Biological Sciences

Tail vein catheterization in mice is a standard technique for precise drug delivery in pharmacological research, offering high accuracy and reproducibility. However, existing techniques face significant limitations in maintaining long-term stable catheter patency in awake, freely moving mice, and there is currently no standardized, detailed protocol for tail vein catheterization. Current methods suffer from high rates of catheter dislodgement, increased animal stress from repeated injections, and movement restrictions, all of which introduce confounding variables in behavioral and pharmacological studies. We have developed a simple and efficient fixation method that maintains stable tail vein catheter patency for more than 60 min while allowing complete freedom of movement. This protocol employs a strain relief loop design and multi-point fixation strategy, effectively preventing catheter dislodgement during extended periods while minimizing animal stress. This protocol has been successfully applied across multiple research areas, including metabolic studies, behavioral assessments, and neuropharmacological research in awake mice, achieving >95% catheter retention with normal animal behavior, providing a reliable technical platform for long-term awake-state research applications.

The CRISPR/Cas9 system is a cornerstone technology in genome editing. Delivery of pre-assembled Cas9 ribonucleoprotein (RNP) complexes exhibits distinct advantages, including reduced off-target effects and lower immunogenicity. Conventional methods for purifying Cas9 protein typically involve multi-step chromatography and the cleavage of fusion tag, which are time-consuming and result in diminished yields. In this study, we present a simplified, one-step purification strategy for functional Streptococcus pyogenes Cas9 (SpCas9) using the ubiquitin (Ub) fusion system in Escherichia coli. The N-terminal Ub fusion not only improves protein solubility but also facilitates high-yield production of the His-Ub-Cas9 fusion protein. Importantly, the Ub tag does not require proteolytic removal during purification, allowing direct one-step purification of the fusion protein via nickel-affinity chromatography. The purified His-Ub-Cas9 retains robust DNA cleavage activity in vivo, as validated in zebrafish embryos. This protocol greatly simplifies the production of functional Cas9 protein, facilitating its broad application in genome editing.

Protoplast systems are widely used in plant research as versatile platforms for studying cellular processes and validating gene editing tools. In maize, they are particularly valuable because stable transformation in immature embryos is slow and labor-intensive, often requiring months to regenerate plants. However, existing protocols often yield inconsistent results in protoplast recovery, transfection efficiency, and viability. We present an optimized protocol for maize mesophyll protoplast isolation and PEG-mediated transfection. Two-week-old etiolated seedlings are processed using vertical cutting, improving the yield and viability of protoplasts. Protoplasts are then immediately transformed with a CRISPR/Cas9 construct after isolation, via PEG4000 with only 10 μg of plasmid DNA, reducing the resource demands of standard methods. Modified washing and storage conditions extend transformed protoplast viability to seven days, enabling longer-term monitoring and expanded downstream analyses. Editing outcomes are quantified by sequencing target sites and calculating efficiency with Cas-Analyzer. This protocol provides a rapid, efficient, and reproducible method for the rapid evaluation of gene editing in maize. This protocol offers a methodology to accelerate agricultural crop studies and broader plant molecular biology.

Labeling cells with reporter genes allows researchers to visually identify specific cells and observe how they interact with each other in dynamic biological systems. Even though various labeling methods are now available, a specific description of gene knock-in labeling methods for human trophoblast stem cells (hTSCs) has not been reported. Here, we present a streamlined protocol for labeling hTSCs with the green fluorescent protein (GFP) reporter gene via CRISPR/Cas9-mediated knock-in of the gene into the adeno-associated virus site 1 (AAVS1) safe harbor locus. A commonly used hTSC cell line, CT29, was transfected with a dual plasmid system encoding the Cas9 endonuclease and an AAVS1-targeted guide RNA in one plasmid and a donor plasmid encoding a puromycin resistance gene and GFP reporter gene flanked by AAVS1 homology arms. Puromycin-resistant clonal cells were isolated, and AAVS1 integration was confirmed via PCR and sequencing of the PCR products. The labeled cells are proliferative and can give rise to extravillous cytotrophoblast cells (EVT) and the syncytiotrophoblast (ST). To our knowledge, this is the first report using the CRISPR/Cas9 system for AAVS1 integration of a reporter gene in human trophoblast stem cells. It provides an efficient tool to facilitate the study of human trophoblast development and function in co-culture systems and will be highly useful in developing clinical gene therapy-related plasmid constructs.

