Transfecting neurons remains technically challenging due to their sensitivity. Conventional methods, such as Lipofectamine 2000 or Lipofectamine RNAiMAX, often result in significant cytotoxicity, which limits their utility. Although lentiviral transfection offers high efficiency, it is hindered by high costs and complex procedures. This experiment employs a small interfering RNA (siRNA)-specific transfection reagent from the Kermey company. This reagent is a novel nanoparticle-based lipid material designed for the efficient delivery of oligonucleotides, including siRNA, into a wide range of cell types. Its efficacy in achieving high transfection efficiency in neurons, however, has not yet been established. After several days of in vitro neuronal culture, researchers can perform a simple transfection procedure using this reagent to achieve robust transfection efficiency. Notably, the protocol does not require medium replacement 6-8 h post-transfection, streamlining the workflow and minimizing cellular stress. Key features • Based on Kermey's siRNA-specific transfection reagent, we present a method for efficient in vitro transfection of siRNA into primary cultured mouse cortical neurons. • No observable adverse effects are detected in the transfected neurons during the entire experiment. • This method enables consistent and efficient knockdown of the target protein. • Phosphoglycerate dehydrogenase (PHGDH) siRNA and siNC (negative control) siRNA can be transfected into neuronal cells after 72 h of in vitro culture.

Expansion microscopy (ExM) is an innovative and cost-effective super-resolution imaging technique that enables nanoscale visualization of biological structures using conventional fluorescence microscopes. By physically enlarging biological specimens, ExM circumvents the diffraction limit and has become an indispensable tool in cell biology. Ongoing methodological advances have further enhanced its spatial resolution, labeling versatility, and compatibility with diverse sample types. However, ExM imaging is often hindered by sample drift during image acquisition, caused by subtle movements of the expanded hydrogel. This drift can distort three-dimensional reconstruction, compromising both visualization accuracy and quantitative analysis. To overcome this limitation, we developed 3D-Aligner, an advanced and user-friendly image analysis software that computationally corrects sample drift in fluorescence microscopy datasets, including but not limited to those acquired using ExM. The algorithm accurately determines drift trajectories across image stacks by detecting and matching stable background features, enabling nanometer-scale alignment to restore structural fidelity. We demonstrate that 3D-Aligner robustly corrects drift across ExM datasets with varying expansion factors and fluorescent labels. This protocol provides a comprehensive, step-by-step workflow for implementing drift correction in ExM datasets, ensuring reliable three-dimensional imaging and quantitative assessment.Key features• 3D-Aligner precisely corrects sample drift in expansion microscopy (ExM) datasets, enabling reliable 3D reconstruction and robust quantitative analysis.• Utilizes background feature detection and feature matching across z-planes to achieve nanoscale-precision drift correction.• 3D-Speckler, which is a MATLAB-based software platform, offers a customizable and user-friendly interface.• Outperforms conventional registration tools across varying expansion factors and labeling conditions and is equally applicable to non-ExM datasets.

Genetic modification is essential for understanding parasite biology, yet it remains challenging in Plasmodium. This is partially due to the parasite’s low genetic tractability and reliance on homologous recombination, since the parasites lack the canonical non-homologous end-joining pathway. Existing approaches, such as the PlasmoGEM project, enable genome-wide knockouts but remain limited in coverage and flexibility. Here, we present the Plasmodium berghei high-throughput (PbHiT) system, a scalable CRISPR-Cas9 protocol for efficient genome editing in rodent malaria parasites. The PbHiT method uses a single cloning step to generate vectors in which a guide RNA (gRNA) is physically linked to short (100 bp) homology arms, enabling precise integration at the target locus upon transfection. The gRNA also serves as a unique barcode, allowing pooled vector transfections and identification of mutants by downstream gRNA sequencing. The PbHiT system reliably recapitulates known mutant growth phenotypes and supports both knockout and tagging strategies. This protocol provides a reproducible and scalable tool for genome editing in P. berghei, enabling both targeted functional studies and high-throughput genetic screens. Additionally, we provide an online resource covering the entire P. berghei protein-coding genome and describe a step-by-step pooled ligation approach for large-scale vector production.Key features• PbHiT provides a high-throughput CRISPR-Cas9 genome editing platform optimised for Plasmodium berghei experimental infections in rodents.• This protocol enables efficient and reproducible generation of knockout and tagged parasite lines using short homology arms.• This protocol is supported by a free online resource for P. berghei gene construct design and requires basic knowledge of cloning.• Transfection of Plasmodium berghei requires experience in handling mice/rats, an ethical permit, and an animal facility.

