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While advances in omics technologies have greatly improved our understanding of cellular heterogeneity, there is an increasing need for complementary approaches that capture the spatial organization and structural dynamics of cells. Image-based single-cell phenotypic profiling provides quantitative information on cell morphology and organelle organization, offering valuable insights into cellular function and regulation. Although numerous image analysis tools are available, establishing a complete analysis workflow, from image preprocessing and segmentation to feature extraction and multivariate analysis, often requires substantial computational expertise and software integration. Here, we describe a Python-based workflow for image-based single-cell phenotypic profiling from immunofluorescence microscopy images and provide a detailed protocol for its implementation. Using synchronized HeLa cells with drug-induced mitotic spindle defects as an example, the workflow covers image loading, cell segmentation, quantitative feature extraction, profile integration, dimensionality reduction, clustering, and data visualization. The protocol is accompanied by example datasets, annotated Jupyter Notebooks, and instructions for execution in either a local Python environment or Google Colab, facilitating straightforward implementation and customization. By integrating the entire analysis pipeline within a single coding environment, this workflow enables reproducible and accessible single-cell morphological profiling without requiring specialized imaging equipment or extensive programming expertise. The workflow therefore provides a practical platform for studying cell morphology, organelle organization, and cellular dynamics across a broad range of biological applications.

The release and updating of whole-genome sequences of several representatives of the human opportunistic pathogen Pseudomonas aeruginosa have laid the ground for investigating the mechanisms of antibiotic resistance, biofilm formation, and virulence, while also offering opportunities for researchers to find new therapeutic targets to control this bacterium using a functional genomics approach. However, there is still a lack of detailed protocols describing gene inactivation methods in P. aeruginosa, resulting in failures and extra time spent designing in-house protocols. Here, we introduce a rapid, efficient, and unmarked deletion mutagenesis method combining overlap extension PCR, efficient conjugation, and the traditionally used sacB-based counter-selection procedure. Efficient generation of deletion mutants using this detailed protocol can be easily completed in one week using standard lab reagents. Importantly, this method may be adaptable to other bacteria where the sacB-based counter-selection system works.

Zebrafish is an excellent in vivo model for high-throughput antiangiogenic drug testing, commonly known as the zebrafish angiogenesis assay. Conventional zebrafish angiogenesis assays are performed in wild-type zebrafish to evaluate the vascular changes in intersegmental and subintestinal vessel regions of the zebrafish larvae at 2 and 3 dpf stages, respectively. However, wild-type zebrafish larvae do not adequately mimic the hypoxia microenvironment and ectopic vessel branching characteristics of cancer. To overcome this limitation, we developed a genetically engineered zebrafish model with constitutive activation of the hypoxia signaling pathway by targeting the vhl, a tumor suppressor, by negative regulation of the hypoxia pathway, using CRISPR mutagenesis. This model exhibits robust ectopic blood vessel branching throughout the larval body, thereby recapitulating pathological angiogenesis. The utility of this zebrafish model system for drug screening was validated with the sorafenib treatment, a known antiangiogenic tyrosine kinase inhibitor. Overall, this proangiogenic hypoxia zebrafish model provides a physiologically relevant platform for testing antiangiogenic drugs.

Plants growing in soil are exposed to several environmental stresses, such as soil compaction, flooding, or drought, and need to quickly acclimate to these conditions to survive. Such acclimations include suberization and lignification of the outer and inner parts of the root. Visualization of suberin and lignin depositions in roots can be done with several well-established staining techniques followed by microscopy, but these methods are mostly qualitative. Radial water loss assays provide a quick, robust, and quantitative method to evaluate the formation of suberized and/or lignified outer apoplastic barriers and how tightly they regulate water loss. Here, we provide a detailed step-by-step protocol, from harvesting the roots from soil conditions and cleaning them, to preparing them for water loss measurements, while using basic lab equipment. Since the data obtained is water loss as mass, quantitative and statistical comparisons can be performed. Finally, while in this protocol roots were grown in soil, the method can be applied to roots growing in hydroponics, agar plates, or any other growth conditions that allow non-destructive harvesting of roots. In summary, the protocol provides a fast and reliable method to assess differences in outer apoplastic barrier development of root tissues, allowing for fast root tissue screenings while obtaining quantitative data without expensive or highly specialized equipment.

