Cell Biology

In response to environmental changes, chloroplasts, the cellular organelles responsible for photosynthesis, undergo intracellular repositioning, a phenomenon known as chloroplast movement. Observing chloroplast movement within leaf tissues remains technically challenging in leaves consisting of multiple cell layers, where light scattering and absorption hinder deep tissue visualization. This limitation has been particularly problematic when analyzing chloroplast movement in the mesophyll cells of C₄ plants, which possess two distinct types of concentrically arranged photosynthetic cells. In response to stress stimuli, mesophyll chloroplasts aggregate toward the inner bundle sheath cells. However, conventional methods have not been able to observe these chloroplast dynamics over time in living cells, making it difficult to assess the influence of adjacent bundle sheath cells on this movement. Here, we present a protocol for live leaf section imaging that enables long-term and detailed observation of chloroplast movement in internal leaf tissues without chemical fixation. In this method, a leaf blade section prepared either using a vibratome or by hand was placed in a groove made of a silicone rubber sheet attached to a glass slide for microscopic observation. This technique allows for the quantitative tracking of chloroplast movement relative to the surrounding cells. In addition, by adjusting the sectioning angle and thickness of the unfixed leaf sections, it is possible to selectively inactivate specific cell types based on their size and shape differences. This protocol enables the investigation of the intercellular interactions involved in chloroplast dynamics in leaf tissues.

Brightfield microscopy is an ideal application for studying live cell systems in a minimally invasive manner. This is advantageous in long-term experiments to study dynamic cellular processes such as stress response. Depending on the sample type and preparation, the inherent qualities of brightfield microscopy, being very low contrast, can contribute to technical issues such as focal drift, sequencing lags, and complete failure of software autofocus systems. Here, we describe the use of microbeads as a focus aid for long-term live cell imaging to address these autofocus issues. This protocol is inexpensive to implement, without extensive additional sample preparation, and can be used to capture focused images of transparent cells in a label-free manner. To validate this protocol, a widefield inverted microscope was used with software-based autofocus to image overnight in time-lapse format, demonstrating the use of the beads to prevent focal drift in long-term experiments. This improves autofocus accuracy on relatively inexpensive microscopes without using hardware-based focus aids. To validate this protocol, the KNIME logistics software was used to train a random forest model to perform binary image classification.

Over the lifespan of an individual, brain function requires adjustments in response to environmental changes and learning experiences. During early development, neurons overproduce neurite branches, and neuronal pruning removes the unnecessary neurite branches to make a more accurate neural circuit. Drosophila motoneurons prune their intermediate axon bundles rather than the terminal neuromuscular junction (NMJ) by degeneration, which provides a unique advantage for studying axon pruning. The pruning process of motor axon bundles can be directly analyzed by real-time imaging, and this protocol provides a straightforward method for monitoring the developmental process of Drosophila motor neurons using live cell imaging.

AMPA-type receptors are transported large distances to support synaptic plasticity at distal dendritic locations. Studying the motion of AMPA receptor⁺ vesicles can improve our understanding of the mechanisms that underlie learning and memory. Nevertheless, technical challenges that prevent the visualization of AMPA receptor⁺ vesicles limit our ability to study how these vesicles are trafficked. Existing methods rely on the overexpression of fluorescent protein-tagged AMPA receptors from plasmids, resulting in a saturated signal that obscures vesicles. Photobleaching must be applied to detect individual AMPA receptor⁺ vesicles, which may eliminate important vesicle populations from analysis. Here, we present a protocol to study AMPA receptor⁺ vesicles that addresses these challenges by 1) tagging AMPA receptors expressed from native loci with HaloTag and 2) employing a block-and-chase strategy with Janelia Fluor-conjugated HaloTag ligand to achieve sparse AMPA receptor labeling that obviates the need for photobleaching. After timelapse imaging is performed, AMPA receptor⁺ vesicles can be identified during image analysis, and their motion can be characterized using a single-particle tracking pipeline.

Amyloplasts, non-photosynthetic plastids specialized for starch synthesis and storage, proliferate in storage tissue cells of plants. To date, studies of amyloplast replication in roots and the ovule nucelli from various plant species have been performed using electron and fluorescence microscopy. However, a complete understanding of amyloplast replication remains unclear due to the absence of experimental systems capable of tracking their morphology and behavior in living cells. Recently, we demonstrated that Arabidopsis ovule integument could provide a platform for live-cell imaging of amyloplast replication. This system enables precise analysis of amyloplast number and shape, including the behavior of stroma-filled tubules (stromules), during proplastid-to-amyloplast development in post-mitotic cells. Here, we provide technical guidelines for observing and quantifying amyloplasts using conventional fluorescence microscopy in wild-type and several plastid-division mutants of Arabidopsis.

Orthoflavivirus is an enveloped, positive-stranded RNA virus that buds into the endoplasmic reticulum (ER) lumen. The budded virus particles are subsequently transported to the Golgi apparatus and secreted into the extracellular environment via the conventional secretion pathway. In this protocol, we describe a method for monitoring the secretion of Orthoflavivirus particles from the ER. To visualize intracellular membrane trafficking, we combine two distinct imaging techniques: the retention using selective hooks (RUSH) system and the split green fluorescent protein (GFP) system. In this approach, GFP11, a peptide tag fused to prME, the outer coat structural protein of Japanese encephalitis virus particles, was co-expressed in HeLa cells along with two additional components: GFP1-10 fused to a streptavidin-binding peptide and a hook construct consisting of streptavidin fused to the ER retention sequence KDEL. Time-lapse imaging was performed after the addition of biotin, which releases the captured GFP-labeled subviral particles from the ER. This method enables synchronized visualization of intracellular subviral particle trafficking and serves as a valuable tool for analyzing the maturation process of Orthoflavivirus particles within cells.

The growth cone is a highly motile tip structure that guides axonal elongation and directionality in differentiating neurons. Migrating immature neurons also exhibit a growth cone–like structure (GCLS) at the tip of the leading process. However, it remains unknown whether the GCLS in migrating immature neurons shares the morphological and molecular features of axonal growth cones and can thus be considered equivalent to them. Here, we describe a detailed method for time-lapse imaging and optical manipulation of growth cones using a super-resolution laser-scanning microscope. To observe growth cones in elongating axons and migrating neurons, embryonic cortical neurons and neonatal ventricular–subventricular zone (V-SVZ)-derived neurons, respectively, were transfected with plasmids encoding fluorescent protein–conjugated cytoskeletal probes and three-dimensionally cultured in Matrigel, which mimics the in vivo background. At 2–5 days in vitro, the morphology and dynamics of these growth cones and their associated cytoskeletal molecules were assessed by time-lapse super-resolution imaging. The use of photoswitchable cytoskeletal inhibitors, which can be reversibly and precisely controlled by laser illumination at two different wavelengths, revealed the spatiotemporal regulatory machinery and functional significance of growth cones in neuronal migration. Furthermore, machine learning–based methods enabled us to automatically segment growth cone morphology from elongating axons and the leading process. This protocol provides a cutting-edge methodology for studying the growth cone in developmental and regenerative neuroscience, being adaptable for various cell biology and imaging applications.

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.