Cell Biology

Time-lapse fluorescence microscopy is a relevant technique to visualize biological events in living samples. Maintaining cell survival by limiting light-induced cellular stress is challenging and requires protocol development and image acquisition optimization. Here, we provide a guide by considering the quartet sample, probe, instrument, and image processing to obtain appropriate resolutions and information for live cell fluorescence imaging. The pleural mesothelial cell line H28, an adherent cell line that is easy to seed, was used to develop innovative advanced light microscopy strategies. The chosen red and near-infrared probes, capable of passively penetrating through the cell plasma membrane, are particularly suitable because their stimulation from 600 to 800 nm induces less cytotoxicity. The labeling protocol describes the concentration, time, and incubation conditions of the probes and associated adjustments for multi-labeling. To limit phototoxicity, acquisition parameters in advanced confocal laser scanning microscopy with a white laser are determined. Light power must be adjusted and minimized at red wavelengths for reduced irradiance (including a 775 nm depletion laser for STED nanoscopy), in simultaneous mode with hybrid detectors and combined with the fast FLIM module. These excellent conditions allow us to follow cellular and intracellular dynamics for a few minutes to several hours while maintaining good spatial and temporal resolutions. Lifetime analysis in lifetime imaging (modification of the lifetime depending on environmental conditions), lifetime dye unmixing (separation with respect to the lifetime value for the spectrally closed dye), and lifetime denoising (improvement of image quality) provide flexibility for multiplexing experiments.

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.

Bone repair is a complex regenerative process relying on skeletal stem/progenitor cells (SSPCs) recruited predominantly from the periosteum. Activation and differentiation of periosteal SSPCs occur in a heterogeneous environment, raising the need for single cell/nucleus transcriptomics to decipher the response of the periosteum to injury. Enzymatic cell dissociation can induce a stress response affecting the transcriptome and lead to overrepresentation of certain cell types (i.e., immune and endothelial cells) and low coverage of other cell types of interest. To counteract these limitations, we optimized a protocol to isolate nuclei directly from the intact periosteum and from the fracture callus to perform single-nucleus RNA sequencing. This protocol is adapted for fresh murine periosteum, fracture callus, and frozen human periosteum. Nuclei are isolated using mechanical extraction combined with fluorescence-based nuclei sorting to obtain high-quality nucleus suspensions. This protocol allows the capture of the full diversity of cell types in the periosteum and fracture environment to better reflect the in vivo tissue composition.

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.

Communication between motor neurons and muscles is established by specialized synaptic connections known as neuromuscular junctions (NMJs). Altered morphology or numbers of NMJs in the developing muscles can indicate a disease phenotype. The distribution and count of NMJs have been studied in the context of several developmental disorders in different model organisms, including zebrafish. While most of these studies involved manual counting of NMJs, a few of them employed image analysis software for automated quantification. However, these studies were primarily restricted to the trunk musculature of zebrafish. These trunk muscles have a simple and reiterated anatomy, but the cranial musculoskeletal system is much more complex. Here, we describe a stepwise protocol for the visualization and quantification of NMJs in the ventral cranial muscles of zebrafish larvae. We have used a combination of existing ImageJ plugins to develop this methodology, aiming for reproducibility and precision. The protocol allows us to analyze a specific set of cranial muscles by choosing an area of interest. Using background subtraction, pixel intensity thresholding, and watershed algorithm, the images are segmented. The binary images are then used for NMJ quantification using the Analyze Particles tool. This protocol is cost-effective because, unlike other licensed image analyzers, ImageJ is open-source and available free of cost.

Gap junctions are transmembrane protein channels that enable the exchange of small molecules such as ions, second messengers, and metabolites between adjacent cells. Gap junctions are found in various mammalian organs, including skin, endothelium, liver, pancreas, muscle, and central nervous system (CNS). In the CNS, they mediate coupling between neural cells including glial cells, and the resulting panglial networks are vital for brain homeostasis. Tracers of sufficiently small molecular mass can diffuse across gap junctions and are used to visualize the extent of cell-to-cell coupling in situ by delivering them to a single cell through sharp electrodes or patch-clamp micropipettes. Here, we describe a protocol for pre-labeling and identification of astrocytes in acute mouse forebrain slices using Sulforhodamine 101 (SR101). Fluorescent cells can then be targeted for whole-cell patch-clamp, which allows for further confirmation of astroglial identity by assessing their electrophysiological properties, as well as for passive dialysis with a tracer such as biocytin. Slices can then be subjected to chemical fixation and immunostaining to detect dye-coupled networks. This protocol provides a method for the identification of astrocytes in live tissue through SR101 labeling. Alternatively, transgenic reporter mice can also be used to identify astrocytes. While we illustrate the use of this protocol for the study of glial networks in the mouse brain, the general principles are applicable to other species, tissues, and cell types.

