Articles in press

Obtaining homozygous mutant and transgenic lines is a critical yet time-consuming step in Arabidopsis thaliana research. Conventional breeding procedures require seeds to undergo complete maturation and natural desiccation before harvest, followed by cold stratification to overcome seed dormancy. This process substantially prolongs generation turnover and delays genetic screening. Here, we describe a rapid germination strategy based on the use of partially dehydrated seeds collected approximately 15 days after pollination (DAP). At this developmental stage, embryos have reached physiological maturity, while the seeds have not yet entered deep desiccation-induced dormancy. After surface sterilization and short-term cold treatment (2–3 days at 4 °C), these seeds readily germinate on 1/2 MS medium and develop into normal seedlings. By bypassing the prolonged maturation and dormancy phases associated with naturally dried seeds, this protocol shortens each generation cycle by approximately 1–2 weeks. The method is particularly useful for accelerating the propagation of transgenic materials and the identification of homozygous mutant lines in Arabidopsis.

Deubiquitinases (DUBs) are attractive therapeutic targets within the ubiquitin-proteasome system, in part because four of the five DUB subfamilies are cysteine proteases amenable to the development of potent, selective inhibitors, as recently demonstrated for USP7. Identifying DUBs that deubiquitylate and stabilize specific human oncogenic proteins is therefore a promising approach to discovering new mechanism-based targets for cancer therapy. Several complementary experimental strategies are typically required to identify bona fide DUB–substrate pairs. Here, we present an efficient, straightforward in vitro immune-complex protocol to validate USP17-mediated deubiquitylation of the transcriptional co-activator β-catenin. In this assay, both β-catenin and USP17 are immunopurified from transiently transfected 293T cells and then combined to assess USP17 enzymatic activity. The protocol describes the in vitro enzymatic assay performed on immunopurified complexes and the immunoblot-based readout. It can be readily adapted to other DUBs and substrates for mechanistic studies.

Neuroinflammation disrupts blood–brain barrier (BBB) integrity, promoting leukocyte recruitment into the central nervous system and contributing to the progression of neurological disorders. This protocol describes a reproducible macrophage adhesion assay to evaluate interactions between immune cells and brain endothelial cells and to screen compounds with potential anti-inflammatory activity. Murine brain endothelial cells (bEnd.3) were cultured to confluency and exposed to inflammatory stimuli, such as lipopolysaccharide (LPS) or pilocarpine, a cholinergic muscarinic receptor agonist reported to induce inflammatory responses through seizure-associated neuroinflammatory mechanisms, in the presence or absence of candidate therapeutic compounds. In these studies, the natural flavonoid quercetin and the synthetic alkyl-lysophospholipid edelfosine were tested for their effects on macrophage adhesion. After 48 h of treatment, fluorescently labeled murine macrophages (RAW 264.7) were added to the endothelial monolayer, and adherent cells were quantified by fluorescence microscopy. The assay was validated using dexamethasone as an anti-inflammatory control and inflammatory stimulation with LPS or pilocarpine. As expected, dexamethasone reduced macrophage adhesion, whereas both LPS and pilocarpine significantly increased adhesion, demonstrating the assay's sensitivity to changes in endothelial inflammatory status. Overall, this protocol provides a reliable and accessible platform for investigating endothelial–immune cell interactions under neuroinflammatory conditions and for evaluating therapeutic compounds that may preserve BBB function and reduce inflammatory cell recruitment in neurological disease models.

