Biochemistry

Divalent metal ion transporters are conserved across all domains of life and play essential roles in diverse processes such as manganese acquisition during nutritional immunity in bacteria and iron homeostasis in higher eukaryotes [1–3]. Traditional techniques, such as electrophysiological assays, are often unsuitable due to the slow kinetics of many membrane transporters, electroneutral nature of certain transporter types, and the influence of other proteins with similar activity. To overcome these limitations and to investigate both the activity and ion selectivity of transporters, also including those normally expressed intracellularly, we have developed a fluorescence-based transport assay using purified proteins. This in vitro assay uses encapsulated fluorophores to monitor the movement of divalent metal ions (e.g., Mn²⁺, Ca²⁺, Mg²⁺) or protons across liposomal membranes reconstituted with purified transporter proteins. This approach provides detailed functional insight that complements structural and cellular data.

3-nitro-tyrosine (nitroTyr) is one of numerous oxidative protein modifications implicated in diseases such as cardiovascular disease, cancer, and amyotrophic lateral sclerosis (ALS). Because of this, the ability to site-specifically encode nitroTyr into recombinant proteins is a powerful approach for studying these disease pathways. However, producing proteins with defined nitration sites is technically challenging due to the limitations of traditional chemical nitration via peroxynitrite, which lacks residue and site-specificity. Genetic code expansion (GCE) offers a solution by enabling precise incorporation of nitroTyr at designated TAG codons using engineered aminoacyl-tRNA synthetase/tRNA pairs from Methanocaldococcus jannaschii and Methanomethylophilus alvus. This protocol provides a reliable, optimized workflow for incorporating nitroTyr into proteins in E. coli using GCE. It guides users through key considerations in selecting cell lines, media conditions, and GCE systems to minimize off-target effects such as release factor 1 competition, near-cognate suppression, and chemical reduction of nitroTyr. The method is demonstrated using wild-type and TAG-containing superfolder GFP but is broadly applicable to other proteins of interest.

Reactive oxygen species (ROS) are central regulators of plant development and stress responses, with hydrogen peroxide (H₂O₂) acting as a key signaling molecule whose spatial distribution determines adaptive versus damaging outcomes. Accurate detection of H₂O₂ at tissue and cellular resolution is therefore essential for understanding redox-dependent regulation of plant growth. A variety of techniques have been used to monitor H₂O₂, including bulk spectrophotometric and fluorometric assays, genetically encoded sensors for real-time measurements, and chemical probes for in situ detection. While these approaches differ in sensitivity, specificity, and temporal resolution, many are limited by a lack of spatial information, technical complexity, or dependence on transgenic material. Here, we present a detailed protocol for 3,3′-diaminobenzidine (DAB)-based histochemical detection of H₂O₂ in seedling roots, covering staining, imaging, and semi-quantitative image analysis using open-source software (FIJI/ImageJ). The method relies on peroxidase-mediated oxidation of DAB, resulting in a stable, light-resistant, and insoluble precipitate that enables visualization of H₂O₂ accumulation with high spatial resolution. This protocol provides a robust, accessible, and genetically independent approach for spatial analysis of H₂O₂ in plant tissues. Its simplicity, compatibility with diverse genotypes and treatments, and suitability for semi-quantitative analysis make it a valuable tool for examining the spatial distribution of H₂O₂, thereby providing spatial insight into redox-related regulatory processes during plant development and stress responses.

The spatiotemporal dynamics and density of actin networks are key determinants of actin cytoskeleton–mediated cellular functions. In vitro reconstitution systems have been widely used to study actin cytoskeletal dynamics; however, many existing approaches offer limited flexibility in controlling the geometry, thickness, and density of the assembled actin networks. Here, we present an in vitro optogenetic protocol that enables precise control of actin network assembly on supported lipid bilayers using an improved light-induced dimer (iLID)-SspB-based light-inducible dimerization system. In this system, His-mEGFP-iLID is anchored to a Ni-NTA-containing lipid bilayer, while SspB-mScarlet-I-VCA, a nucleation-promoting factor fused with SspB, together with other actin cytoskeletal proteins, is supplied in bulk solution. Upon blue light illumination, SspB-mScarlet-I-VCA is recruited to the membrane in a spatially and temporally defined manner, inducing localized actin polymerization. By tuning illumination patterns and duration, actin networks with defined density, thickness, and geometry can be generated, and polymerization can be rapidly halted by stopping illumination. This protocol provides a versatile platform for reconstructing actin networks with controlled spatial organization and density, enabling quantitative analysis of density-dependent interactions between actin networks and actin-binding proteins.