Extracellular vesicle (EV) proteomics can be used to study intercellular communication and find biomarkers of non-invasive diseases. Traditional separation methods (ultracentrifugation, size exclusion chromatography) and data-dependent acquisition (DDA) mass spectrometry usually have the drawbacks of copurification of pollutants, poor reproducibility, and insufficient sampling of low-abundance peptides. This protocol describes a workflow for label-free extracellular vesicle proteomics, which combines size exclusion chromatography for vesicle separation, data-independent acquisition (DIA) for deep discovery, and parallel reaction monitoring (PRM) for targeted verification. Plasma-derived extracellular vesicles are treated by standardized lysis, digestion, and LC-MS/MS procedures, so that the protein group of extracellular vesicles can be identified and quantified robustly. The main advantages of this scheme are that it can achieve high quantitative reproducibility, unbiased detection of low-intensity peptides, and seamless transition from discovery to targeted verification, while also being suitable for liquid biopsy samples and various cardiovascular diseases.

Spatial transcriptomics enables genome-wide gene expression profiling while preserving tissue architecture, making it a powerful approach for studying plant developmental transitions. However, preparing small and structurally complex plant tissues for spatial transcriptomics remains technically challenging because samples must be rapidly preserved, precisely oriented, serially sectioned, and accurately positioned within the limited capture area of the Visium slides. Here, we describe an optimized workflow for cryo-embedding, serial cryosectioning, section placement, and data analysis of small plant samples for 10x Genomics Visium spatial transcriptomics. Using maize seedling shoot apices as the target, this protocol includes preparation of custom molds for optimal cutting temperature embedding, rapid fresh sample embedding, serial cryosectioning, section-position marking for Visium HD workflows, and morphological quality assessment of replicate tissue slides before transcript capture. The associated data analysis workflow includes Space Ranger processing, Seurat-based normalization and Harmony integration, anatomical domain annotation, pseudobulk and developmental trend analyses, RNA velocity, pseudotime analysis, transcription factor network analysis, single-cell reference mapping, and 3D transcriptome reconstruction. This computational workflow was developed and tested using maize Visium V1 data, but not Visium HD data. This protocol was used to generate serial spatial transcriptomes of maize shoot apices and developing leaf primordia, enabling reconstruction of gene expression transitions from the shoot apical meristem to sequential leaf developmental stages. The approach is also applicable to other small plant tissues, including Arabidopsis first true leaves and Marchantia thalli.

Region-specific RNA sequencing is a powerful approach for investigating tissue differentiation and dynamic changes in gene expression during embryonic development. The chicken embryo has long served as an important model system in developmental biology. However, the limited availability of tissue-specific reporter lines makes region-specific RNA-seq approaches particularly valuable in this organism. Here, we applied photo-isolation chemistry-based RNA sequencing (PIC-RNA-seq) to the somatic lateral plate mesoderm (sLPM) before the emergence of limb bud progenitor cells (LPCs) and to early LPCs in chicken embryos. These analyses revealed the upregulation of multiple genes, including Hox genes, in LPCs, suggesting the initiation of their regional patterning program. This workflow enables visualization of dynamic changes in the gene expression profile of LPCs and should also be applicable to other tissues in avian embryos.