Whole-mount 3D imaging of multilayered biological tissues enables quantitative analysis of cell states and organization in their spatial context. However, extracting unbiased and meaningful quantitative information from dense, multilayered samples remains challenging due to imaging artifacts, increased density, and limited signal-to-noise ratio. Open source bioimage analysis workflows tailored to this type of analysis are scarce, and analysis bottlenecks like image curation or cell segmentation are seldom available without coding expertise. Here, we present a step-by-step computational protocol for the analysis of dense 3D organoid datasets using the Tapenade (Thorough Analysis PipEliNe for Advanced DEep imaging) workflow. Starting from multichannel image stacks, the protocol guides users through software installation, registration and fusion of multi-view datasets, preprocessing, and nuclei segmentation. It further details the generation of quantitative outputs, including morphometric measurements, deformation fields, and spatial correlation analyses. The workflow can be executed through open-source Python scripts or user-friendly Napari interfaces, allowing interactive parameter tuning and 3D visualization at each stage. This pipeline provides an accessible and modular framework for nonspecialist users to perform reproducible, multiscale quantitative analysis of 3D organoid images, while retaining flexibility for advanced users to customize individual steps.

Platelets originate from megakaryocytes, whose generation involves a series of biological processes including directed differentiation, proliferation, polyploidization, and maturation of hematopoietic stem cells. Abnormalities in megakaryocyte development and maturation can lead to quantitative and functional defects in platelets, thereby contributing to hemostatic or thrombotic disorders as well as the development of malignancies. Investigating megakaryocyte development and maturation and platelet production can provide important theoretical foundations for the diagnosis and treatment of thrombocytopenia, thrombotic diseases, and myeloproliferative neoplasms. Currently, there are three main clinical sources of hematopoietic stem cells (HSCs): bone marrow (BM), peripheral blood (PBSC), and umbilical cord blood (UCB). Among these, umbilical cord blood (UCB)-derived HSCs, due to their higher differentiation efficiency and stronger proliferative capacity, are the preferred starting cell source for studying megakaryocyte (MK) development and maturation and the mechanisms of platelet production. This article describes a detailed protocol covering all necessary steps for isolating CD34⁺ hematopoietic stem cells from umbilical cord blood, followed by in vitro induction culture with stem cell factor (SCF) and thrombopoietin (TPO) to generate mature megakaryocytes that highly express early megakaryocyte markers (CD41a, CD61) and late maturation markers (CD42a, CD42b). This protocol provides an effective tool for studying megakaryocyte development and platelet production and holds potential value for application in research on megakaryocyte-related diseases.

Memory is a fundamental process, regulated by protein–protein interactions within neuronal proteome networks. Learning-dependent changes in specific brain regions important for memory have been detected by mass spectrometry, by comparing proteins from animals trained to learn with mock-trained controls. Detection through this method relies on relative protein abundance; brain dissection is readily available for macroscopic animals to spatially control protein identification by mass spectrometry. In the nematode C. elegans, however, such spatial control is limited due to its microscopic size, hindering its utilization in proteomics. A protocol to address this limitation would strengthen an already excellent model to study memory, given that many proteins for learning are evolutionarily conserved in the worm and single-cell expression is uniquely defined across all 302 neurons. We modified existing protocols to enable (i) proximity labeling detection of neuronal proteins in C. elegans and (ii) high-throughput enrichment of these proteins from >3,000 whole worm bodies simultaneously, to assess trained vs. mock-trained proteomes. This involved the biotin ligase enzyme TurboID, which promiscuously labels nearby proteins with its substrate biotin. Enzyme expression was transgenically restricted to the nervous system, and biotin supplementation was limited to the training (or mock training) period in a classical (gustatory) conditioning paradigm. Labeled proteins were enriched by pull-down using streptavidin, which has a high binding affinity to biotin, and then processed for mass spectrometry runs and qualitative data analysis. This protocol is uniquely advantageous in that it minimizes proteins present before a temporal window of interest (training/mock training), improving the detection of lowly abundant proteins from a specific tissue in the worm (neurons). We have demonstrated that the protocol can sufficiently detect novel learning regulators, thus providing a useful framework to interrogate proteomes in microscopic brains.

