Developmental Biology

Centrosomes are vital eukaryotic organelles involved in regulating cell adhesion, polarity, mobility, and microtubule (MT) spindle assembly during mitosis. Composed of two centrioles surrounded by the pericentriolar material (PCM), centrosomes serve as the primary microtubule-organizing centers (MTOCs) in proliferating cells. The PCM is crucial for MT nucleation and centriole biogenesis. Centrosome numbers are tightly regulated, typically duplicating once per cell cycle, during the S phase. Deregulation of centrosome components can lead to severe diseases. While traditionally viewed as stable structures, centrosomes can be inactivated or disappear in differentiating cells, such as epithelial cells, muscle cells, neurons, and oocytes. Despite advances in understanding centrosome biogenesis and function, the mechanisms maintaining mature centrosomes or centrioles, as well as the pathways regulating their inactivation or elimination, remain less explored. Studying centrosome maintenance is challenging as it requires the uncoupling of centrosome biogenesis from maintenance. Tools for acute spatial-temporal manipulation are often unavailable, and manipulating multiple components in vivo is complex and time-consuming. This study presents a protocol that decouples centrosome biogenesis from maintenance, allowing the study of critical factors and pathways involved in the maintenance of the integrity of these important cellular structures.

Traditional tissue dissociation methods for bulk- and single-cell sequencing use various protease and/or collagenase combinations at temperatures ranging from 28 to 37 °C, which cause transcriptional cell stress that may alter data interpretation. Such artifacts can be reduced by dissociating cells in cold-active proteases, but few studies have shown that this improves cell-type specific transcription, particularly in tissues hypersensitive to mechanical integrity and extracellular matrix (ECM) interactions. To address this, we have dissociated zebrafish tendons and ligaments in subtilisin A at 4 °C and compared the results with 37 °C collagenase dissociation using bulk RNA sequencing. We find that high-temperature collagenase dissociation causes general cell stress in tendon fibroblasts (tenocytes) as reported in previous studies with other cell types, but also that high temperature specifically downregulates hallmark genes involved in tenocyte specification and ECM production in vivo. Our results suggest that cold-protease dissociation reduces transcriptional artifacts and increases the robustness of RNA-sequencing datasets such that they better reflect native in vivo tissue microenvironments.

One of the major factors contributing to aging and age-related diseases is the well-understood decline in the function of adult stem cells. Quantifying the degree of aging in adult stem cells is essential for advancing anti-aging mechanisms and developing anti-aging agents. However, no systematic approach to this exists. In this study, we developed a method to quantitatively assess the degree of aging in adult intestinal stem cells using a Drosophila midgut model and two aging markers. First, aging was induced in Drosophila with the desired genotype, and the anti-aging agent was administered 7 days before dissection. Then, the levels of two intestinal stem cell aging markers found in Drosophila (PH3 and γ-tubulin) were measured using immunohistochemistry. Finally, fluorescence microscopy was employed to count the number of aging markers and take images, which were analyzed using image analysis software. Using this approach, we quantitatively analyzed the effects of anti-aging agents on the aging of adult intestinal stem cells. This methodology is expected to significantly expedite the development of anti-aging agents and substantially reduce the research costs associated with aging-related studies.

Our goal is to understand how hematopoietic stem cell precursors emerge from vessels and to visualize their settling in developmental and more definitive niches that will persist in the adult. For this, we use as a biological model the zebrafish, which offers invaluable advantages owing to its transparency and small size, allowing high-resolution imaging and investigations of the entire animal. In vertebrate species, precursors of hematopoietic stem cells emerge from arterial vessels, mainly from the ventral side of the dorsal aorta. From there, they can either reside in the underlying vascular niche and/or pass through the vein to enter the blood circulation and conquer the caudal hematopoietic tissue, a functional equivalent to the fetal liver in mammals. Here, we provide experimental details of a protocol we have recently optimized to identify, based on mRNA in situ hybridization, precursors of hematopoietic stem cells while still embedded in the aortic wall (at the embryonic stage) as well as when they reside in specific niches a few days after emergence (at the early larval stage). Our experimental approach uses RNAscope technology, which allows combining high-sensitivity mRNA detection with high-resolution fluorescence confocal imaging to achieve spatial transcriptomics. Importantly, the small size of the probes allows better penetration inside tissues, which is a significant improvement in comparison to long mRNA probes; this is an invaluable advantage for reaching deeply embedded niches such as the ones of the pronephros region in the larva and, in addition, provides an increased signal-to-noise ratio.

The fate mapping technique is essential for understanding how cells differentiate and organize into complex structures. Various methods are used in fate mapping, including dye injections, genetic labeling (e.g., Cre-lox recombination systems), and molecular markers to label cells and track their progeny. One such method, the FlashTag system, was originally developed to label neural progenitors. This technique involves injecting carboxyfluorescein diacetate succinimidyl ester (CFSE) into the lateral ventricles of mouse embryos, relying on the direct uptake of dye by cells. The injection of CFSE into the lateral ventricle allows for the pulse labeling of mitotic (M-phase) neural progenitors in the ventricular zone and their progeny throughout the brain. This approach enables us to trace the future locations and differentiation paths of neural progenitors. In our previous study, we adapted this method to selectively label central nervous system–associated macrophages (CAMs) in the lateral ventricle by using a lower concentration of CFSE compared to the original protocol. Microglia, the brain's immune cells, which play pivotal roles in both physiological and pathological contexts, begin colonizing the brain around embryonic day (E) 9.5 in mice, with their population expanding as development progresses. The modified FlashTag technique allowed us to trace the fate of intraventricular CAMs, revealing that certain populations of microglia are derived from these cells. The optimized approach offers deeper insights into the developmental trajectories of microglia. This protocol outlines the modified FlashTag method for labeling intraventricular CAMs, detailing the CFSE injection procedure, evaluation of CFSE dilution, and preparation of tissue for immunohistochemistry.

