细胞生物学

The cAMP-dependent protein kinase (PKA) is one of the most extensively distributed kinases among intracellular signal cascades, with a pivotal role in the regulation of various processes, including the capacitation of sperm cells. Traditional assessments of PKA activity rely on the utilization of [γ-³²P] ATP and the Kemptide peptide as a substrate. This strategy presents several major drawbacks, including high costs and health risks derived from the manipulation of radioactive isotopes. In this work, we introduce an enhanced non-radioactive assay to quantify PKA activity, termed kinase mobility shift assay (KiMSA), based on the use of a fluorescent-labeled Kemptide (Kemptide-FITC). Once the kinase reaction is terminated, the products can be easily resolved through electrophoresis on an agarose gel and quantified by fluorescence densitometry. We show that KiMSA is suitable for isolated PKA as well as for the enzyme in cell extracts. In addition, it enables quantification of PKA activity during the progression of mouse sperm capacitation. Furthermore, the assay enables monitoring the inhibition of PKA with pharmacological inhibitors in live cells. Therefore, the experimental and optimal assay conditions are set so that KiMSA can be used to assess in vitro as well as in vivo PKA activity in sperm cells. Finally, this method allows for measurement of cAMP concentrations, rendering a versatile technique for the study of cAMP/PKA pathways.

The subcellular localization of RNA plays a critical role in various biological processes, including development and stress response. Proximity labeling eases the detection of localized transcripts and protein enrichment compared to previous techniques that rely on biochemical isolation of subcellular structures. The rapid reaction and small labeling radius of APEX2 make it an attractive alternative to other proximity labeling approaches, such as BioID. However, we found that standard protocols for APEX proximity labeling fail in human induced pluripotent stem cells. Moreover, standard protocols yield heterogeneous labeling of biomolecules across single cells in MCF10A breast epithelial cells. Our results indicate that low biotin permeability in these cell lines is the main cause for failed or inefficient labeling. This protocol outlines improved labeling by combining the rapid hydrogen peroxide-driven APEX2 reaction with the addition of a mild detergent during biotin incubation. This adaptation leads to efficient proximity labeling in hiPSCs and more homogeneous biotinylation across single cells in MCF10As. The adapted protocol extends the use of APEX2 proximity labeling to cell lines with poor biotin permeability.

Zika virus (ZIKV), an arthropod-borne orthoflavivirus, has emerged as a global health concern due to its ability to cause severe fetal neurological disorders, leading to the congenital Zika syndrome (CZS) in neonates. Vertical transmission during pregnancy can alter neural progenitor cell (NPC) proliferation and differentiation and induce apoptosis, leading to microcephaly and other neurodevelopmental abnormalities. While mammalian models have been used to study the impact of ZIKV on NPC behavior, limitations such as high costs, dedicated time, and ethical constraints have fostered the exploration of alternative systems. The zebrafish embryo constitutes an advantageous in vivo model for studying ZIKV neuropathogenesis. Indeed, ZIKV infection phenocopies several features of the CZS while sharing a conserved neuroanatomical layout and offering genetic plasticity and unique accessibility to the infected brain compared to mammals. Here, we describe a protocol for characterizing ZIKV-induced defects of NPCs in this zebrafish model, relying on whole animal flow cytometry.

Epithelial tissues form barriers to the flow of ions, nutrients, waste products, bacteria, and viruses. The conventional electrophysiology measurement of transepithelial resistance (TEER/TER) can quantify epithelial barrier integrity, but does not capture all the electrical behavior of the tissue or provide insight into membrane-specific properties. Electrochemical impedance spectroscopy, in addition to measurement of TER, enables measurement of transepithelial capacitance (TEC) and a ratio of electrical time constants for the tissue, which we term the membrane ratio. This protocol describes how to perform galvanostatic electrochemical impedance spectroscopy on epithelia using commercially available cell culture inserts and chambers, detailing the apparatus, electrical signal, fitting technique, and error quantification. The measurement can be performed in under 1 min on commercially available cell culture inserts and electrophysiology chambers using instrumentation capable of galvanostatic sinusoidal signal processing (4 μA amplitude, 2 Hz to 50 kHz). All fits to the model have less than 10 Ω mean absolute error, revealing repeatable values distinct for each cell type. On representative retinal pigment (n = 3) and bronchiolar epithelial samples (n = 4), TER measurements were 500–667 Ω·cm² and 955–1,034 Ω·cm² (within the expected range), TEC measurements were 3.65–4.10 μF/cm² and 1.07–1.10 μF/cm², and membrane ratio measurements were 18–22 and 1.9–2.2, respectively.

