发育生物学

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.

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.

Metamorphosis in class Insecta is defined as the drastic transition of an organism from a larval stage to a morphologically very different adult form. Insects such as Drosophila melanogaster undergo metamorphosis, which requires precise temporal control, exerted by ecdysteroids, the primary regulators of these developmental transitions. The active form of ecdysone is 20-hydroxyecdysone (20E), which acts in pulses that trigger the onset of different developmental stages. A massive pulse of 20E occurs between the prepupal and pupal stages, triggering pupariation. Precise quantification of 20E levels is critical for understanding how this hormonal signaling governs the developmental timing of different stages. This protocol is sensitive for detecting low levels of 20E and describes a methanol-based extraction method coupled with a competitive ELISA to quantify whole-body 20E levels in third-instar Drosophila larvae. Further, this protocol enables detection of differences in ecdysone titers among genotypes affecting ecdysone biosynthesis, particularly at the late third-instar larval stage. This protocol can also be utilized for disease-causing insects (Aedes aegypti), as well as economically relevant beneficial insects (Apis mellifera), to study the effects of ecdysteroid analogs during development.

The aorta–gonad–mesonephros (AGM) region is the site where hematopoietic stem cells (HSCs) first emerge during development, and is therefore widely used to study in vivo hematopoiesis and to discover novel regulatory mechanisms. The endothelial-to-hematopoietic transition (EHT) process can be directly observed via immunofluorescence on frozen sections of the AGM region. However, the mouse AGM region is extremely delicate and lies deep within the embryo, between the notochord and the somatic mesoderm. Here, we present a step-by-step protocol covering embryo collection, fixation, dehydration, and embedding with a defined orientation, followed by frozen sectioning, immunofluorescence staining, and confocal imaging. The protocol is highly reproducible and easy to follow and provides clear instructions on orienting the embryo and anatomically locating AGM. By filling a technical gap, the protocol can enable researchers to reliably study HSC emergence and EHT in the mouse embryonic AGM.

Limb development requires the coordination of multiple cell types, including the limb bud mesoderm and surface ectoderm, by the apical ectodermal ridge (AER), a specialized signaling center secreting numerous morphogens. Characterizing these cell–cell interactions is crucial for understanding limb morphogenesis, but they are challenging to study in vivo. Furthermore, existing in vitro models do not capture the multilineage complexity of the limb. We recently developed a robust 7-day differentiation protocol using mouse embryonic stem cells (mESCs) to generate heterogeneous cultures containing cells with characteristics of the limb bud mesoderm, surface ectoderm, and AER. Dissociating and reaggregating these cultures in low attachment 96-well plates forms budoids, organoids that display certain limb bud–like features. Budoids undergo chondrogenesis-mediated symmetry breaking and elongation within 5 days of culture. Altogether, our protocols have enabled the study of cell–cell interactions in limb development and provide an easily scalable model adaptable for various applications, including drug testing and congenital disorder modeling.

Embryonic blood formation encompasses the independent generation of different cell types in distinct cellular and anatomical environments, reflecting highly coordinated specific hierarchies of interacting tissues. Despite widespread use of embryonic stem cells (ESC) and induced pluripotent stem cell (iPSC)-based models to attempt to capture blood development in vitro and generate hematopoietic stem cells (HSC), a system that fully captures the spatial and temporal complexity of embryonic hematopoiesis is still lacking. In recent years, gastruloid models have emerged as powerful representations of early development, demonstrating self-organizing behaviors such as symmetry breaking, elongation, multi-axis formation, somitogenesis, and early organogenesis, with striking parallels to embryonic processes. Here, we present a protocol to generate hemogenic gastruloids (haemGx) from mouse ESC (mESC) that closely recapitulates the multi-stage, multi-niche process of blood formation and generates developmentally accurate hematopoietic progenitors. The haemGx model has been proven valuable in understanding embryonic hematopoiesis, as well as an in vitro model of forms of infant leukemia with an embryonic, in utero origin.