Leptospirosis is a widespread zoonotic disease caused by pathogenic bacteria of the genus Leptospira. The isolation and comprehensive characterization of circulating strains within a region are essential for understanding the local epidemiology and improving public health surveillance. Historically, whole blood has been the specimen of choice for isolation; however, its efficiency can be limited by the presence of inhibitory substances in the sample, and Leptospira viability may depend on rapid processing and inoculation. Here, we present an in-house culture protocol for the isolation of Leptospira from serum samples previously maintained under refrigeration (i.e., 4–8 °C) for up to 10 days. The protocol employs a real-time PCR-guided strategy by first screening specimens for the lipL32 gene. Positive samples are then inoculated into specialized EMJH media supplemented with AFAS and EMJH+AFAS supplemented with STAFF antibiotic cocktail, followed by incubation at 30 °C for up to six months. Growth is monitored weekly through visual inspection and, once turbid, the presence of Leptospira is determined via dark-field microscopy prior to downstream serogroup and genomic characterization. A significant advantage of this method is the successful recovery of viable Leptospira from non-fresh serum specimens stored under refrigeration, even in samples with low bacterial loads. Additionally, the protocol facilitates broader surveillance by repurposing serum samples already collected for routine serology, increasing the probability of identifying diverse strains without further clinical collection

Cellular stress induces profound changes in cytoskeletal organization and biomolecular condensate formation. Traditional approaches for monitoring cellular stress often require multi-component biosensors, endpoint staining procedures, or indirect biochemical measurements. Here, we describe a protocol for the use of OptoProfilin, a genetically encoded single-component optogenetic biosensor derived from Profilin-1 fused to Cryptochrome 2 (Cry2) and mCherry. Following transient expression in mammalian cells, OptoProfilin exhibits light-dependent localization to focal adhesions under non-stressed conditions and transitions to punctate condensates under energetic, oxidative, osmotic, and senescence-associated stress conditions. The protocol includes transient transfection, induction of cellular stress, live-cell imaging, immunofluorescence validation, and quantitative image analysis. While this protocol describes imaging on a Leica widefield fluorescence microscope, it can readily be extended to other microscopy platforms. As a stand-alone biosensor that produces visually distinct responses in stressed versus non-stressed cells, OptoProfilin provides a convenient platform for investigating stress-associated cytoskeletal remodeling and biomolecular condensate formation.

Human bone marrow organoids provide a tractable three-dimensional platform for modeling hematopoiesis and hematologic disease in a human niche–like context. Here, we describe a stepwise protocol for utilizing human induced pluripotent stem cell (iPSC)-derived bone marrow organoids that support autonomous hematopoiesis for hematopoietic disease modeling, mouse xenograft hematopoiesis, and drug sensitivity testing. The workflow combines embryoid body formation, early mesoderm/angiogenic induction under hypoxia, hemogenic endothelial commitment, maturation within a collagen-containing hydrogel, and subsequent suspension culture as individual organoids. The resulting organoids contain endothelial, stromal, and hematopoietic components and reproduce key structural and cellular features of human marrow. We further describe procedures for engraftment of normal donor- or patient-derived CD34+ cells and implantation of mature organoids under the renal capsule of immunodeficient mice to assess in vivo hematopoietic maintenance. In prior applications of this platform, donor-derived CD34+ cells were shown to engraft within the organoid niche and undergo multilineage differentiation, enabling detection of selective erythroid defects caused by DDX41 deficiency and assessment of therapeutic suppression of JAK2V617F-mutant patient-derived hematopoietic cells in a human marrow–like microenvironment. This protocol, therefore, enables disease modeling, in vivo xenograft assessment, and ex vivo functional analysis of patient-derived hematopoietic cells using relatively small input samples.

