细胞生物学

Local mRNA translation in axons is crucial for the maintenance of neuronal function and homeostasis, particularly in processes such as axon guidance and synaptic plasticity, due to the long distance from axon terminals to the soma. Recent studies have shown that RNA granules can hitchhike on the surface of motile lysosomal vesicles, facilitating their transport within the axon. Accordingly, disruption of lysosomal vesicle trafficking in the axon, achieved by knocking out the lysosome–kinesin adaptor BLOC-one-related complex (BORC), decreases the levels of a subset of mRNAs in the axon. This depletion impairs the local translation of mitochondrial and ribosomal proteins, leading to mitochondrial dysfunction and axonal degeneration. Various techniques have been developed to visualize translation in cells, including translating RNA imaging by coat protein knock-off (TRICK), SunTag, and metabolic labeling using the fluorescent non-canonical amino acid tagging (FUNCAT) systems. Here, we describe a sensitive technique to detect newly synthesized proteins at subcellular resolution, the puromycin proximity ligation assay (Puro-PLA). Puromycin, a tRNA analog, incorporates into nascent polypeptide chains and can be detected with an anti-puromycin antibody. Coupling this method with the proximity ligation assay (PLA) allows for precise visualization of newly synthesized target proteins. In this article, we describe a step-by-step protocol for performing Puro-PLA in human induced pluripotent stem cell (iPSC)-derived neuronal cultures (i3Neurons), offering a powerful tool to study local protein synthesis in the axon. This tool can also be applied to rodent neurons in primary culture, enabling the investigation of axonal protein synthesis across species and disease models.

The reduction in intracellular neuronal chloride concentration is a crucial event during neurodevelopment that shifts GABAergic signaling from depolarizing to hyperpolarizing. Alterations in chloride homeostasis are implicated in numerous neurodevelopmental disorders, including autism spectrum disorder (ASD). Recent advancements in biosensor technology allow the simultaneous determination of intracellular chloride concentration of multiple neurons. Here, we describe an optimized protocol for the use of the ratiometric chloride sensor SuperClomeleon (SClm) in organotypic hippocampal slices. We record chloride levels as fluorescence responses of the SClm sensor using two-photon microscopy. We discuss how the SClm sensor can be effectively delivered to specific cell types using virus-mediated transduction and describe the calibration procedure to determine the chloride concentration from SClm sensor responses.

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.

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.

Fluorescence lifetime imaging microscopy (FLIM) is a highly valuable technique in the fluorescence microscopy toolbox because it is essentially independent of indicator concentrations. Conventional fluorescence microscopy analyzes changes in emission intensity. In contrast, FLIM assesses the fluorescence lifetime, which is defined as the time a fluorophore remains in an excited state before emitting a photon. This principle is advantageous in experiments where fluorophore concentrations are expected to change, e.g., due to changes in cell volume. FLIM, however, requires collecting a substantial number of photons to accurately fit distribution plots, which constrains its ability for dynamic imaging. This limitation has recently been overcome by rapidFLIM, which utilizes ultra-low dead-time photodetectors in conjunction with sophisticated rapid electronics. The resulting reduction in dead-time to the picosecond range greatly enhances the potential for achieving high spatio-temporal resolution. Here, we demonstrate the use of multi-photon-based rapidFLIM with the sodium indicator ION NaTRIUM Green-2 (ING-2) for the quantitative, dynamic determination of Na⁺ concentrations in neurons in acute rodent brain tissue slices. We describe the loading of the dye into neurons and present a procedure for its calibration in situ. We show that rapidFLIM not only allows the unbiased determination of baseline Na⁺ concentrations but also allows dynamic imaging of changes in intracellular Na⁺, e.g., induced by inhibition of cellular ATP production. Overall, rapidFLIM, with its greatly improved signal-to-noise ratio and higher spatio-temporal resolution, will also facilitate dynamic measurements using other FLIM probes, particularly those with a low quantum yield.

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.

Mitochondrial cristae, formed by folding the mitochondrial inner membrane (IM), are essential for cellular energy supply. However, the observation of the IM is challenging due to the limitations in spatiotemporal resolution offered by conventional microscopy and the absence of suitable in vitro probes specifically targeting the IM. Here, we describe a detailed imaging protocol for the mitochondrial inner membrane using the Si-rhodamine dye HBmito Crimson, which has excellent photophysical properties, to label live cells for imaging via stimulated emission depletion (STED) microscopy. This allows for STED imaging over more than 500 frames (approximately one hour), with a spatial resolution of 40 nm, enabling the observation of cristae dynamics during various mitochondrial processes. The protocol includes detailed steps for cell staining, image acquisition, image processing, and resolution analysis. Utilizing the superior resolution of STED microscopy, the structure and complex dynamic changes of cristae can be visualized.

Cryo-electron microscopy (cryo-EM) is a powerful technique capable of investigating samples in a hydrated state, compared to conventional high-vacuum electron microscopy that requires samples to be completely dry. During the drying process, numerous features and details may be lost due to damage caused by dehydration. Cryo-EM circumvents these problems by cryo-fixing the samples, thereby retaining the intact and original features of hydrated samples. This protocol describes a step-by-step cryo-scanning electron microscopy (cryo-SEM) experimental procedure with Chlorella sorokiniana as the subject. By employing filter paper as the sample substrate, we propose a simple and reliable method for cryo-fixation and freeze-fracture of Chlorella sorokiniana in water suspension. The advantage of using filter paper as a substrate lies in its ability to support a thin film of sample, enabling a cold knife to make a cut effortlessly and produce a clean freeze-fractured surface for SEM investigation. By following the approach described in this protocol, both the internal structure and surface morphology of Chlorella sorokiniana can be easily resolved with high quality. This protocol is highly versatile and can be applied to samples dispersed in water or solvents, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto filter paper.

The mammalian kinetochore is a multi-layered protein complex that forms on the centromeric chromatin. The kinetochore serves as the attachment hub for the plus ends of microtubules emanating from the centrosomes during mitosis. For karyokinesis, bipolar kinetochore-microtubule attachment and subsequent microtubule depolymerization lead to the development of inter-kinetochore tension between the sister chromatids. These events are instrumental in initiating a signaling cascade culminating in the segregation of the sister chromatids equally between the new daughter cells. Of the hundreds of conserved proteins that constitute the mammalian kinetochore, many that reside in the outermost layer are loaded during early mitosis and removed around metaphase-anaphase. Dynamically localized kinetochore proteins include those required for kinetochore-microtubule attachment, spindle assembly checkpoint proteins, various kinases, and molecular motors. The abundance of these kinetochore-localized proteins varies at prometaphase, metaphase, and anaphase, and is thus considered diagnostic of the fidelity of progression through these stages of mitosis. Here, we document detailed, state-of-the-art methodologies based on high-resolution fluorescence confocal microscopy followed by quantification of the levels of kinetochore-localized proteins during mitosis. We also document methods to accurately measure distances between sister kinetochores in mammalian cells, a surrogate readout for inter-kinetochore tension, which is essential for chromosome segregation.