Biological Sciences

Peptergent is a novel class of amphipathic peptides that enables detergent-free extraction of membrane proteins (MPs) from lipid bilayers. This reagent self-assembles around hydrophobic transmembrane regions, forming stable, water-soluble complexes that can be isolated directly from biological membranes. Peptergent therefore bypasses the limitations imposed by traditional detergents, which often destabilize protein assemblies. Since detergents are completely avoided, MPs are directly amenable to structural and mass spectrometry (MS) analysis, thereby addressing their persistent underrepresentation in proteomic datasets and improving their accessibility in drug-screening strategies. We present here a streamlined protocol for MPs extraction with the Peptergent PDET-1, followed by exchange into His-tagged Peptidiscs for Ni-NTA-based affinity purification. The method encompasses membrane isolation, peptide preparation, protein extraction, clarification, and MPs exchange from Peptergents to Peptidiscs. This workflow yields an enriched membrane proteome compatible with downstream LC-MS/MS analysis for improved identification of multi-pass MPs.

Bottom-up proteomics workflows encompass several key stages, including sample preparation, data acquisition, and data analysis. Of these, sample preparation is the initial and critical stage, as it significantly influences the depth, reproducibility, and reliability of subsequent mass spectrometry–based analyses. While several main digestion strategies exist, including in-gel, in-solution, and filter-aided methods, each presents distinct trade-offs in terms of throughput, contamination removal, and applicability to complex biological matrices. The Suspension Trapping (S-Trap) method offers a compelling alternative by efficiently capturing and digesting proteins while removing interferents like sodium dodecyl sulfate (SDS), which can compromise downstream LC–MS/MS performance. This protocol details a S-Trap workflow optimized for biofluid proteomics, specifically plasma, serum, and cerebrospinal fluid (CSF). We describe two complementary formats: a manual tube-based procedure for individual or small-batch samples and a 96-well-plate-based system enabling high-throughput processing. The protocol integrates optional high-abundance protein depletion to enhance coverage of low-abundance analytes and includes steps for reduction, alkylation, digestion, and peptide elution for low total protein content samples, such as plasma, serum, and cerebrospinal fluid. By providing a detailed protocol, this work aims to improve the consistency and accessibility of S-Trap-based sample preparation, facilitating robust and reproducible discoveries in bottom-up proteomics.

The rising global incidence of pancreatitis, pancreatic cancer, and diabetes has increased the need for efficient in vivo gene manipulation approaches to study the pancreas and develop new therapies. Although transgenic mouse models are widely used, they are time-consuming and costly to generate and maintain. Systemic viral delivery methods offer greater flexibility but often lack pancreatic specificity and require high viral doses. Here, we describe a streamlined protocol for intrapancreatic ductal delivery of adeno-associated viruses (AAVs) for targeted gene delivery. Our protocol requires standard surgical equipment and can be implemented in most laboratories. Specifically, we adopted a clamping strategy at the hepatopancreatic duct near the liver, as well as beneath the major duodenal papilla at the duodenum. This strategy exposes the duodenal papilla, facilitating viral delivery, preventing backflow, and enabling efficient pancreatic transduction at lower viral doses. Overall, this method provides a fast, simple, and effective approach for pancreas-targeted gene manipulation, facilitating preclinical studies of pancreatic biology and disease.

The emergence of antimicrobial resistance and the persistence of Klebsiella pneumoniae biofilms represent significant challenges to public health. Hermetia illucens (HI) larvae are considered a sustainable reservoir of novel bioactive compounds. This protocol details a method for extracting fatty acids from HI larvae fat (AWME3 fraction) and studying their effects on multidrug-resistant and hypervirulent Klebsiella pneumoniae strains. Effects are evaluated by crystal violet and ethidium bromide uptake assays, motility assays (swimming, swarming, and twitching), minimal biofilm inhibitory and eradication concentration tests (MBIC/MBEC) for single, mixed, and mature biofilms, light, fluorescence, and scanning electron microscopy imaging, and microbial adhesion to solvents (MATS). This protocol offers a reliable methodology for evaluating the anti-biofilm and anti-virulence properties of natural compounds.

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.

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.

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.

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.

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.

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.