During herpesvirus replication, capsids are assembled inside the nucleus and translocated into the cytosol by a non-canonical nucleocytoplasmic export process termed nuclear egress. Traditional methods of measuring nuclear egress rely on imaging-based technologies such as confocal and electron microscopy. These techniques are labor-intensive, limited by the number of cells that can be examined, and may not accurately represent the entire population, generating a potential bias during data interpretation. To overcome these problems, we have developed a flow cytometry–based method to measure HSV-1 nuclear egress that we termed FLARE (FLow cytometry–based Assay of nucleaR Egress). This assay uses a double fluorescent reporter system, utilizing HSV-1-tdTomato to identify infected cells and an Alexa Fluor-488-conjugated, capsid-specific antibody to detect cytosolic capsids, thereby distinguishing infected cells with nuclear egress from those without it. This assay provides more quantitative results than traditional methods, enables large-scale high throughput, and can be adapted for use with other herpesviruses.Key features• Quantification of HSV-1 nuclear egress by flow cytometry using a double fluorescent reporter system.• The assay is suitable for large-scale high-throughput screens, e.g., CRISPR/Cas9.• The assay can be adapted for use with other herpesviruses, provided a mature capsid-specific antibody is available.

Traditional methods for studying protein–protein interactions often lack the resolution to quantitatively distinguish distinct oligomeric states, particularly for membrane proteins within their native lipid environments. To address this limitation, we developed SiMPull-POP (single-molecule pull-down polymeric nanodisc photobleaching), a single-molecule technique designed to quantify membrane protein oligomerization with high sensitivity and in a near-native context. The goal of SiMPull-POP is to enable precise, quantitative analysis of membrane protein assembly by preserving native lipid interactions using diisobutylene maleic acid (DIBMA) to form nanodiscs. Unlike ensemble methods such as co-immunoprecipitation or FRET, which average out heterogeneous populations, SiMPull-POP uses photobleaching to resolve monomeric, dimeric, and higher-order oligomeric states at the single-molecule level. We validated SiMPull-POP using several model systems. A truncated, single-pass transmembrane protein (Omp25) appeared primarily monomeric, while a membrane-tethered FKBP protein exhibited ligand-dependent dimerization upon addition of the AP ligand. Applying SiMPull-POP to EphA2, a receptor tyrosine kinase, we found it to be mostly monomeric in the absence of its ligand, Ephrin-A1, and shifting toward higher-order oligomers upon ligand binding. To explore factors influencing ligand-independent assembly, we modulated membrane cholesterol content. Reducing cholesterol induced spontaneous EphA2 oligomerization, indicating that cholesterol suppresses receptor self-association. Overall, SiMPull-POP offers significant advantages over conventional techniques by enabling quantitative, single-molecule resolution of membrane protein complexes in a native-like environment. This approach provides critical insights into how membrane properties and external stimuli regulate protein assembly, supporting broader efforts to understand membrane protein function in both normal and disease states.Key features• Precise determination of membrane protein stoichiometry (e.g., monomer, dimer, oligomer) by directly counting photobleaching steps, overcoming the averaging limitations of bulk assays.• By incorporating membrane proteins into DIBMA lipid particles (DIBMALPs), this preserves native lipid interactions, offering a more physiologically relevant context for studying protein assembly.• Sensitively detects ligand-induced or membrane property-driven changes in oligomerization, making it a powerful tool for investigating both constitutive and regulated protein interactions.