Human intestinal barrier function is crucial for health. Beneficial microbes, such as commensal gut bacteria and probiotics, are known to contribute to the regulation of this barrier function. Interactions between bacteria and human intestinal cells can be analyzed by co-culturing bacteria with mammalian cells in vitro. Here, we describe a method to assess the effect of individual bacterial strains on intestinal barrier function using automated transepithelial electrical resistance (TEER) measurements. Caco-2 cells are used as a model of the intestinal epithelium, as these cells spontaneously differentiate into small intestinal epithelial-like cells characterized by tight junctions between adjacent cells. These cells are seeded on polyester filter inserts and cultured for 17 days to form a differentiated monolayer prior to the co-culture experiment. Bacteria are grown on agar, and a single colony is used to prepare a liquid culture in bacterial broth appropriate for the bacteria of interest. On the day of the co-culture experiment, the bacterial culture is resuspended in cell culture medium at the desired concentration. Inserts are transferred to cellZscope cell modules to enable automated TEER measurements, and the medium in the insert is replaced with cell culture medium containing the bacteria of interest. This method allows for intestinal tight junction barrier function to be assessed non-invasively and in real-time in response to probiotics. The use of the automated cellZscope system eliminates the need for labor-intensive manual TEER measurements, which reduces the variability in data that results from human handling and temperature changes that occur when cells are removed from the incubator.

The Seahorse 96 XF Analyzer (Agilent Technologies, Santa Clara, CA, USA) has been an effective tool in non-invasively measuring mitochondrial function for the past decade. It is a high-throughput respirometer that is considered the “gold standard” for quantifying mitochondrial function and bioenergetics in cells. Peripheral blood mononuclear cells (PBMCs) play a selective role in immune system responses and are key components of human immunity. Recent studies have suggested that these cell populations provide an overview of systemic changes within the body and therefore provide a source of sensitive biomarkers. Assessing mitochondrial function in PBMCs has been shown to provide an indication of metabolic stress associated with diseases such as diabetes and neurodegenerative conditions such as Alzheimer’s disease. In this protocol, we use two adhesive compounds, Poly-D-Lysine (PDL) and Poly-L-Lysine (PLL), at 50 μg/mL each per well, to immobilize PBMCs to a specialized Seahorse microplate to perform mitochondrial stress assay using the Seahorse Analyzer. We compared six cell densities of PBMCs to identify the optimal cell density for use in Seahorse Mito Stress analysis. This protocol includes the immobilization of freshly isolated PBM cells into a Seahorse microplate, hydration and calibration of the sensor cartridge, cell seeding, running the Seahorse Analyzer for the Mito Stress test, and simple data analysis to compare the effectiveness of PLL and PDL as the coating agent for PBMCs. The data analysis indicates that there is no statistical difference between PLL and PDL.

Plastic pollution presents a looming danger to the environment and virtually all life on planet Earth. Especially pernicious are nanoplastics (NPs), which are plastic fragments with dimensions ≤1 μm. Conventional detection methods are ineffective for NPs, while their high specific surface area renders them efficient carriers of toxic substances; additionally, they may even be inherently toxic. Although NP waste chiefly arises from environmental weathering of larger plastic fragments, most published studies employed manufactured pristine NPs of uniform size and shape. Furthermore, almost all NP effects were studied using polystyrene (PS) as a convenient model material, despite PS accounting for <6% of all plastic pollution. There is thus an urgent need to expand investigations of environmental NP pollution and effects on biota. The present work provides a comprehensive roadmap for studying the effects of “real-world” NP pollution on living systems, using, for example, lung alveolar epithelial cells on which such NPs deposit by breathing ambient air. Herein, we describe detailed in-house methods to fabricate various NPs that are weathered with UV light and O₃ gas exposure to more closely mimic real environmental NPs. We also illustrate a simple and straightforward bioelectrical method for assessing passive and active ion transport properties of primary rat lung alveolar epithelial cell monolayers as a model for the distal mammalian lung exposed to one of the generated NPs. This protocol allows researchers to rapidly and more accurately assess the biological impact of various simulated environmental NPs on a vulnerable air–blood barrier in the lung.

