Cell Biology

Antibody therapeutics have demonstrated transformative impacts on improving the quality of life of millions of patients, whereas advances in antibody discovery technologies have imposed a significant production challenge for the generation of a large diversity of therapeutic antibody candidates. A demand for the rapid production of dozens of purified antibodies in 10-mg quantities is entailed for functional screening and molecular assessment studies. Here, we present a robust semi-automated production protocol that bridges the gap between miniaturized high-throughput screenings and conventional custom-scale workflows. This methodology and workflow utilize a simple high-titer transient Chinese hamster ovary (CHO) cell host–CHO4Tx^® expression system, a procedure of magnetic protein-A bead in-culture antibody capturing, and a semi-automated purification process with the GenScript AmMag^TM SA Plus system. This production protocol has been proven to be robust and valuable for the routine production of dozens of antibody constructs per week in sufficient quality and quantity for cell-based and biophysical studies.

Understanding epithelial barrier function is essential for studying both its normal physiology and its role in disease, yet choosing an appropriate experimental model remains challenging. Animal models are commonly used but often suffer from interspecies differences that limit translational relevance. Human-derived cell lines offer a more suitable alternative, although establishing them often requires immortalisation strategies that involve overexpression of oncogenes, which can introduce phenotypic and functional changes. In contrast, primary cells, such as human small airway epithelial cells (HSAECs), provide a more physiologically accurate model. A critical aspect of replicating the native respiratory environment is maintaining continuous air exposure, which can be achieved through air–liquid interface (ALI) culture. This protocol provides a unified, step-by-step workflow for cultivating primary HSAECs under ALI conditions, covering the entire process from initial recovery after cryopreservation to the formation of a barrier-like layer. The protocol incorporates non-invasive methods such as transepithelial electrical resistance (TEER) measurements to monitor its integrity. While individual elements of this workflow have been described separately in different studies, a consolidated version encompassing the full workflow has not been widely available. This resource is intended for researchers with limited experience in airway epithelial culture and offers practical, clear guidance through each step of the process.

Fluorescence in situ hybridization (FISH) is a cytological method used to visualize specific oligonucleotide sequences within the cell. This method relies on the specific binding of a fluorescence-tagged probe, a short stretch of single-stranded polynucleotide, to its complementary sequence in the DNA or RNA, forming stable double-stranded hybrids. Fluorochromes, such as fluorescein, Alexa Fluor, cyanine dyes, or rhodamine, are attached to these probes to help in detecting their presence within the cell. Based on sequence complementarity, FISH allows for the visualization of the DNA or RNA with which they have hybridized. The distribution of these fluorochrome-tagged probes can be observed under a fluorescence or confocal microscope. The oligo(dT) FISH technique specifically utilizes a fluorochrome-tagged stretch of 40–50 thymidine (T) oligonucleotides that binds to the poly(A) tails of mature mRNAs within the cell. Newly transcribing pre-mRNAs and certain non-coding RNAs may not have poly(A) tails and therefore cannot be detected by this method. This step-by-step protocol outlines the oligo(dT) FISH technique for visualizing the cellular distribution of polyadenylated mRNAs in the tissues of Drosophila and other related model organisms.

The mammalian central circadian clock resides in the suprachiasmatic nucleus (SCN) of the hypothalamus in the brain and is responsible for coordinating daily rhythms of biological processes spanning from gene expression to behavior. Light, the primary environmental zeitgeber, entrains the SCN via melanopsin-expressing intrinsically photosensitive retinal ganglion cells that project through the retino-hypothalamic tract. Altered circadian rhythms are common in individuals diagnosed with neurodevelopmental and neurodegenerative disorders, and often, associated with structural alterations of the SCN and impaired retinal input; importantly, these anomalies can be recapitulated in animal models. Here, we describe step-by-step protocols for quantitative histomorphometrical analysis of the SCN and the assessment of retinal–SCN connectivity, previously used in mouse models of neurodevelopmental and neurodegenerative disorders. These include measurement of the SCN area, perimeter, height and width using Nissl- or DAPI-stained coronal sections, as well as densitometric and plot profile analyses of cholera toxin β-subunit–labeled retinal projections using Axiovision or Fiji/ImageJ. The protocols incorporate standardized region-of-interest, measurements by masked observers, and consistent scaling procedures to enhance reproducibility. These methods provide a rigorous framework for detecting structural anomalies and connectivity defects in the circadian system and can be broadly applied to other experimental models of circadian dysfunction.