Xenopus laevis oocytes are widely used as a heterologous expression system for investigating the function of membrane proteins due to robust expression of heterologous protein and a low endogenous transport background. Traditionally, transporter activity in oocytes has been assessed using electrophysiology or radiolabeled uptake assays, approaches that are constrained by the requirement for electrogenicity of the transport process, availability of radiolabeled compounds, and instrumentation. Here, we describe a fluorescence-based uptake assay that enables direct and rapid quantification of transporter activity using a fluorescence plate reader. The protocol uses the Arabidopsis thaliana sucrose transporter 1 (SUC1) and its fluorescent substrate esculin as a case for how to set up the assay. The workflow includes optimizing assay conditions, sample preparation, fluorescent measurements, and downstream data analysis using R. This method can readily be adapted to other transporter-substrate pairs, and it supports applications such as transporter inhibitor screening, mutational analysis, characterization of kinetic properties, or indirect substrate specificity testing through competition assays. Overall, this protocol provides a simple and scalable alternative to traditional techniques, eliminating the need for radiolabeled compounds or electrophysiology while enabling easy quantitative assessment of transporter activity.

Human induced pluripotent stem cell (hiPSC)-derived liver organoids have emerged as valuable models for studying human liver development. However, existing organoid systems often lack developmentally matched cell populations, particularly fetal hepatic stellate cells (HSCs), limiting their ability to recapitulate key developmental processes. Current approaches for generating HSCs rely on primary cells, immortalized cell lines, or hiPSC differentiation methods that frequently produce activated HSC-like cells and often require cell sorting. Here, we describe an efficient protocol for generating expandable fetal-like HSCs from hiPSCs through a stepwise differentiation strategy that mimics embryonic HSC development. The resulting cells can be robustly expanded while maintaining characteristic molecular and functional features of fetal HSCs. This protocol provides a reproducible and scalable source of fetal-like HSCs without cell sorting and supports the generation of multicellular liver organoids containing developmentally relevant stromal components. Beyond the validation of the protocol in studies of liver maturation and vascularization, it can be applied to investigations of HSC biology and congenital liver diseases.

This protocol describes a targeted workflow for identifying and quantitatively comparing kinase autophosphorylation sites, using in-gel proteolytic digestion, LC-MS/MS analysis, phosphosite localisation, and extracted ion chromatogram (XIC) analysis. The detailed protocol begins with an excised SDS-PAGE gel band containing the immunoprecipitated kinase of interest. Standard cell culture, expression, immunoprecipitation, and SDS-PAGE procedures can be used to generate this starting material according to the experimental system under investigation; the specific conditions used for NEK1 are described in our associated research article. Here, the protocol is applied to characterise phosphorylation sites on NEK1—a serine/threonine kinase implicated in amyotrophic lateral sclerosis (ALS)—using a homozygous NEK1 knockout cell background in which GFP-tagged wild-type or kinase-dead NEK1 constructs are re-expressed. This experimental system enables accurate assessment of kinase-dependent phosphorylation events in the absence of endogenous NEK1. A key methodological feature of the workflow is the use of two parallel MS2 fragmentation strategies: higher energy collisional dissociation (HCD) and electron-transfer/HCD (EThcD). EThcD generates complementary b/y and c/z ion series, providing superior localisation confidence for labile phosphorylation events that are difficult to resolve using HCD alone. Relative phosphopeptide abundance across conditions can be assessed using extracted ion chromatograms generated in FreeStyle, or using Skyline software with normalisation to the total ion current (TIC), as performed in the associated NEK1 study. Although demonstrated here using NEK1, the protocol is readily adaptable to other kinases amenable to immunoprecipitation and will be of interest to researchers studying kinase signalling, post-translational modification biology, and disease-associated phosphorylation events.

Multicellular organization relies on reciprocal interactions between molecular events, such as gene expression and protein state, and higher-scale properties, such as spatial patterning and tissue architecture. Understanding these processes requires methods that enable quantitative measurements at subcellular resolution, while maintaining the three-dimensional tissue organization. Conventional immunofluorescence imaging captures spatial information but is limited to the number of fluorescence markers that can be imaged simultaneously, whereas dissociation-based single-cell approaches can profile multimodal cellular states but lack positional information. Here, we describe 3D in toto iterative immunofluorescence imaging, termed 3D-4i, which enables up to ten-plex protein and protein state measurements in early zebrafish embryos. Leveraging sample immobilization on 96-well plates together with a gentle liquid handling system and high-content spinning disc confocal microscopy, this method comprises repeated rounds of antibody staining, optical clearing, confocal imaging, and antibody elution. Subsequent image analysis allows segmentation of nuclei and cells, extraction of quantitative single-cell features, and integration of molecular measurements with spatial context. Altogether, 3D-4i provides a scalable platform for investigating diverse biological processes in intact embryos, while maintaining both subcellular resolution and three-dimensional context.