Zebrafish and medaka are valuable model vertebrates for genetic studies. The advent of CRISPR-Cas9 technology has greatly enhanced our capability to produce specific gene mutants in zebrafish and medaka. Analyzing the phenotypes of these mutants is essential for elucidating gene function, though such analyses often yield unexpected results. Consequently, providing researchers with accessible and cost-effective phenotype analysis methods is crucial. A prevalent technique for investigating calcified bone development in these species involves using transgenic fish that express fluorescent proteins labeling calcified bones; however, acquiring these fish and isolating appropriate crosses can be time-consuming. We present a comprehensive protocol for visualizing ossified bones in zebrafish and medaka larvae and juveniles using calcein and alizarin red S staining, which is both economical and efficient. This method, applicable to live specimens during the ossification of bones, avoids apparent alterations in skeletal morphology and allows for the use of different fluorescent dyes in conjunction with transgenic labeling, thus enhancing the analysis of developmental processes in calcifying bones, such as vertebrae and fin rays.

Developing a physiologically relevant in vitro model of the respiratory epithelium is critical for understanding lung development and respiratory diseases. Here, we describe a detailed protocol in which the fetal mouse proximal epithelial progenitors were differentiated into 3D airway organoids, which contain terminal-differentiated ciliated cells and basal stem cells. These differentiated airway organoids could constitute an excellent experimental model to elucidate the molecular mechanisms of airway development and epithelial cell fate determination and offer an important tool for establishing pulmonary dysplasia disease in vitro.

Osteoclasts are terminally differentiated multinucleated giant cells that mediate bone resorption and regulate skeletal homeostasis under physiological and pathological states. Excessive osteoclast activity will give rise to enhanced bone resorption, being responsible for a wide range of metabolic skeletal diseases, ranging from osteoporosis and rheumatoid arthritis to tumor-induced osteolysis. Therefore, the construction of in vitro models of osteoclast-mediated bone resorption is helpful to better understand the functional status of osteoclasts under (patho)physiological conditions. Notably, it is essential to provide an in vivo–relevant bone substrate that induces osteoclasts to generate authentic resorption lacunae and excavate bone. Here, we summarize the experimental design of a reproducible and cost-effective method, which is suitable for evaluating the regulatory mechanisms and influence of molecular agonists and antagonists as well as therapeutics on osteoclast-mediated bone-resorbing activity.

Neuroscience incorporates manipulating neuronal circuitry to enhance the understanding of intricate brain functions. An effective strategy to attain this objective entails utilizing viral vectors to induce varied gene expression by delivering transgenes into brain cells. Here, we combine the use of transgenic mice, neonatal transduction with adeno-associated viral constructs harboring inhibitory designer receptor exclusively activated by designer drug (DREADD) gene, and the DREADD agonist clozapine N-oxide (CNO). In this way, a chemogenetic approach is employed to suppress neuronal activity in the region of interest during a critical developmental window, with subsequent investigation into its effects on the neuronal circuitry in adulthood.

All living organisms require the division of a cell into daughter cells for their growth and maintenance. During cell division, both genetic and cytoplasmic contents are equally distributed between the two daughter cells. At the end of cell division, cytoplasmic contents and the plasma membrane are physically separated between the two daughter cells via a process known as cytokinesis. Hundreds of proteins and lipids involved in the cytokinetic process have been identified; however, much less is known about the mechanisms by which these molecules regulate cytokinesis, being therefore an intense area of current research. Male meiotic cytokinesis in Drosophila melanogaster testes has been shown to be an excellent model to study cytokinesis in vivo. Currently, several excellent protocols are available to study cytokinesis in Drosophila testes. However, improved methods are required to study cytokinesis under in vitro and ex vivo conditions. Here, we demonstrate a simple method to perform live imaging on individual spermatocyte cysts isolated from adult testes. We evaluate amenability of this in vitro method for treatment with pharmacological agents. We show that cytokinesis is strongly inhibited upon treatment with Dynasore, a dynamin inhibitor known to block clathrin-mediated endocytosis. In addition, we also demonstrate an ex vivo method to perform live imaging on whole mount adult testes on gas permeable membrane chambers. We believe the protocols described here are valuable tools to study cytokinetic mechanisms under various genetic and treatment conditions.

Key features

• In vitro method to study male meiotic cytokinesis in dissected spermatocyte cysts.

• In vitro method allows acute treatment with various pharmacological agents to study cytokinesis.

• Ex vivo method to image male meiosis cytokinesis in intact adult testes.

• Requires 15–60 min to set up and could be imaged up to 6–12 h.

Graphical overview

In vitro and ex vivo live imaging of male meiotic cytokinesis in adult Drosophila testes