发育生物学

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.

Fluorescence in situ hybridization (FISH) is a cytological method used to visualize specific oligonucleotide sequences within the cell. This method relies on the specific binding of a fluorescence-tagged probe, a short stretch of single-stranded polynucleotide, to its complementary sequence in the DNA or RNA, forming stable double-stranded hybrids. Fluorochromes, such as fluorescein, Alexa Fluor, cyanine dyes, or rhodamine, are attached to these probes to help in detecting their presence within the cell. Based on sequence complementarity, FISH allows for the visualization of the DNA or RNA with which they have hybridized. The distribution of these fluorochrome-tagged probes can be observed under a fluorescence or confocal microscope. The oligo(dT) FISH technique specifically utilizes a fluorochrome-tagged stretch of 40–50 thymidine (T) oligonucleotides that binds to the poly(A) tails of mature mRNAs within the cell. Newly transcribing pre-mRNAs and certain non-coding RNAs may not have poly(A) tails and therefore cannot be detected by this method. This step-by-step protocol outlines the oligo(dT) FISH technique for visualizing the cellular distribution of polyadenylated mRNAs in the tissues of Drosophila and other related model organisms.

Extracellular vesicles (EVs) are critical mediators of cell–cell communication and play a key role in male reproductive biology by modulating sperm function. This protocol describes a robust and reproducible workflow for isolating EVs from ram seminal plasma using size-exclusion chromatography (SEC) and assessing their uptake by ram spermatozoa. In contrast to ultracentrifugation-based methods, SEC provides a gentle and more efficient isolation approach that preserves EV integrity and functionality. A central innovation of this protocol is the use of carboxyfluorescein succinimidyl ester (CFSE)-labeled seminal plasma EVs (SP-EVs) to evaluate their incorporation into sperm cells through two complementary detection platforms: (i) flow cytometry with standard resolution and (ii) confocal microscopy, for spatial confirmation of EV–sperm interactions. By bridging the gap between EV isolation and functional analysis, this protocol provides a valuable tool for investigating the role of EV–cell interactions. Specifically, it offers potential applications in male fertility preservation, biomarker discovery, and the development of EV-based therapeutic strategies in reproductive medicine.

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.