Aging is associated with progressive accumulation of senescent cells, which contribute to tissue dysfunction and organismal decline. Conventional approaches for assessing cellular senescence, such as histological or immunofluorescence analyses of fixed tissue sections and flow cytometry, require tissue collection, thereby precluding longitudinal in vivo studies. To enable the analysis of cellular senescence in a living vertebrate model, we have previously generated a cdkn1a (p21)-driven GFP reporter line that was established in the transparent klara background of Nothobranchius furzeri. Here, we describe a protocol for in vivo light-sheet microscopy of the reporter activity as readout for senescence-associated cell cycle arrest with single-cell resolution. The procedure involves anesthesia and mounting of fish for stable positioning within the imaging chamber, with particular attention to animal welfare considerations. It further includes the acquisition of three-dimensional image stacks and subsequent image processing. The workflow allows monitoring of GFP-positive cells in intact living killifish at different developmental stages. Although imaging depth remains limited despite organismal transparency, this method provides high-resolution volumetric imaging with minimal phototoxicity and enables analysis of senescence dynamics in a short-lived vertebrate model. It is currently performed as a terminal procedure under approved ethical regulations, but longitudinal imaging would also be possible with additional ethical authorization.

Thin membrane protrusions in cells help them communicate, create traction forces during their movement, and coordinate complex development in multicellular organisms. These structures include cytonemes, tunneling nanotubes, and microtubule-based nanotubes (MT-nanotubes), each with a different cytoskeletal constitution and function. Actin-based cytonemes help deliver signaling molecules, while microtubule-based nanotubes assist with transporting vesicles and organelles. Despite their physiological role, we still do not fully understand how these thin membrane protrusions form and function. In this study, we introduce an improved live-cell imaging method to observe polar cell protrusions during micropyle morphogenesis in developing Drosophila eggs. This technique combines precise developmental staging, careful dissection, and optimized ex vivo culture conditions to maintain tissue health during extended imaging. We also fine-tuned the imaging settings to reduce phototoxicity and thermal stress. This allows for continuous, high-resolution tracking of protrusion dynamics in real time. Our protocol addresses major drawbacks of fixed-tissue methods by capturing the entire process of protrusion formation, extension, and remodeling in intact living tissue. Additionally, it works well with drugs, making it a useful tool for functional studies. Overall, this approach builds a strong foundation for exploring membrane protrusion biology. It can also be applied to investigate similar developmental processes in other systems, aiding our understanding of normal development and diseases.

Whole-mount techniques are widely used in medical and biological research to analyze protein expression and tissue organization in intact specimens. Traditional approaches for protein localization include section-based immunohistochemistry and in situ hybridization; however, these methods can be limited by tissue disruption and loss of spatial context. Whole-mount protocols generally involve tissue fixation, permeabilization, and staining with specific probes, but their effectiveness varies depending on the antigen–antibody combination and the specimen type. Consequently, no universal protocol is suitable for all experimental conditions. This protocol presents a detailed whole-mount immunostaining protocol for evaluating tyrosine hydroxylase (TH) expression, a key marker of dopaminergic neurons, in zebrafish (Danio rerio) larvae. The procedure outlines critical steps from sample preparation to staining optimization to ensure reproducible and specific signal detection. This approach enables accurate visualization and analysis of dopaminergic neuron distribution in intact larvae. The protocol offers a reliable and adaptable approach that preserves tissue integrity, enables three-dimensional visualization, and is particularly well-suited for developmental and neurobiological studies using zebrafish larvae.

Neural tube closure is a critical process that transforms the neural plate, an open epithelial tissue, into the closed tube that serves as the structural basis of the central nervous system. Defects in this process are among the most common and severe developmental diseases in the human population, with failures in cranial closure accounting for approximately one-third of total defects. However, the cell and tissue mechanisms that drive cranial closure remain opaque relative to the better studied process of spinal closure, in large part due to the unique challenges in characterizing cranial tissues. Here, we present protocols for quantifying cell dynamics and tissue-level remodeling events that enable highly spatiotemporally resolved investigations of the causes of cranial closure defects in mouse embryos. These include brightfield morphometric approaches, fluorescent staining and confocal imaging, and quantitative pipelines to analyze these image-based datasets. At the conclusion of these approaches, users will be able to quantify several parameters of overall tissue shape in the cranial neural tissues and produce rich quantitative datasets about cell-level parameters, particularly apical cell area. These can be used to identify correlative and causative differences between mutants and control embryos. Given their flexibility, many of these approaches can be generalized to other tissue morphogenetic contexts.