Conventional Schlieren optics equipment typically operates on a large optical table, which is inconvenient for imaging small samples or thin layers of transparent materials. We describe an imaging device based on Schlieren optics, aided by a slight shift in light reflected from two surfaces. The device is designed to place the sample between a thick concave mirror and a camera next to a point-light source located at the spherical origin of the concave mirror. The compact device is portable and convenient. It is similarly capable of sensitively detecting patterns in gaseous or liquid media created by a density gradient when the optical effect is too subtle to be detectable by regular cameras and scanners. The new device is particularly suitable for detecting translucent samples, including thin fluid films on the order of micrometers, tissue slices, and other biological samples. We show two examples of how our device can be applied to imaging biological samples. The first compares images acquired using several techniques of a bacterial swarm spread over an agar plate; the second is a set of images of human cells grown on a tissue culture plate.Key features• The protocol presents the design of a compact Schlieren optics device (CSOD), with image boundaries enhanced by a slight shift in two overlapping, virtual images.• The CSOD captures high-resolution images of a transparent medium with variation in thickness or index of refraction.• The CSOD can detect transparent samples with thickness in the order of 1 µm; it is simple to build, user-friendly, and portable.• As a cheaper and portable complement to a phase contrast microscope, the device can image large samples more conveniently.

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.Key features• Subcutaneous transplantation of fetal kidneys into recipient fetuses minimizes surgical invasiveness and significantly improves fetal survival.• Natural delivery enables pups to nurse from the dam, allowing extended postnatal observation.• Use of green fluorescent protein (GFP)-expressing donor tissue permits real-time visualization of graft location and growth.• The protocol is readily adaptable for xenotransplantation and studies of immunological tolerance during fetal development.

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.Key features• First report to constitutively express a fluorescent label in hTSCs by applying a CRISPR/Cas9 knock-in approach and an AAVS1 safe harbor locus.• Provides an efficient tool to facilitate the study of human trophoblast development and function, particularly in heterologous co-culture systems.• Offers an approach for developing clinical gene therapy–related plasmid constructs that allow insertion of therapeutic genes without associated disruption of essential genes.• Widely applicable approach to label other human cell lines.

Most viruses extensively remodel their host cells to establish productive infection. Visualization of virus-induced cellular remodeling by electron microscopy (EM) has been revolutionized in recent years by advances in cryo-focused ion beam (cryo-FIB) milling paired with cryo-electron tomography (cryo-ET). As cryo-FIB/ET becomes more widely available, there is a need for beginner-friendly guides to optimize the preparation of virus-infected mammalian cells on EM grids. Here, we provide an in-house protocol for new users for preparing samples of cells infected with herpes simplex virus 1 (HSV-1) for cryo-FIB/ET. This protocol guides users in how to seed infected cells onto grids, blot, and plunge-freeze grids using basic, manual equipment. It also provides tips on how to screen and prioritize grids for efficient milling and data collection.Key features• A beginner-friendly protocol for users without access to a cryo-EM core/suite at their institution that utilizes basic equipment.• This protocol focuses on optimizing cell seeding and blotting to yield grids with thin ice and evenly distributed cells.• Grids prepared using this protocol can be used for focused ion beam milling.

Obesity is a risk factor for many diseases. The 3T3-L1 cell line is often used to obtain mature adipocytes, but these lack the inflammatory phenotype observed in obesity. Using a cocktail of cytokines that mimics the secretome of macrophages found in the inflammatory adipose tissue, we developed a protocol for obtaining mature inflammatory adipocytes. This model was validated at gene (RT-qPCR) and protein levels (multiplex adipokine array) as we found a decrease of adipogenic markers (C/EBPα, PPARУ, adiponectin, and CD36) and an increase of pro-inflammatory cytokines (IL-6, IL-1β, CXCL1, CXCL10, TNF-α, ICAM-1, and lipocalin-2). We provide a relevant in vitro model for studying the impact of low-grade chronic inflammation caused by obesity and its downstream effects on metabolic disorders and tumor microenvironments. Key features • Currently available protocols of adipocyte differentiation are not relevant for studying obesity in vitro. • We developed a simple and reproducible method to generate inflammatory adipocytes in vitro using a cytokine cocktail. • Gene expression analysis (qPCR) confirms the downregulation of adipogenic and protective markers (e.g., adiponectin, CD36) and the upregulation of pro-inflammatory cytokines (e.g., IL-6, IL-1β, TNF-α). • Adipokine array reveals decreased secretion of anti-inflammatory molecules (adiponectin, IGFBPs, FGF-21, HGF) and increased release of pro-inflammatory adipokines (serpin E1, IGF-1, lipocalin-2, IL-6, ICAM-1). • This protocol provides a relevant and versatile method for investigating obesity-related inflammation and its role in disease progression.