发育生物学

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.

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.

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.

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.

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.

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.

Repetitive increases of intracellular calcium ions (Ca²⁺ oscillations) control cellular functions in various biological events, including meiotic resumption after fertilization. Sperm-derived substances enter the cytoplasm of mature oocytes by sperm fusion, causing Ca²⁺ oscillations. Sperm-independent Ca²⁺ oscillations are also induced in immature oocytes isolated from the ovaries of neonatal to adult mice. The presence of Ca²⁺ oscillations may contribute to subsequent oocyte quality; however, its physiological role and molecular mechanism are unclear. Here, we describe a method of collecting immature oocytes from the ovaries of juvenile (12, 15, and 21 days after birth) and adult mice and monitoring their Ca²⁺ oscillations. Since mouse oocytes are larger than other types of cells, they are a useful model for studying spatiotemporal patterns and the mechanism of Ca²⁺ oscillations in various types of cells. This method can be applied to other rodents due to similarities in oocyte size and developmental processes. Furthermore, the use of various fluorescent probes enables visualization of organelle rearrangement. The mechanism of interaction between oocytes and somatic cells differs between juvenile and adult mice. Therefore, two distinct methods are employed for oocyte collection.

Nowadays, the use of 3D cultures (organoids) is considered a valuable experimental tool to model physiological and pathological conditions of organs and tissues. Organoids, retaining cellular heterogeneity with the presence of stem, progenitor, and differentiated cells, allow the faithful in vitro reproduction of structures resembling the original tissue. In this context, the growth of endometrial organoids allows the generation of 3D cultures characterized by a hollow lumen, secretory activity, and apicobasal polarity and displaying phenotypical modification in response to hormone stimulation. However, a limitation in currently used models is the absence of stromal cells in their structure; as a result, they miss epithelial–stromal interactions, which are crucial in endometrial physiology. We developed a novel 3D model to generate endometrial organoids grown in floating Matrigel^TM droplets in the presence of standard culture medium. From a structural point of view, these novel floating 3D cultures develop as gland-like structures constituted by epithelial cells organized around a central lumen and retain the expression of endometrial and decidual genes, like previously published organoids, although with a phenotype resembling hormonally differentiated structures. Importantly, floating organoids retain stromal cells which grow in close contact with the epithelial cells, localized within the internal or external portion of the organoid structure. In summary, we present a simple and rapid model for generating 3D endometrial organoids that preserve epithelial–stromal cell interactions, promoting the formation of differentiated organoids and enabling the study of reciprocal modulation between epithelium and stroma.

Zebrafish are a powerful model for investigating vascular and lymphatic biology due to their genetic tractability and optical transparency. While translating ribosome affinity purification (TRAP) has been widely applied in other systems, its application in zebrafish has remained limited. Here, we present an optimized TRAP protocol for isolating ribosome-associated mRNAs from endothelial cells in vivo, without the need for cell dissociation or sorting. Using a novel transgenic zebrafish line, which expresses HA-tagged Rpl10a under the mrc1a promoter, we enriched actively translating endothelial transcripts. Differential expression analysis revealed robust upregulation of vascular and lymphatic genes including flt4, kdrl, and lyve1b. This approach captures the endothelial cell translatome with high specificity and offers a robust platform for investigating the molecular mechanisms of endothelial biology under genetic, environmental, or toxicological perturbations.

Adipose cells vary functionally, with white adipocytes storing energy and brown/beige adipocytes generating heat. Mouse and human subcutaneous white adipose tissue (WAT)-derived stromal vascular fraction (SVF) provides mesenchymal stem cells (MSCs) that can be differentiated into thermogenic adipocytes using pharmacological cocktails. After six days of browning induction, these cells exhibited significant upregulation of thermogenic markers (UCP1, Cidea, Dio2, PRDM16) along with adipogenic genes (PPARγ, aP2), showing enhanced thermogenic potential. This in vitro system offers a practical platform to study adipogenesis and thermogenic regulation.