Among the biophysical techniques used in fragment-based drug discovery (FBDD) campaigns, crystallography is the most sensitive, allowing for the identification of low-affinity ligands and the characterization of protein–ligand complexes at atomic resolution. Although powerful, the proper application of this technique depends on obtaining crystals capable of diffracting X-rays at high resolution. Additionally, in crystallographic compound screening, the crystals must be resistant to multiple organic solvents used in chemical libraries, such as DMSO. In this protocol, we describe recombinant protein production suitable for crystallization and procedures for X-ray crystallographic screening of a library of 768 fragments. As a case study, we used the Schistosoma mansoni thioredoxin glutathione reductase (SmTGR), a redox enzyme with a key role in controlling oxidative stress in parasites of the Schistosoma genus, which causes schistosomiasis. As a validated pharmacological target, SmTGR is used in the development of new schistosomicidal drugs. The experimental pipeline includes SmTGR expression, purification, and crystallization, crystal soaking, diffraction data collection, and refinement. The 768 fragments from the Diamond-SGC Poised Library (DSPL) were individually soaked onto the crystals, and diffraction data were collected and processed at the I04-1 beamline of the Diamond Light Source synchrotron. Diffraction data were subsequently analyzed using PanDDA to identify fragment-binding events and to enable reliable detection of low-occupancy ligands within the protein crystal structures. In addition to the core experimental steps, this protocol incorporates systematic approaches to overcome limitations frequently encountered in crystallographic screening campaigns, including assessment of crystal solvent tolerance, acceleration of crystal mounting through the use of auxiliary devices, acoustic dispensing–based soaking of hundreds of fragments for low material consumption and high throughput, automated data collection, and efficient data analysis pipeline for the detection of weakly bound ligand. This protocol can be broadly applied to screen diverse compound sets against multiple targets amenable to crystallization.

Cyclic GMP–AMP synthase (cGAS) is a key cytosolic double-stranded DNA sensor that activates innate immune responses. Upon binding double-stranded DNA, cGAS undergoes conformational activation and catalyzes the synthesis of the second messenger 2′3′-cyclic GMP–AMP (2′3′-cGAMP) from ATP and GTP. 2′3′-cGAMP then triggers a downstream signaling cascade that induces type-I interferon and inflammatory gene expression and has been shown to exert antitumor effects in the context of cancer. Accurate measurement of this enzymatic activity is therefore important for mechanistic studies. Traditional kinetic methods such as radiolabeling, HPLC, or mass spectrometry provide precise results but require specialized equipment and expertise. Here, we describe a rapid and accessible ELISA-based protocol to quantify 2′3′-cGAMP product formation and derive cGAS enzymatic parameters. Reactions are initiated with defined DNA ligands and quenched at multiple time points, and product accumulation is quantified by a commercially available 2′3′-cGAMP ELISA. Time course measurements are used to calculate initial velocities, which can be plotted against substrate concentration to obtain Michaelis–Menten parameters. This approach enables direct, product-specific quantification of 2′3′-cGAMP formation using only an absorbance plate reader. The protocol provides a sensitive and broadly applicable alternative to traditional methods, allowing laboratories without advanced instrumentation to perform reliable cGAS enzyme kinetics.