The transient receptor mucolipin subtype 1 (TRPML1) is a ubiquitously expressed ion channel involved in lysosomal homeostasis. Recent pharmaceutical interest in developing agonist ligands has emerged due to beneficial effects in neurodegenerative diseases. The major high-throughput screening techniques to investigate this ion channel involve fluorescent calcium imaging and electrophysiology. Despite their high capacity for screening compounds, it is well known that both methods face hurdles, such as the need for expensive, specialized equipment. Here, we present a novel technique to screen for ligands of TRPML1 using a bioluminescence resonance energy transfer (BRET) assay. This assay consists of a target engagement assay in live cells, which permits the determination of binding constants between ligands and the target of interest in equilibrium or time-dependently. We employ a full-length TRPML1 C-terminally tagged with the small bioluminescent protein nanoluciferase. This ensures the correct localization of the ion channel in the lysosomal membrane and an optimal placement of the luciferase in the cytoplasm. We also developed a cell- and lysosome-permeable fluorescent BRET tracer that gives a BRET signal only when bound to the ion channel. This new protocol allows researchers worldwide to screen compounds that would interact with TRPML1 by using any plate reader with luminescent and fluorescence filters.

Multiple sclerosis (MS) is a chronic autoimmune disease characterized primarily by inflammatory demyelination of the central nervous system and is one of the leading causes of non-traumatic neurological disability in young and middle-aged adults worldwide. Myelin loss leads to impaired neural conduction, while progressive axonal degeneration resulting from failed remyelination constitutes a major pathological basis for irreversible disability in patients. Among currently approved treatments for MS, effective therapies that directly promote remyelination are still lacking; therefore, establishing animal models that can precisely recapitulate the myelin injury-repair process is essential for elucidating the mechanisms of remyelination and screening remyelination-promoting drugs. Focal demyelination models are important tools for investigating the mechanisms of remyelination and for developing therapeutic strategies for demyelinating diseases such as multiple sclerosis. Unlike the inflammation-driven injury of the experimental autoimmune encephalomyelitis (EAE) model and the systemic metabolic toxicity-induced demyelination of the cuprizone model, the lysophosphatidylcholine (LPC) injection model directly disrupts myelin in the corpus callosum through local injection of a membrane-solubilizing lipid, inducing focal demyelinating lesions and enabling investigators to study, in a controlled manner, the recruitment and differentiation of oligodendrocyte progenitor cells as well as the dynamic process of remyelination. This protocol describes the complete workflow for establishing focal demyelinating lesions by stereotaxic injection of LPC into the mouse corpus callosum, covering surgical preparation, coordinate localization, controlled injection, and postoperative care. Compared with existing methods, its main advantages lie in the precise control of the lesion and the synchronization of the post-injury repair phase, making it highly suitable for quantitative comparisons. Beyond the corpus callosum, this method is also broadly applicable to focal demyelination studies in other white matter tracts (including the spinal cord, optic nerve, and others), serving as a versatile platform for investigating region-specific myelin injury and repair.

Cytoplasmic protein aggregation is a defining feature of multiple neurodegenerative diseases, including amyotrophic lateral sclerosis, frontotemporal dementia, Huntington’s disease, and certain forms of motor neuron disease. Recent evidence indicates that promyelocytic leukemia protein (PML) and engineered PML-derived variants can act as versatile aggregate-remodeling factors. In particular, cytoplasmically redirected PML variants recognize pathological cytoplasmic inclusions and promote their clearance. Here, we describe a protocol to generate and validate two engineered cytoplasmic PML variants: full-length mPML, which is redirected to the cytoplasm by disruption of its nuclear localization sequence, and the truncated mPML^ΔRBC variant, which lacks the RING, B-box, and coiled-coil domain but retains aggregate-reducing activity. The protocol integrates fluorescence-based imaging, bimolecular fluorescence complementation, detergent-soluble/insoluble fractionation, and validation in primary rat cortical neurons. This workflow provides a practical platform for assessing cytoplasmic aggregate burden and for comparing the aggregate-remodeling activities of PML-derived constructs. It can also be adapted to other disease-associated aggregation-prone proteins, including TDP-43, SOD1, FUS, tau, polyGA, and polyQ-expanded proteins.

