Developmental Biology

Congenital renal disorders, such as the Potter sequence, result from renal dysgenesis. To explore a prenatal therapeutic approach for fetuses with kidney insufficiency, we established an in utero transplantation protocol using donor fetal kidneys. Although numerous rodent studies have reported cellular injections into fetal recipients, no protocol to date has described whole-organ transplantation during gestation. Here, we present a step-by-step method for grafting donor fetal kidneys (embryonic day 14.0–16.5) into allogeneic rat fetuses at embryonic day 18.0–18.5, resulting in term neonates that retain the grafts postnatally. A 15–16 G needle preloaded with the donor kidney is inserted transuterinely, depositing the organ into the subcutaneous space of the fetus. Four days later, the term pups are delivered naturally and evaluated for graft development. This protocol enables organ-level transplantation and longitudinal assessment of graft maturation within the unique fetal environment, which differs markedly from adult settings in terms of growth factor availability and immune reactivity. To our knowledge, this is the first protocol to successfully achieve whole-organ transplantation directly into fetuses in utero. Therefore, the model provides a valuable platform for studying developmental organogenesis, fetal immunology, and regenerative strategies that leverage embryonic cues.

Underwater noise is a growing source of anthropogenic pollution in aquatic environments. However, few studies have evaluated the impact of underwater noise on aquatic invertebrates. More importantly, studies involving early developmental stages have been poorly addressed. Significant limitations are due to the lack of standardized protocols for working in the laboratory. Particularly, the design of uniform procedures in the laboratory is important when working with species that inhabit short-term changing habitats, such as estuaries, which makes it difficult to carry out repeated experiments in the natural habitat. Besides, controlling for environmental variables is also important when assessing the effect of a stressor on the physiological parameters of individuals. This experimental protocol addresses that gap by offering an adaptable laboratory-based method to evaluate sublethal physiological responses to sound exposure under highly controlled conditions. Here, we present a reproducible and accessible laboratory protocol to expose crabs to recorded boat noise and evaluate physiological responses using oxidative stress biomarkers. The method is designed for ovigerous females, as we evaluated the effects on embryos and early life stages (i.e., larvae), but it can be readily adapted to different life stages of aquatic invertebrates. A key strength of this protocol is its simplicity and flexibility: animals are exposed to noise using submerged transducers under well-controlled laboratory conditions, ensuring consistency and repeatability. Following exposure, tissues or whole-body samples can be processed for a suite of oxidative stress biomarkers—glutathione-S-transferase (GST), catalase (CAT), lipid peroxidation (LPO), and protein oxidation. These biomarkers are highly responsive, cost-effective indicators that provide a sensitive and early readout of sublethal stress. Together, the exposure and analysis steps described in this protocol offer a powerful and scalable approach for investigating the physiological impacts of underwater noise in crustaceans and other aquatic invertebrates.

Primary cilia are evolutionarily conserved organelles that play critical roles in brain development. In the developing cortex, neural progenitors extend their primary cilia into the ventricular surface, where the cilia act as key signaling hubs. However, visualizing these cilia in a systematic and intact manner has been challenging. The commonly used cryostat sectioning only provides a limited snapshot of cilia on individual sections, and this process often disrupts the ciliary morphology. By contrast, the previously established whole-mount technique has been shown to preserve ciliary architecture in the adult mouse cortex. Here, we adapt and optimize the whole-mount approach for embryonic and neonatal brain, allowing robust visualization of ciliary morphology at the ventricular surface during development. This protocol describes step-by-step procedures for whole-mounting and immunostaining delicate embryonic and neonatal mouse cortices, enabling direct visualization of cilia in neural progenitors in the developing brain.

In mammals, the semen is ejaculated into the female reproductive tract, and the sperm travel to the oviduct to fertilize the egg. A comprehensive understanding of the pre- and post-ejaculatory intrauterine environment is one of the key points for overcoming infertility; however, the dynamics of the intrauterine environment and its physiological role in the uterus, namely in the internal fertilization process, remain unclear. Conventional methods for collecting uterine fluids from the uterus post-ejaculation of mice show challenges regarding the ambiguous ejaculation timing. Here, we established a method for a mating environment with exact ejaculation timing. We also created a simple method for collecting pre- and post-ejaculatory uterine fluid without using forceps. Our methods achieved time-dependent biochemical and histological analyses of uterine fluids to provide fundamental information regarding protein composition and uterine structure changes during pre- and post-ejaculation. This protocol is suitable for analyzing temporal changes in reproductive phenomena, thereby contributing to elucidating the physiological role of the uterus in the process of intrauterine fertilization.

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.