The cellular secretome is a rich source of biomarkers and extracellular signaling molecules, but proteomic profiling remains challenging, especially when processing culture volumes greater than 5 mL. Low protein abundance, high serum contamination, and sample loss during preparation limit reproducibility and sensitivity in mass spectrometry–based workflows. Here, we present an optimized and scalable protocol that integrates (i) 50 kDa molecular weight cutoff ultrafiltration, (ii) spin column depletion of abundant serum proteins, and (iii) acetone/TCA precipitation for protein recovery. This workflow enables balanced recovery of both low- and high-molecular-weight proteins while reducing background from serum albumin, thereby improving sensitivity, reproducibility, and dynamic range for LC–MS/MS analysis. Validated in human mesenchymal stromal cell cultures, the protocol is broadly applicable across diverse cell types and experimental designs, making it well-suited for biomarker discovery and extracellular proteomics.

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 offer numerous advantages as a vertebrate model because of their rapid development, high fecundity, transparent embryos, and ease of genetic manipulation. A wide variety of transgenic and mutant fish lines have been generated, and efficiently sharing these resources is crucial for advancing research. Zebrafish lines have typically been exchanged as early embryos, adult fish, or cryopreserved sperm, making transportation costly and logistically challenging. Here, we provide a protocol for preserving functional zebrafish sperm for more than 7 days at room temperature and subsequent in vitro fertilization using the preserved sperm. In this protocol, sperm collected either from the cloaca of an anesthetized male or from dissected testes is stored in L-15-based storage medium. Importantly, the storage medium, originally developed for zebrafish, is also applicable to medaka, another widely used vertebrate model. This sperm storage method allows researchers to ship sperm using low-cost methods and to investigate key factors for motility and fertilizing ability in those sperm.

Protein S-nitrosylation is a critical post-translational modification that regulates diverse cellular functions and signaling pathways. Although various biochemical methods have been developed to detect S-nitrosylated proteins, many suffer from limited specificity and sensitivity. Here, we describe a robust protocol that combines a modified biotin-switch technique (BST) with streptavidin-based affinity enrichment and quantitative mass spectrometry to detect and profile nitrosylated proteins in cultured cells. The method involves blocking free thiols, selective reduction of nitrosothiols, biotin labeling, enrichment of biotinylated proteins, and identification by tandem mass tag (TMT)-based quantitative mass spectrometry. Additionally, site-directed mutagenesis is employed to generate “non-nitrosylable” mutants for functional validation of specific nitrosylation sites. This protocol provides high specificity, quantitative capability, and versatility for both targeted and global analysis of protein nitrosylation.

The RNA-guided CRISPR-Cas9 endonuclease has been a transformative tool for laboratory biochemistry with huge potential as a precision therapeutic. This tool site-specifically cleaves double-stranded DNA following the recognition of a unique protospacer-adjacent motif (PAM). Activation of the protein–nucleic acid Cas complex has also been widely recognized to feature an allosteric mechanism dependent on structural remodeling and interdomain crosstalk. Biophysical methods have probed the impact of allosteric perturbations on cleavage and specificity of Cas9, with the aim of engineering enhanced Cas effectors. These studies include Cas9 from thermophilic organisms that edit at higher temperatures and are active in human plasma. Validation of biophysical insights has necessitated the quantitation of DNA cleavage in vitro and, subsequently, the adaptation of established protocols to encompass temperature-dependent function that is evident in extremophilic Cas systems, such as Cas9 from Geobacillus stearothermophilus and the mesophilic SpCas9. This protocol is advantageous for probing functional temperature ranges of DNA cleavage that can theoretically be applied to any Cas-RNP system.

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.