Postnatal mouse retinal vascular development is a widely used model for studying retinal vascular diseases and evaluating candidate therapies. This is particularly relevant for inherited disorders such as familial exudative vitreoretinopathy (FEVR), in which impaired vascular growth and organization are central to disease pathogenesis. Numerous approaches have been used to assess retinal vasculature in mouse flat mounts, ranging from qualitative descriptions to limited quantitative measurements of vascular growth. However, phenotypic variability across genetic models, including different models of FEVR, complicates comparisons and underscores the need for standardized, comprehensive multi-parameter analyses that are suitable for rapid and cost-effective screening studies. We describe a standardized morphometric protocol using ImageJ software to quantitatively analyze mouse retinal vasculature in a reproducible manner. The protocol begins with measurement of areas of vascular disorganization (meshes) as well as total vascular and retinal area. Two defined regions in the peripheral and midperipheral retina are then selected to quantify cell clusters, followed by image processing, binarization, and skeletonization. From these processed images, vascular density, branch number, branch length and thickness, junction number, triple points, and box-counting fractal dimension and lacunarity are quantified. Overall, this protocol provides a rapid, cost-effective, and standardized framework for quantifying retinal vascular phenotypes across diverse mouse models. By capturing multiple structural features and accommodating phenotypic variability, it is well-suited for comparative studies and therapeutic screening in retinal vascular disease.

The activity of chloroplast ATP synthase (CF_oCF₁) is precisely regulated through a thioredoxin (Trx)-mediated dithiol/disulfide reaction in response to varying light conditions. This regulatory mechanism is further controlled by ΔpH formation across the thylakoid membrane. To better understand this complicating regulatory function of CF_oCF₁, a method is required to evaluate the extent of CF_oCF₁ reduction by Trx under controlled ΔpH conditions and to directly evaluate the redox state of CF_oCF₁. In this study, we present a simple in vitro procedure to assess the CF_oCF₁ reduction system using spinach thylakoids. The method consists of three key steps: (A) simple preparation of intact thylakoids from spinach leaves; (B) reduction of CF_oCF₁ on the thylakoid membrane using recombinant Trx under light irradiation; and (C) in situ determination of the redox state of CF_oCF₁ by labeling thiol groups with a maleimide reagent followed by protein detection using western blotting. The redox state of CF_oCF₁ was determined by mobility shifts on non-reducing SDS-PAGE. This protocol provides a refined strategy for elucidating the regulatory mechanism controlling energy conversion by CF_oCF₁ under fluctuating photosynthetic conditions.

Conventional light microscopy is limited in resolution by the diffraction limit of light, restricting the visualization of the nanoscale organization of biomolecules. Expansion microscopy (ExM) has emerged as a powerful technique to overcome this barrier by physically expanding the specimen embedded in a swellable hydrogel without requiring specialized or high-cost imaging hardware. ExM is widely used in animal models, whereas its application to plant tissues has been challenging due to their multicellularity, in which each cell is encompassed by the rigid cell wall, which resists the expansion forces and prevents isotropic swelling. Here, we describe a robust and optimized ExM protocol specifically designed for Arabidopsis thaliana root tissues. This protocol details critical steps, including immunostaining, anchoring, gelation, denaturation, cell wall digestion, and expansion. Our method achieves an expansion factor of approximately 4.3×, enabling an effective lateral resolution of ~60 nm using a standard confocal microscope. We demonstrate the visualization of microtubules with preserved ultrastructure. This accessible protocol allows plant researchers to perform super-resolution imaging without specialized optical equipment, facilitating detailed structural analysis of plant cells.

Animal and human stem cell–derived three-dimensional models to study physio-pathological brain functioning are becoming a gold standard for in vitro electrophysiology, as they enable the recapitulation of complex network properties by accounting for spatial architectural features that better reflect in vivo conditions than simpler 2D models. Standard planar multielectrode arrays (MEAs), typically providing tens of recording electrodes, are commonly used to record activity from 2D neuronal cultures. However, when adapted for use with 3D models, planar 2D MEAs showed limited effectiveness. The main issues are limited specimen adhesion to the chip, a low number of sensing elements, inability to retrieve signals from within the tissue, and reduced perfusion and vitality of the tissue in contact with sensors. To overcome these limitations, a new generation of microchip-based 3D high-density MEAs (3D HD-MEA) has been developed and validated in recent years. This technological advancement has improved the sensing capabilities and the vitality of 3D models, providing a tool tailored to maximize their potential. Here, we present an optimized protocol for neural network activity recordings in 3D models (including acute slices, brain spheroids, and organoids) from various brain regions using 3D HD-MEAs. First, we summarize the critical steps for 1) obtaining viable acute slices from the mouse cerebellum, cortico-hippocampal circuit, and prefrontal cortex, 2) establishing efficient coupling of the slices with the chip, and 3) performing recordings and analyses. We then describe the main procedures required to obtain human and animal brain spheroids and neural organoids, as well as standardized routines to perform effective recordings and analyses. For each section, we highlight the crucial steps, identify tips for specific applications, and propose troubleshooting procedures. For example, the same type of preparation (e.g., acute slices) requires different adjustments when working with different brain areas. The specific information provided here is intended to assist researchers in their daily efforts to obtain efficient and reproducible functional recordings from 3D models by using the cutting-edge technique of 3D HD-MEA.