In nature, filamentous fungi interact with plants. These fungi are characterized by rapid growth in numerous substrates and under minimal nutrient requirements. Investigating the interaction of these fungi with their plant hosts under controlled conditions is of importance for many researchers aiming to proceed with molecular or microscopical investigations of their favorite plant–fungus interaction system. The speed of growth of these fungi complicates transferring plant–fungal interaction systems in laboratory conditions. The issue is more complicated when monoxenic conditions are desired, to ensure that only two members (a fungus and a plant) are present in the system under study. Here, two simple closed systems for investigating plant–filamentous fungi associations under laboratory, monoxenic conditions are described, along with their limitations. The plant and fungal growth conditions, methods for sampling, staining, sectioning, and subsequent microscopical imaging of colonized plant tissues with affordable, common laboratory tools are described.

Calcium-permeable AMPA receptors (CP-AMPARs) and kainate receptors (CP-KARs) play crucial roles in synaptic plasticity and are implicated in various neurological processes. Current methods for identifying neurons expressing these receptors, such as electrophysiological recordings and immunostaining, have limitations in throughput or inability to distinguish functional receptors. This protocol describes a novel approach for the vital identification of neurons containing CP-AMPARs and CP-KARs using calcium imaging. The method involves loading neurons with Fura-2 AM, a calcium-sensitive fluorescent probe, KCl application to identify all neurons, and further addition of specific AMPAR agonists (e.g., 5-fluorowillardiine) in the presence of voltage-gated calcium channel blockers and NMDAR/KAR antagonists to identify CP-AMPAR-containing neurons. CP-KAR-containing neurons are identified using domoic acid applications in the presence and absence of NASPM (a CP-AMPAR antagonist). This technique offers several advantages over existing methods, including the ability to assess large neuronal populations simultaneously, distinguish between different receptor types, and provide functional information about CP-AMPAR and CP-KAR expression in living neurons, making it a valuable tool for studying synaptic plasticity and neurological disorders.

The existence and functional relevance of DNA and RNA G-quadruplexes (G4s) in human cells is now beyond debate, but how did we reach such a level of confidence? Thanks to a panoply of molecular tools and techniques that are now routinely implemented in wet labs. Among them, G4 imaging ranks high because of its reliability and practical convenience, which now makes cellular G4 detection quick and easy; also, because this technique is sensitive and responsive to any G4 modulations in cells, which thus allows gaining precious insights into G4 biology. Herein, we briefly explain what a G4 is and how they can be visualized in human cells; then, we present the strategy we have been developing for several years now for in situ click G4 imaging, which relies on the use of biomimetic G4 ligands referred to as TASQs (for template-assembled synthetic G-quartets) and is far more straightforward and modular than classically used immunodetection methods. We thus show why and how to illuminate G4s with TASQs and provide a detailed, step-by-step methodology (including the preparation of the materials, the methodology per se, and a series of notes to address any possible pitfalls that may arise during the experiments) to make G4 imaging ever easier to operate.

Cell viability and cytotoxicity assays are commonly used to investigate protein function and to evaluate drug efficacy in cancer and other disease models. Cytotoxicity is the measure of dead or damaged cells and is often quantified using assays based on cellular characteristics such as membrane integrity or mitochondrial metabolism. However, these assays are typically limited to endpoint analysis and lack emulation of physiological conditions. The IncuCyte Live and Dead Cell assay described here leverages common cell permeability methodologies but uses fluorescence microscopy channels to image both live and dead cells over time and phase microscopy channels to measure confluency. Cytotox green reagent is a cell membrane–impermeable dye that can only be taken up by cells with poor cell membrane integrity. NucLight rapid red dye is a cell membrane–permeable nuclear dye that can be taken up by all cells. Based on dye uptake and fluorescence intensity, the IncuCyte software can be used to analyze images for live and dead cell detection and quantification. Phase microscopy is used to determine confluency and can be further quantified using the IncuCyte software. We provide an application of this assay, using it to calculate IC₅₀ and EC₅₀ values for the assessment of drug efficacy.