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.

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.

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.

Primary human intestinal stem cells (ISCs) can be cultured and passaged indefinitely as two-dimensional monolayers grown on soft collagen. Culturing ISCs as monolayers enables easy access to the luminal side for chemical treatments and provides a simpler topology for high-resolution imaging compared to cells cultured as three-dimensional organoids. However, the soft collagen required to support primary ISC growth can pose a challenge for live imaging with an inverted microscope, as the collagen creates a steep meniscus when poured into wells. This may lead to uneven growth toward the center of the well, with cells at the edges often extending beyond the working distance of confocal microscopes. We have engineered a 3D-printed collagen mold that enables the preparation of chamber slides with flat, smooth, and reproducible thin collagen layers. These layers are adequate to support ISC growth while being thin enough to optimize live imaging with an inverted microscope. We present methods for constructing the collagen press, preparing chamber slides with pressed collagen, and plating primary human ISCs for growth and analysis.

Drug-induced hearing injury (ototoxicity) is a common, debilitating side effect of many antibiotic regimens that can be worsened by adverse drug interactions. Such adverse drug interactions are often not detected until after drugs are already on the market because of the difficulty of measuring all possible drug combinations. While in vivo mammalian assays to screen for ototoxic damage exist, they are currently time-consuming, costly, and limited in throughput, which limits their utility in assessing drug interaction outcomes. To facilitate more rapid quantification of ototoxicity and assessment of adverse drug interactions that impact ototoxicity, we have developed a high-throughput workflow we call parallelized evaluation of protection and injury for toxicity assessment (PEPITA). PEPITA uses zebrafish larvae to quantify ototoxic damage and protection. Previous work has shown that hair cells (HCs) in the zebrafish lateral line are very similar to human inner ear HCs, meaning zebrafish are a viable model to test drug-induced ototoxicity. In PEPITA, we expose zebrafish larvae to different combinations of drugs, fluorescently label the HCs, and subsequently use microscopy to quantify the brightness of the fluorescently labeled HCs as an assay for ototoxic damage and hair-cell viability. PEPITA is a reproducible, low-cost, technically accessible, and high-throughput assay. These advantages allow many experiments to be conducted in parallel, paving the way for systematic evaluation of drug-induced hearing injury and other multidrug interactions.

Phosphoinositides are rare membrane lipids that mediate cell signaling and membrane dynamics. PI(4)P and PI(3)P are two major phosphoinositides crucial for endolysosomal functions and dynamics, making them the lipids of interest in many studies. The acute modulation of phosphoinositides at a given organelle membrane can reveal important insights into their cellular function. Indeed, the localized depletion of PI(4)P and PI(3)P is a viable tool to assess the importance of these phosphoinositides in various experimental conditions. Here, we describe a live imaging method to acutely deplete PI(4)P and PI(3)P on endolysosomes. The depletion assay utilizes the GAI-GID1 or the FRB-FKBP inducible dimerization system to target the catalytic domain of the PI(4)P phosphatase, Sac1, or the PI(3)P phosphatase domain of MTM1 to the endolysosome for localized depletion of these phosphoinositides. By using the fluorescently tagged biosensors, 2xP4M and PX, we can validate and monitor the depletion of PI(4)P and PI(3)P, respectively, on endolysosomes in real-time. We discuss a method for normalizing the fluorescence measurements to appropriate the relative amount of these phosphoinositides in the organellar membranes (endolysosomes), which is required for monitoring PI(4)P or PI(3)P levels during the acute depletion assay. Since the localization of the dimerization partners is specified by the membrane targeting signal, our protocol will be useful for studying the signaling and functions of phosphoinositides at any membrane.

Protein misfolding fuels multiple neurodegenerative diseases, but existing techniques lack the resolution to pinpoint the location and physical properties of aggregates within living cells. Our protocol describes high-resolution confocal and fluorescent lifetime microscopy (Fast 3D FLIM) of an aggregation probing system. This system involves a metastable HaloTag protein (HT-aggr) labeled with P1 solvatochromic fluorophore, which can be targeted to subcellular compartments. This strategy allows to distinguish between aggregated and folded probe species, since P1 fluorophore changes its lifetime depending on the hydrophobicity of its microenvironment. The probe is not fluorescence intensity-dependent, overcoming issues related to intensity-based measurements of labeled proteins, such as control of probe quantity due to differences in expression or photobleaching of a proportion of the fluorophore population. Our approach reports on the performance of the machinery dealing with aggregation-prone substrates and thus opens doors to studying proteostasis and its role in neurodegenerative diseases.

As an essential process for the maintenance of cellular homeostasis and function, autophagy is responsible for the lysosome-mediated degradation of damaged proteins and organelles; therefore, dysregulation of autophagy in humans can lead to a variety of diseases. The link between impaired autophagy and disease highlights the need to investigate possible interventions to address dysregulations. One possible intervention is hyperthermia, which is described in this protocol. To investigate these interventions, a method for absolute quantification of autophagosomal compartments is required that allows comparison of autophagosomal activity under different conditions. Existing methods such as western blotting and immunohistochemistry for analysing the location and relative abundance of intracellular proteins associated with autophagy, or transmission electron microscopy (TEM), which are either very time-consuming, expensive, or both, are less suitable for this purpose. The method described in this protocol allows the absolute quantification of autophagosomes per cell in human fibroblasts using the CYTO-ID^® Autophagy Detection Kit after heat therapy compared to a control. The Cyto-ID^® assay is based on the use of a specific dye that selectively stains autophagic compartments, combined with an additional Hoechst 33342 dye for nuclear staining. The subsequent recognition of these stained compartments by the Cytation Imager enables the software to determine the number of autophagosomes per nucleus in living cells. Additionally, this absolute quantification uses an image-based method, and the protocol is easy to use and not time-consuming. Furthermore, the method is not only suitable for heat therapy but can also be adapted to any other desired therapy or substance.

All aerial organs in plants originate from the shoot apical meristem, a specialized tissue at the tip of a plant, enclosing a few stem cells. Understanding developmental dynamics within this tissue in relation to internal and external stimuli is of crucial importance. Imaging the meristem at the cellular level beyond very early stages requires the apex to be detached from the plant body, a procedure that does not allow studies in living, intact plants over longer periods. This protocol describes a new confocal microscopy method with the potential to image the shoot apical meristem of an intact, soil-grown, flowering Arabidopsis plant over several days. The setup opens new avenues to study apical stem cells, their interconnection with the whole plant, and their responses to environmental stimuli.

Calcium signalling in the endocardium is critical for heart valve development. Calcium ion pulses in the endocardium are generated in response to mechanical forces due to blood flow and can be visualised in the beating zebrafish heart using a genetically encoded calcium indicator such as GCaMP7a. Analysing these pulses is challenging because of the rapid movement of the heart during heartbeat. This protocol outlines an imaging analysis method used to phase-match the cardiac cycle in single z-slice movies of the beating heart, allowing easy measurement of the calcium signal.