears to be an alternative to conventional refolding strategies, with potential improvements in the reproducibility and yield of soluble protein compared to dialysis or dilution. The proposed approach can be extended to a broad range of aggregation-prone proteins and is considered a promising strategy for obtaining otherwise insoluble recombinant proteins.

Although protein–protein interactions (PPIs) are central to nearly all biological processes, identifying and engineering high-affinity intracellular binders remains a significant challenge due to the complexity of the cellular environment and the folding constraints of proteins. Here, we present a two-stage complementary platform that combines magnetic-activated cell sorting (MACS)-based yeast surface display with functional ligand-binding identification by twin-arginine translocation (Tat)-based recognition of associating proteins (FLI-TRAP), a bacterial genetic selection system for efficient screening, validation, and optimization of PPIs. In the first stage, MACS-based yeast display enables the rapid high-throughput identification of candidate binders for a target antigen from a large synthetic-yeast display library through extracellular interaction screening. In the second stage, an antigen-focused library is subcloned into the FLI-TRAP system, which exploits the hitchhiker export process of the Escherichia coli Tat pathway to evaluate binder–antigen binding in the cytoplasm. This stage is achieved by co-expressing a Tat signal peptide–tagged protein of interest with a β-lactamase-tagged antigen target, such that only binder–antigen pairs with sufficient affinity are co-translocated into the periplasm, thus rendering the bacterium β-lactam antibiotic resistant. Because Tat-dependent export requires fully folded and soluble proteins, FLI-TRAP further serves as a stringent in vivo filter for intracellular compatibility, folding, and stability. Therefore, this approach provides a powerful and cost-effective pipeline for discovering and engineering intracellular protein binders with high affinity, specificity, and functional expression in bacterial systems. This workflow holds promise for several applications, including synthetic biology and screening of theragnostic proteins and PPI inhibitors.

Underwater noise is a growing source of anthropogenic pollution in aquatic environments. However, few studies have evaluated the impact of underwater noise on aquatic invertebrates. More importantly, studies involving early developmental stages have been poorly addressed. Significant limitations are due to the lack of standardized protocols for working in the laboratory. Particularly, the design of uniform procedures in the laboratory is important when working with species that inhabit short-term changing habitats, such as estuaries, which makes it difficult to carry out repeated experiments in the natural habitat. Besides, controlling for environmental variables is also important when assessing the effect of a stressor on the physiological parameters of individuals. This experimental protocol addresses that gap by offering an adaptable laboratory-based method to evaluate sublethal physiological responses to sound exposure under highly controlled conditions. Here, we present a reproducible and accessible laboratory protocol to expose crabs to recorded boat noise and evaluate physiological responses using oxidative stress biomarkers. The method is designed for ovigerous females, as we evaluated the effects on embryos and early life stages (i.e., larvae), but it can be readily adapted to different life stages of aquatic invertebrates. A key strength of this protocol is its simplicity and flexibility: animals are exposed to noise using submerged transducers under well-controlled laboratory conditions, ensuring consistency and repeatability. Following exposure, tissues or whole-body samples can be processed for a suite of oxidative stress biomarkers—glutathione-S-transferase (GST), catalase (CAT), lipid peroxidation (LPO), and protein oxidation. These biomarkers are highly responsive, cost-effective indicators that provide a sensitive and early readout of sublethal stress. Together, the exposure and analysis steps described in this protocol offer a powerful and scalable approach for investigating the physiological impacts of underwater noise in crustaceans and other aquatic invertebrates.

