Developmental Biology

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.

Telomere shortening is a hallmark of human aging, and telomerase regulation plays a critical role in cellular proliferation and replicative senescence. In human cells, telomere length imposes a limit on proliferative potential, a phenomenon known as the Hayflick limit. However, species-specific differences in telomere dynamics and telomerase regulation between humans and mice present challenges to using mice as accurate models for human telomere-related research. To address this limitation, we engineered a humanized telomerase gene (hmTert) in mice by replacing the non-coding sequences within the mouse Tert locus (mTert) with corresponding regulatory sequences from the human TERT gene. Breeding of these genetically modified mice resulted in progressive telomere shortening over successive generations, ultimately reaching human-like lengths (below 10 kb). This protocol outlines the development of this humanized telomere mouse model, referred to as HuT mice, offering a robust platform for studying human telomere biology and aging-related diseases.

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.