Larval zebrafish are often mounted laterally to ensure consistent anatomical positioning and to standardize imaging of body axes across early development. However, this conventional approach often tethers sample orientation to a single microscope configuration and limits optical accessibility. We present a mounting protocol for larval zebrafish that enables optical access from both dorsal and ventral orientations while preserving lateral sample position. This approach uses common laboratory consumables to establish a mounting platform that eliminates any need to remount samples between the use of upright and inverted microscopes. By establishing a hydrophobic seal, mounted embryos can be inverted with ease to access the sample from either orientation. A seamless transition here facilitates reliable identification and longitudinal tracking of the same biological region of interest across microscope configurations. This protocol is broadly applicable to live imaging experiments requiring flexibility in imaging geometry, minimal sample handling, and high reproducibility.

Neural tube closure is a critical process that transforms the neural plate, an open epithelial tissue, into the closed tube that serves as the structural basis of the central nervous system. Defects in this process are among the most common and severe developmental diseases in the human population, with failures in cranial closure accounting for approximately one-third of total defects. However, the cell and tissue mechanisms that drive cranial closure remain opaque relative to the better studied process of spinal closure, in large part due to the unique challenges in characterizing cranial tissues. Here, we present protocols for quantifying cell dynamics and tissue-level remodeling events that enable highly spatiotemporally resolved investigations of the causes of cranial closure defects in mouse embryos. These include brightfield morphometric approaches, fluorescent staining and confocal imaging, and quantitative pipelines to analyze these image-based datasets. At the conclusion of these approaches, users will be able to quantify several parameters of overall tissue shape in the cranial neural tissues and produce rich quantitative datasets about cell-level parameters, particularly apical cell area. These can be used to identify correlative and causative differences between mutants and control embryos. Given their flexibility, many of these approaches can be generalized to other tissue morphogenetic contexts.

Three-dimensional immunohistochemistry (3D-IHC) shows the organization of molecular assemblies in the context of tissue architecture. Deep and rapid antibody penetration into 3D tissues and highly sensitive detection are crucial for high-throughput analysis of 3D-IHC imaging. Here, we provide a detailed protocol for a nanobody (nAb)-based 3D-IHC technique, namely POD-nAb/FT-GO 3D-IHC, for high-speed and high-sensitivity detection of targets within 1-mm-thick mouse brain tissues. Peroxidase-fused nAb (POD-nAb) is a genetically encoded recombinant antibody, which consists of a camelid nAb and a variant of horseradish peroxidase, and fluorochromized tyramide-glucose oxidase (FT-GO) is a fluorescent tyramide signal amplification (TSA) system. POD-nAb/FT-GO 3D-IHC incorporates three main components: 1) tissue permeabilization, 2) POD-nAb binding, and 3) 3D-TSA reaction with FT-GO. POD-nAbs enhance signal penetration depth and allow for highly sensitive detection when combined with FT-GO signal amplification. By using the 3D-IHC protocol provided herein, we can visualize target molecules in mouse brain tissues of 1-mm thickness with drastic signal enhancement within three days. This protocol for POD-nAb/FT-GO 3D-IHC could facilitate structural and molecular interrogation of 3D tissues.

Human mononuclear cells derived from peripheral blood and bone marrow are valuable resources for the study of hematological malignancies, including acute myeloid leukemia (AML) and chronic myelomonocytic leukemia (CMML). Cryopreservation enables long-term storage of patient samples for downstream assays; while thawing protocols have been described, subsequent recovery of viable cells after thawing can be challenging, particularly for fragile blast and monocyte populations. Here, we describe a reliable protocol for thawing cryopreserved AML and CMML mononuclear cells designed to preserve post-thaw viability, recovery, and functional integrity. The method incorporates controlled dilution of cells out of cryoprotectant with anticoagulant-supplemented thaw buffer, DNase I treatment, and gentle resuspension steps. Using this approach, post-thaw viability consistently exceeded 80% with a mean recovery of 55.6% across samples. Recovered cells retained functional capacity, as demonstrated by colony-forming assays, and maintained immunophenotypic characteristics by flow cytometry. This protocol provides a robust and reproducible method for the recovery of cryopreserved AML and CMML mononuclear cells and may be broadly applicable to other fragile or monocyte-rich patient-derived hematopoietic samples.