The protochlorophyllide (Pchlide) level is a crucial indicator of plant fitness. Precise quantification of Pchlide content is necessary not only in studies of flu-related mutants that over-accumulate Pchlide in the dark but also for research on plants suffering from environmental stresses. Due to its low content and interference of chlorophylls, quantitative determination of Pchlide content is a challenge. Here, we describe an optimized protocol for Pchlide extraction from Arabidopsis thaliana seedlings and subsequent analysis using high-performance liquid chromatography (HPLC) coupled with fluorescence detection. Divinyl-Protochlorophyllide (DV-Pchlide, the major form of Pchlide in plants) quantification is achieved by interpolating fluorescence peak areas against an experimentally derived standard curve. This protocol provides a reliable workflow for Pchlide quantification, facilitating the deciphering of the underlying mechanism of plant environmental resilience.

Small GTPases function as molecular switches in cells, and their activation triggers diverse cellular responses depending on the GTPase type. Therefore, visualizing small GTPase activation in living cells is crucial because their activity is tightly regulated in space and time, and this spatiotemporal pattern of activation often determines their specific cellular functions. Various biosensors, such as relocation-based sensors and fluorescence resonance energy transfer (FRET)-based sensors, have been developed. However, these methods rely on interactions between activated GTPases and their downstream effectors, which limits their applicability for detecting activation of GTPases with unknown or atypical effectors. Recently, we developed a novel method utilizing split fluorescence technology to detect membrane recruitment of small GTPases upon activation, designated the Small GTPase ActIvitY ANalyzing (SAIYAN) system. This approach offers a new strategy for monitoring small GTPase activation based on membrane association and is potentially applicable to a wide range of small GTPases, including those with uncharacterized effectors.

Traditional methods for studying protein–protein interactions often lack the resolution to quantitatively distinguish distinct oligomeric states, particularly for membrane proteins within their native lipid environments. To address this limitation, we developed SiMPull-POP (single-molecule pull-down polymeric nanodisc photobleaching), a single-molecule technique designed to quantify membrane protein oligomerization with high sensitivity and in a near-native context. The goal of SiMPull-POP is to enable precise, quantitative analysis of membrane protein assembly by preserving native lipid interactions using diisobutylene maleic acid (DIBMA) to form nanodiscs. Unlike ensemble methods such as co-immunoprecipitation or FRET, which average out heterogeneous populations, SiMPull-POP uses photobleaching to resolve monomeric, dimeric, and higher-order oligomeric states at the single-molecule level. We validated SiMPull-POP using several model systems. A truncated, single-pass transmembrane protein (Omp25) appeared primarily monomeric, while a membrane-tethered FKBP protein exhibited ligand-dependent dimerization upon addition of the AP ligand. Applying SiMPull-POP to EphA2, a receptor tyrosine kinase, we found it to be mostly monomeric in the absence of its ligand, Ephrin-A1, and shifting toward higher-order oligomers upon ligand binding. To explore factors influencing ligand-independent assembly, we modulated membrane cholesterol content. Reducing cholesterol induced spontaneous EphA2 oligomerization, indicating that cholesterol suppresses receptor self-association. Overall, SiMPull-POP offers significant advantages over conventional techniques by enabling quantitative, single-molecule resolution of membrane protein complexes in a native-like environment. This approach provides critical insights into how membrane properties and external stimuli regulate protein assembly, supporting broader efforts to understand membrane protein function in both normal and disease states.

The cellular secretome is a rich source of biomarkers and extracellular signaling molecules, but proteomic profiling remains challenging, especially when processing culture volumes greater than 5 mL. Low protein abundance, high serum contamination, and sample loss during preparation limit reproducibility and sensitivity in mass spectrometry–based workflows. Here, we present an optimized and scalable protocol that integrates (i) 50 kDa molecular weight cutoff ultrafiltration, (ii) spin column depletion of abundant serum proteins, and (iii) acetone/TCA precipitation for protein recovery. This workflow enables balanced recovery of both low- and high-molecular-weight proteins while reducing background from serum albumin, thereby improving sensitivity, reproducibility, and dynamic range for LC–MS/MS analysis. Validated in human mesenchymal stromal cell cultures, the protocol is broadly applicable across diverse cell types and experimental designs, making it well-suited for biomarker discovery and extracellular proteomics.