Developmental Biology

The female reproductive tract is comprised of different regions, each with distinctive physiological characteristics. One of them is the fallopian tubes, which are vital for human reproductive health and success. The ability to model their function and physiology is of utmost importance. So far, in vitro models have been based on a few immortalized or cancer cell lines derived from fallopian tube cells that lacked differentiated, specialized cell types and did not allow for the study of cancer initiation due to their implicit biases. Organoids, in contrast, overcome these limitations and provide an advanced, three-dimensional system for the study of healthy fallopian tube physiology and pathology. Fallopian tube organoids are comprised of epithelial progenitors that can be enriched using chemical or hormonal treatment into the different cell types that are found in the in vivo tissue, namely detyrosinated-tubulin-positive ciliated cells or paired-box protein 8 (PAX8)-positive secretory cells. This protocol provides a step-by-step guide for the establishment and maintenance of a long-term culture of organoids from healthy human fallopian tube tissue. The organoid model described here closely mimics the in vivo physiology and anatomy of human fallopian tube epithelium and provides a comprehensive basis for future studies on its underlying molecular characteristics and possible pathology.

Infertility has emerged as a global health concern, impacting around 8%–12% of couples during their reproductive years. Due to limitations in obtaining human biological samples, mouse models have been widely used for investigating gene functions. Fertility assessment in mouse models is a critical component in reproductive biology for studying gene function and elucidating mechanisms of reproductive disorders. However, natural mating observation of mice may yield inconsistent results, especially in the absence of standard guidelines, prolonged experimental cycles, and operational complexity. This protocol establishes a comprehensive breeding strategy for evaluating murine fertility through systematic vaginal plug monitoring and litter size quantification within defined timeframes. Key steps include (1) standardized male–female pairing protocols, (2) daily vaginal plug inspection, and (3) longitudinal tracking of pregnancy outcomes. This protocol presents a straightforward and easily implementable protocol for mouse mating cage setup and statistical analysis, enabling reliable fertility assessment under natural breeding conditions.

Immunofluorescence staining is a technique that permits the visualization of components of various cell preparations. Manchette, a transient structure that is only present in elongating spermatids, is involved in intra-manchette transport (IMT) for sperm flagella formation. Sperm flagella are assembled by intra-flagellar transport (IFT). Due to the big complexes formed by IMT and IFT components, it has been challenging to visualize these components in tissue sections. This is because the proteins that make up these complexes overlap with each other. Testicular tissue is digested by a combination of DNase I and Collagenase IV enzymes and fixed by paraformaldehyde and sucrose. After permeabilization with Triton X-100, testicular cells are incubated with specific antibodies to detect the components in the manchette and developing sperm tails. This method allows for cell type–specific resolution without interference from surrounding cells like Sertoli, Leydig, or peritubular myoid cells. Additionally, isolated cells produce cleaner immunofluorescence signals compared to other methods like tissue section/whole mount, making this method the best fit for visualizing protein localization in germ cells when spatial context is not being considered. Hence, this protocol provides the detailed methodology for isolating male mice germ cells for antibody-targeted immunofluorescence assay for confocal/fluorescence microscopy.

CRISPR-Cas9 has democratized genome engineering due to its simplicity and efficacy. Adapted from a bacterial defense mechanism, CRISPR-Cas9 comprises the Cas9 endonuclease and a site-specific guide RNA. In vivo, the Cas9 ribonucleoprotein (RNP) can target specific genomic loci and generate double-strand breaks. Eukaryotic endogenous DNA repair mechanisms recognize the cut site and attempt to repair the DNA either by non-homologous end joining, which introduces insertions/deletions, resulting in a loss of reading frame in coding genes, or through homology-directed repair that maintains the reading frame. The latter approach allows the insertion of fluorescent reporter sequences in frame with protein-coding genes in order to monitor gene expression and protein dynamics in cells and whole organisms. Here, we provide a protocol for targeting endogenous genes to introduce sequences coding for fluorescent reporters in medaka (Oryzias latipes). The method is simple, robust, and efficient, thus facilitating straightforward organismal genome editing.

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.

The silkworm Bombyx mori has been extensively utilized in sericulture and serves as a representative model insect of Lepidoptera in various fields of life sciences and applied research. In recent years, its significance has further increased in molecular genetics and functional genomics. Germline transformation and genome editing in B. mori require the injection of vector solutions into early embryos; however, the thick eggshell of B. mori presents a significant challenge for microinjection. Conventional methods involve arranging eggs, pre-pierced with a tungsten needle, followed by solution injection, making the process both time-consuming and technically demanding. Here, we describe a simplified and more efficient microinjection protocol. Unlike conventional approaches, our method eliminates the need for egg removal from the egg-laying sheet and egg alignment on the slide glass by allowing injections to be performed directly on eggs retained on the egg-laying sheet. A thick-walled glass capillary, capable of penetrating the rigid eggshell, is used to directly pierce the eggshell and deliver the solution. By eliminating the need for egg alignment and micromanipulator operation, this protocol significantly enhances efficiency, enabling higher-throughput embryo injections within a shorter time frame. Moreover, this approach holds potential for application to other insect species with similarly thick eggshells.

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.

Super-resolution imaging of RNA–protein (RNP) condensates has shown that most are composed of different immiscible phases reflected by a heterogenous distribution of their main components. Linking RNA–protein condensate’s inner organization with their different functions in mRNA regulation remains a challenge, particularly in multicellular organisms. Drosophila germ granules are a model of RNA–protein condensates known for their role in mRNA storage and localized protein production in the early embryo. Present at the posterior pole of the embryo within a specialized cytoplasm called germplasm, they are composed of maternal mRNAs as well as four main proteins that play a key role in germ granule formation, maintenance, and function. Germ granules are necessary and sufficient to drive germ cell formation through translational regulation of maternal mRNAs such as nanos. Due to their localization at the posterior tip of the ovoid embryo and small size, the classical imaging setup does not provide enough resolution to reach their inner organization. Here, we present a specific mounting design that reduces the distance between the germ granule and the objectives. This method provides optimal resolution for the imaging of germ granules by super-resolution microscopy, allowing us to demonstrate their biphasic organization characterized by the enrichment of the four main proteins in the outermost part of the granule. Furthermore, combined with the direct visualization of nanos mRNA translation using the Suntag approach, this method enables the localization of translation events within the germ granule’s inner organization and thus reveals the spatial organization of its functions. This approach reveals how germ granules serve simultaneously as mRNA storage hubs and sites of translation activation during development. This work also highlights the importance of considering condensates’ inner organization when investigating their functions.