Peripheral nerve injuries (PNIs) often result in incomplete functional recovery due to insufficient or misdirected axonal regeneration. Balanced regeneration of myelinated A-fibers and unmyelinated C-fibers is essential for functional recovery, making it crucial to understand their differential regeneration patterns to improve PNI treatment outcomes. However, immunochemical staining does not clearly differentiate between A- and C-fiber axons in whole-mount nerve preparations. To overcome this limitation, we developed a modified protocol by optimizing the immunostaining to restrict the antibody access to myelinated axons. This enables visualization of A-fibers by myelin sheath labeling, while allowing selective staining of unmyelinated C-fiber axons. As a result, A- and C-fibers can be reliably distinguished, facilitating accurate analysis of their regeneration in both normal and post-injury conditions. Combined with confocal microscopy, this approach supports efficient screening of whole-mount nerve preparations to evaluate fiber density, spatial distribution, axonal sprouting, and morphological characteristics. The refined technique provides a robust tool for advancing PNI research and may contribute to the development of more effective therapeutic strategies for nerve repair.

N⁶-methyladenosine (m⁶A) is the most abundant internal modification in mRNA and is regulated primarily by the balance between the METTL3 methylase complex and two demethylases, FTO (fat mass and obesity-associated protein) and ALKBH5 (α-ketoglutarate-dependent dioxygenase alkB homolog). Reflecting this prevalence, m⁶A participates in virtually every step of RNA metabolism, influencing a wide range of physiological and pathological processes. The first step in studying m⁶A is genome-wide mapping, typically performed by m⁶A-seq, which sequences RNA fragments immunoprecipitated with an m⁶A-specific antibody. This is followed by identification of RRACH motifs (R = A or G; H = A, C, or U) within these sequences, with m⁶A being located at the third nucleotide. The second step involves mutating the putative m⁶A sites to establish a causal link between the modification and downstream biological effects. Since the mapping step has been covered in several detailed protocols, this article focuses on the second step—mutagenesis of RRACH motifs and subsequent functional analysis of the mutations by ectopic expression. The 3′ untranslated region (UTR) of the mouse Runx2 gene is used as an example. The mutant and wild-type sequences are inserted into a luciferase reporter vector and transfected into 293FT cells to evaluate how loss of m⁶A affects luciferase protein levels. The same reporter plasmids are also used in an RNA stability assay with a transcription inhibitor. Although site-specific demethylation of endogenous mRNA would be preferable, it remains technically challenging despite many attempts. Thus, ectopic expression of the mutated target gene remains a widely used and practical alternative.

Long noncoding RNAs (lncRNAs) are increasingly understood to play important roles in cell biology, development, and disease, though the vast majority of annotated lncRNAs have yet to be functionally characterized. Disrupting lncRNAs is often challenging owing to their tolerance for mutations (e.g., single-nucleotide polymorphisms and short indels) along with the limitations of other genetic knockdown strategies such as RNA interference (RNAi). Here, we describe a protocol to achieve robust knockdown of lncRNAs in the fruit fly Drosophila using a self-cleaving ribozyme. The 111-bp ribozyme cassette, which consists of the N79 hammerhead ribozyme flanked by flexible linker sequences, is inserted into transcript regions of lncRNA genes using CRISPR/Cas9-mediated homology-directed repair (HDR). The fluorescent eye transformation marker is then removed using a piggyBac transposase, leaving no other modifications at the lncRNA locus save the ribozyme cassette insertion. When transcribed as part of the lncRNA, the ribozyme folds and catalyzes its own self-cleavage, resulting in two RNA cleavage fragments. The efficacy of lncRNA knockdown is then evaluated using reverse transcription quantitative PCR (RT-qPCR) and single-molecule RNA fluorescence in situ hybridization (smFISH). This approach has resulted in efficient knockdown of both nuclear and cytoplasmic lncRNAs in Drosophila, with knockdown of steady-state RNA levels in 3' cleavage fragments typically exceeding 90% and no evidence of off-target effects. The method can also be applied to protein-coding genes in order to knock down specific mRNA isoforms. Thus, self-cleaving ribozymes are a valuable addition to the genetic toolkit in Drosophila.

Translation is a key step in decoding the genetic information stored in DNA. Regulation of translation is an important step in gene expression control and is essential for healthy organismal development and behavior. Despite the importance of translation regulation, its impact and dynamics remain only partially understood. One reason is the lack of methods that enable the real-time visualization of translation in the context of multicellular organisms. To overcome this critical gap, microscopy-based methods that allow visualization of translation on single mRNAs in living cells and animals have been developed. A powerful approach is the SunTag system, which enables real-time imaging of nascent peptide synthesis with high spatial and temporal resolution. This protocol describes the implementation and use of the SunTag translation imaging system in the small round worm Caenorhabditis elegans. The protocol provides details on how to design, carry out, and interpret experiments to image translation dynamics of an mRNA of interest in a cell type of choice of living C. elegans. The ability to image translation live enables better understanding of translation and reveals the mechanisms underlying the dynamics of cell type–specific and subcellular localization of translation in development.