In vitro lymphocyte proliferation assays are essential for assessing immune responses and antiproliferative drug efficacy. Such assays rely on antigen presentation or mitogen stimulation, with performance determined by reagent concentration and incubation time. Although splenocytes are often used, peripheral blood mononuclear cells (PBMCs) offer more accessible and practical sampling. However, a streamlined protocol for porcine PBMCs proliferation with robust batch analysis has been lacking. We therefore developed a detailed workflow for inducing proliferation in cryopreserved porcine PBMCs using 5 μg/mL concanavalin A (ConA). The protocol covers cell isolation, cryopreservation, ConA stimulation, CD4+ T-cell staining, flow cytometry acquisition and gating on an Attune NxT instrument, and batch analysis with FCS Express^TM 7.18. This approach yielded 78.9% viable cells, of which 33.8% were CD4+ lymphocytes. Moreover, 93.9% (n = 216) of cells proliferated, yielding up to nine cell generations. Batch analysis in FCS Express^TM enhanced the accuracy and interpretation of proliferation metrics. This validated protocol provides a reliable framework for generating consistent proliferation data in porcine immunology studies.

Microglial cells are crucial patrolling immune cells in the brain and pivotal contributors to neuroinflammation during pathogenic or degenerative stress. Microglia exhibit a heterogeneous "dendrite-like" dense morphology that is subject to change depending on inflammatory status. Understanding the association between microglial morphology, reactivity, and neuropathology is key to informing treatment design in diverse neurodegenerative conditions from inherited encephalopathies to traumatic brain injuries. However, existing protocols for microglial morphology analyses lack standardization and are too complex and time-consuming for widescale adoption. Here, we describe a customized pipeline to quantitatively assess intricate microglial architecture in three dimensions under various conditions. This user-friendly workflow, comprising standard immunofluorescence staining, built-in functions of standard microscopy image analysis software, and custom Python scripts for data analysis, allows the measurement of important morphological parameters such as soma and dendrite volumes and branching levels for users of all skill levels. Overall, this protocol aims to simplify the quantification of the continuum of microglial pathogenic morphologies in biological and pharmacological studies, toward standardization of microglial morphometrics and improved inter-study comparability.

In response to DNA-damaging physical or chemical agents, the DNA damage repair (DDR) pathway is activated in eukaryotic cells. In the radiobiology field, it is important to assess the DNA damage effect of a certain irradiation regime on cancer cells and compare it to the effect on non-transformed cells exposed to identical conditions. The first step in the DNA repair mechanism consists of the attachment of proteins such as the phosphorylated histone γ-H2AX (p-γ-H2AX) to DNA double-strand breaks (DSB) in the nucleus, which leads to the formation of repairing foci. Therefore, imaging methods were established to evaluate the presence of foci inside the nucleus after exposure to DNA-damaging agents. This approach is superior in sensitivity to other methods, such as the comet assay or the pulsed-field gel electrophoresis (PFGE), that allow direct detection of cleaved DNA fragments. These electrophoresis-based methods require high ionizing radiation dosages and are difficult to reproduce compared to imaging-based assays. Conventionally, the number of foci is determined visually, with limited accuracy and throughput. Here, by exploring the effect of laser-plasma accelerated electrons FLASH irradiation on cancer cells, we describe an image cytometry protocol for the quantification of foci with increased throughput, upon large areas, with increased precision and sample-to-sample consistency. It consists of the automatic scanning of fluorescently labeled cells and using a gating strategy similar to flow cytometry to discriminate cells in co-culture based on nuclei elongation properties, followed by automatic quantification of foci number and statistical analysis. The protocol can be used to monitor the kinetics of DNA repair by quantification of p-γ-H2AX at different time points post-exposure or by quantification of other DNA repair proteins that form foci at the DNA DSB sites. Also, the protocol can be used for quantifying the response to chemical agents targeting DNA. This protocol can be performed on any type of cancer cells, and our gating strategy to discriminate cells in co-culture can also be used in other research applications.