Reliable DNA extraction is essential for genetic research on marine species; however, obtaining sufficient DNA from single fish eggs remains challenging. Existing protocols often require optimization to achieve high PCR efficiency. The optimized phenol–chloroform–isoamyl extraction protocol presented in this paper improves DNA yield and quality from individual eggs of Atlantic bluefin tuna (Thunnus thynnus), bogue (Boops bops), saddled seabream (Oblada melanura), and painted comber (Serranus scriba) by modifying buffer volumes, incubation times, and washing steps, following prior micropuncturing of eggs on a glass slide. DNA quality is confirmed by spectrophotometry, PCR amplification of the mitochondrial COI gene, electrophoresis, and Sanger sequencing. This method provides a low-cost and effective approach for species identification from individual fish eggs.

Spatial organization of macromolecules is fundamental to cellular function, with colocalization providing key insights into molecular interactions and biological processes. However, quantification remains challenging due to diverse localization patterns and irregular sample geometries. Here, we present a protocol for analyzing colocalization between two fluorescent probes using coAnalyzer, a MATLAB-based software package. coAnalyzer features a user-friendly graphical interface and supports region of interest (ROI) selection, image merging, line scanning, signal isolation, scatterplot generation, and quantitative analysis. coAnalyzer enables colocalization analysis of any two fluorophores, regardless of the proteins or dyes involved. Its broad applicability across a wide range of organisms and sample types demonstrates the robustness, flexibility, and versatility of the platform.

Molecular tension sensors enable the visualization of forces acting on specific intracellular proteins in living cells. Most established genetically encoded sensors rely on Förster resonance energy transfer (FRET), requiring donor–acceptor imaging and correction for spectral bleed-through, which can complicate their application in heterogeneous tissues. We developed non-FRET α-actinin and α-catenin tension indicators containing a force-responsive tension sensor (TS) module composed of an optimized circularly permuted enhanced green fluorescent protein (cpEGFP) scaffold and an elastic (GPGGA)₈ linker. The TS module is incorporated into α-actinin or α-catenin, while a C-terminal mCherry serves as a force-insensitive reference for indicator abundance. This protocol describes indicator expression in cultured cells, validation using the myosin II inhibitor blebbistatin, two-color time-lapse imaging and live-cell super-resolution imaging using SRRF-Stream reconstruction based on super-resolution radial fluctuations (SRRF), and calculation of the green/red fluorescence ratio and normalized relaxation ratio. The protocol also specifies essential quality-control procedures, including imaging under nonsaturating conditions with fixed channel-specific acquisition settings across comparisons, registration of the green and red channels, and the use of appropriate vehicle and negative controls. For SRRF-Stream analysis, the green and red channels are reconstructed separately using identical reconstruction settings before ratio calculation. Representative reconstructed images should be compared with the corresponding conventional images to check for reconstruction artifacts. Optional procedures describe imaging of isolated cardiomyocytes and freshly isolated organs from tension-indicator mice. Because molecular loading is inferred from changes in TS fluorescence normalized to mCherry rather than from energy transfer between two fluorophores, the method avoids FRET-specific bleed-through correction; nevertheless, controlled acquisition and channel registration remain necessary. The indicators report relative changes in molecular loading and are particularly useful for resolving protein-specific and subcellular heterogeneity in tension dynamics.

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.

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.

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.

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.