ADP ribosylation (ADPr) is a crucial post-translational modification that plays a vital role in DNA damage repair. Catalyzed by ADP ribose polymerases using NAD⁺ as a substrate, ADPr activates DNA repair pathways rapidly, thereby maintaining genomic integrity. The involvement of ADP ribose hydrolases in this process is significant, as they hydrolyze PAR chains, facilitating the release of ADPr-modified proteins from DNA or other proteins, which is essential for subsequent DNA repair steps. This protocol outlines a high-throughput screening method for identifying inhibitors of ADP ribose hydrolases, utilizing His-Tb-conjugated and ADPr-modified His-ADP ribose polymerase as the signal donor, and GST-d2-conjugated GST-XRCC1 as the signal receptor. The detection of time-resolved fluorescence signals enables efficient evaluation of compounds with potential therapeutic activity against cancer.

Immunohistochemistry (IHC) is a highly specific and widely used laboratory technique for assessing protein localization and expression in tissue samples. Interpretation of 3,3’ diamino benzidine (DAB)-based IHC is often based on observer-dependent manual scoring or traditional imaging software, which may show variability in DAB staining quantification. Furthermore, conventional image analysis tools often face limitations in precisely defining cell boundaries and quantifying membrane-specific signals. In this study, we present a standardized image analysis workflow using CellProfiler, an open-source software for image analysis for the quantification of membrane staining intensity in IHC images captured from slides prepared using formalin-fixed paraffin-embedded (FFPE) human cervical cancer tissue sections. The image analysis workflow was demonstrated using ASCT2 (SLC1A5), a membrane-localized amino acid transporter, as a representative biomarker for membrane-associated protein expression. This protocol involves image preprocessing, object identification, segmentation, and intensity measurement modules to distinguish cell membranes from cytoplasmic regions, enabling automated quantification of membrane intensity signals. The CellProfiler pipeline demonstrated improved accuracy in cell boundary identification and quantification of membrane-specific staining intensity. This is a rapid quantification process, since processing of each image only takes a few seconds; therefore, the analysis for 100 images can be performed within 10–15 min. This segmentation and quantification strategy is applicable to other membrane-based biomarkers after appropriate optimization of segmentation parameters. Following further minor modifications to the object identification modules, this pipeline can be used to detect and quantify cytoplasm- or nuclei-localized DAB-IHC markers across different tissue types. Overall, this protocol provides a standardized, user-friendly, and reproducible workflow for quantitative IHC image analysis that can be broadly applied to the study of protein biomarkers of different localizations, such as nuclei, cytoplasm, and cell membranes from different tissue types.

Nucleases are key tools in molecular biology, enabling controlled nucleic acid digestion for applications such as ribosome profiling. Micrococcal nuclease (MNase) from Staphylococcus aureus is widely used as a tool in molecular biology and biochemistry, but its reduced activity under high-salt conditions necessitates higher enzyme input to achieve efficient digestion, increasing costs in studies of halophilic organisms. Here, we present an optimized protocol for the heterologous expression and purification of the recombinant staphylococcal MNase. The procedure enables reproducible production of a highly active, stable enzyme and incorporates an enzymatic activity assay to standardize batches to minimize variability. The resulting MNase exhibits robust activity in high-salt environments and remains stable during storage, providing a cost-effective and reliable alternative to commercial nucleases for ribosome profiling and related applications.

Most existing preclinical models have been limited in their predictive value to mimic patients’ responses, which is a major drawback in drug development and the identification of predictive biomarkers. To overcome these limitations, patient-derived three-dimensional in vitro models have been proposed. One of them is the organoid model, which preserves the original cellular heterogeneity and recapitulates epithelial architecture and functionality. Recently, studies using patient-derived organoids for drug screening applications have increased in quantity, and organoids have already been applied to pancreatic, colon, and lung cancers and female gynecological malignancies. Here, we established a multiplex workflow to analyze longitudinal therapeutic effects of anti-cancer therapeutics on organoid growth, viability, and cytotoxicity by combining state-of-the-art viability measurement with automated live cell imaging. This workflow can be used for the prediction of patient-specific treatment response, high-throughput screening of potential anticancer drugs, and downstream analysis to identify novel therapeutic targets.