Organic solvent–based tissue clearing methods are widely used for whole-brain imaging but often compromise endogenous fluorescence. Existing protocols, such as iDISCO and fluorescence-preserving variants, have improved optical transparency but still present trade-offs between fluorescence retention, tissue stability, and workflow complexity. Here, we present MDISCO, a modified iDISCO-based clearing protocol designed to enhance preservation of endogenous fluorescence while maintaining high transparency and stable tissue morphology. MDISCO is directly compared with FDISCO+, an established fluorescence-preserving protocol, for the preservation of endogenous tdTomato and YFP. Performance across clearing steps is evaluated by measuring brain weight, anteroposterior and mediolateral dimensions, and optical transparency before and after solvent clearing and refractive index matching. Fluorescence preservation is assessed using whole-brain light-sheet microscopy with standardized imaging parameters to enable direct comparison. This protocol provides an accessible and high-throughput, reproducible workflow for solvent-based clearing with robust endogenous fluorescence preservation, offering clear advantages for whole-brain 3D imaging of genetically encoded fluorescent reporters.

Small ubiquitin-related modifiers (SUMOs) are covalently conjugated onto the proteome and serve as signaling molecules in many aspects of eukaryotic cell biology, from S. cerevisiae and C. elegans to H. sapiens. The conjugatable SUMO variants, SUMO1 and the almost identical SUMO2 and SUMO3 (designated SUMO2/3), are processed by an E1(SAE1:SAE2)-E2(UBC9)-E3 enzyme cascade to produce SUMO-modified proteins. The prerogative of the SUMO biology field is to identify and study the specific proteins undergoing SUMOylation, which grants us insights into the biological pathway of interest. This protocol was developed using the human osteosarcoma cell line U2OS to enable the investigation of SUMO conjugates in mitosis, the cell division phase of the cell cycle. We enrich the cell population for mitotic cells, which are isolated and subjected to stringent lysis conditions involving a high concentration of SDS and DTT in RIPA buffer, to promote complete protein denaturation. The lysates in high SDS RIPA buffer are diluted to reduce the overall SDS concentration and undergo conventional immunoprecipitation using SUMO1- or SUMO2/3-specific antibodies bound to protein A/G agarose beads. The samples are then compatible with downstream readouts such as western blots and mass spectrometry. This protocol detects endogenous SUMOylated proteins and avoids exogenous SUMO overexpression, which can alter SUMO conjugate formation. Furthermore, this denaturing protocol ensures only SUMOylated proteins are immunoprecipitated, and not their interactors.

Laccase2 (Lac2), a member of the phenoloxidase (PO) family, is an essential oxidase for melanin pigmentation in insects. The identification of the in vivo spatial distribution of Lac2 is crucial for understanding the molecular mechanisms underlying color pattern formation. However, it is technically difficult to determine the distribution because Lac2 expression peaks at late pupal stages, when adult cuticle sclerotization has already begun. Here, we report a simple and rapid protocol for estimating the distribution of endogenous PO proteins, prophenoloxidases (proPOs) and phenoloxidases (POs), in insect tissues. In this method, the spatial distribution of endogenous PO proteins is estimated based on staining patterns formed by dopamine melanin synthesis in tissues incubated in a solution containing isopropanol and dopamine. We validated that tissues collected at approximately 80% of the total pupal duration yielded staining patterns corresponding to adult melanin-forming regions in three insect species. By comparing staining patterns across developmental stages, this protocol enables estimation of the timing of color pattern formation. Furthermore, the contrast between stained and unstained regions within the same tissue allows region-specific sampling, thereby facilitating an investigation of the underlying molecular mechanisms regulating spatial PO distribution. Taken together, this method facilitates the study of melanin biosynthesis and enables the identification of the genes involved in regulating color pattern formation. This protocol does not require antibodies, transgenic lines, or specialized equipment and can be completed within a short time frame. Its effectiveness has been validated in multiple coleopteran and lepidopteran species, demonstrating its broad applicability as a versatile tool for studying insect pigmentation and color pattern formation.

The genome of the nematode Caenorhabditis elegans encodes at least 160 predicted peptide precursor genes that can generate over 300 bioactive peptides, the functions of most of which remain unknown. Phenotypes resulting from deletion or transgenic expression of peptide genes are readily assayed, but genetic dissection of individual peptide activities is often confounded when a single gene encodes multiple peptides or when distinct peptides act redundantly. Here, we describe a protocol for direct microinjection of chemically synthesized peptides into individual worms. This approach permits investigation of the effects of an individual peptide while providing precise temporal control over peptide delivery.