Biochemistry

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.

Extracellular vesicles (EVs) are nanoscale particles secreted by all cells and present in all biological fluids, where they carry molecular cargo reflective of health and disease states. Their diagnostic potential is often obscured by the high abundance of non-EV proteins and lipoproteins (e.g., albumin, apolipoproteins) that complicate proteomic analysis of primary biofluids, such as ascites fluid. Conventional isolation strategies face a persistent trade-off between EV purity and yield. To overcome this, a magnetic bead-based protocol (Mag-Net) to enrich EVs according to electrochemical surface charge using strong anion-exchange chemistry (SAX) was adapted for proteomics. Our workflow is specifically adapted to ascites fluid from human or murine sources. This approach effectively separates EVs from high-abundance proteins and lipoproteins, enabling proteomic profiling from as little as 2 μL of ascites fluid. Demonstrated in both murine and human ovarian cancer models, Mag-Net offers a reproducible, scalable, and automation-ready solution for EV isolation from various biofluids.

Transcription factors (TFs) regulate gene expression by binding to cis-regulatory elements in the genome. Understanding transcriptional regulation requires genome-wide characterization of TF occupancy across different chromatin contexts, yet simultaneous assessment of TF binding for multiple factors remains technically challenging. Here, we describe a detailed and reproducible protocol for cFOOT-seq, a cytosine deaminase–based genomic footprinting assay by sequencing, which enables antibody-independent, base-resolution profiling of chromatin accessibility, nucleosome organization, and TF occupancy. In cFOOT-seq, the double-stranded DNA (dsDNA) cytosine deaminase SsdA_tox converts cytosine to uracil in accessible chromatin, whereas TF binding and nucleosome occupancy locally protect DNA from deamination. Using the FootTrack analysis framework, deamination patterns generated by cFOOT-seq are quantitatively analyzed to derive standardized footprint and chromatin organization profiles at base resolution across the genome. Because cFOOT-seq preserves genomic DNA integrity during deamination-based footprinting, it is compatible with ATAC-seq-based chromatin enrichment. ATAC-combined implementations reduce sequencing depth requirements and improve scalability for footprint-focused analyses, supporting applications in low-input and single-cell settings. This protocol provides a practical framework for genome-wide TF footprint profiling and can be readily applied to dissect gene regulatory mechanisms in development, immunity, and disease, including cancer.

Anthocyanins are specialized flavonoid pigments that play critical roles in plant coloration, photoprotection, and responses to environmental stress. Arabidopsis thaliana serves as a valuable genetic model for dissecting anthocyanin biosynthesis and regulatory networks. Conventional methods for anthocyanin quantification, such as crude spectrophotometric assays, often compromise pigment integrity, yield inconsistent results, and provide limited information on compound composition. Here, we describe a simple, reproducible, and high-fidelity protocol for the induction, extraction, quantification, and chromatographic profiling of anthocyanins in Arabidopsis thaliana seedlings. The workflow employs well-defined anthocyanin-inductive conditions (AIC), methanol/formic acid extraction, lyophilization for dry-weight normalization, and dual quantification via spectrophotometry and High-performance liquid chromatography with diode-array detection (HPLC-DAD) analysis. This protocol enables accurate comparison between wild-type and mutant genotypes, facilitating both mutant screening and metabolic pathway analysis. The approach minimizes pigment degradation, enhances reproducibility across replicates, and offers a robust tool for research in plant metabolism, stress physiology, and flavonoid biochemistry.

The deletion and mutation of Topoisomerase 3β (TOP3B) is linked to multiple neurological disorders and is the only known topoisomerase that is also catalytically active on RNA in vitro and in cells. Uniquely, TOP3B is primarily localized to the cytoplasm, binds to open reading frames of mRNA, and regulates mRNA stability and translation in a transcript-specific manner. A common approach for studying TOP3B activity in cells is immunodetection of TOP3B•RNA covalent intermediates after bulk RNA isolation. However, in this approach, the RNA species is unknown and is not selective for the major TOP3B substrate, mRNA. In this protocol, we describe a recently developed and optimized protocol for capturing TOP3B•mRNA covalent intermediates using oligo-dT isolation of mRNA under protein-denaturing conditions. Covalent intermediates are then detected by a dual membrane slot blotting strategy with nitrocellulose and positively charged nylon membranes. Nitrocellulose membrane-bound TOP3B•mRNA covalent intermediates are analyzed by immunodetection, and nylon membrane-bound free mRNA is stained with methylene blue. The protocol detailed below has been validated with wildtype and mutant 3xFLAG-tagged TOP3B expressed in Neuro2A cells, with additional optimization for slot blotting using recombinant EGFP.

ADGRL4 is an adhesion G protein–coupled receptor (aGPCR) implicated in multiple tumours. In our experience, conventional insect cell-based baculovirus expression systems have not yielded sufficient correctly folded ADGRL4 protein for purification and cryo-electron microscopy (cryo-EM) analysis. Here, we describe aGPCR-HEK, a six-week protocol that establishes stable tetracycline-inducible mammalian HEK293S GnTI- TetR cell lines expressing N-terminally HA- and GFP-tagged aGPCRs. The method comprises lentiviral production in Lenti-X 293T cells, transduction of target adherent HEK293S GnTI- TetR cells, flow cytometry enrichment of uninduced GFP-positive cells displaying leaky expression, adaptation to suspension culture, and large-scale tetracycline induction and harvesting of cells for downstream purification and cryo-EM. The system yields reproducible, milligram-scale quantities of folded aGPCR suitable for structural and biochemical studies.