细胞生物学

When plants undergo senescence or experience carbon starvation, leaf cells degrade proteins in the chloroplasts on a massive scale via autophagy, an evolutionarily conserved process in which intracellular components are transported to the vacuole for degradation to facilitate nutrient recycling. Nonetheless, how portions of chloroplasts are released from the main chloroplast body and mobilized to the vacuole remains unclear. Here, we developed a method to observe the autophagic transport of chloroplast proteins in real time using confocal laser-scanning microscopy on transgenic plants expressing fluorescently labeled chloroplast components and autophagy-associated membranes. This protocol enabled us to track changes in chloroplast morphology during chloroplast-targeted autophagy on a timescale of seconds, and it could be adapted to monitor the dynamics of other intracellular processes in plant leaves.

Cellular phenomena such as signal integration and transmission are based on the correct spatiotemporal organization of biomolecules within the cell. Therefore, the targeted manipulation of such processes requires tools that can precisely induce the localizations and interactions of the key players relevant to these processes with high temporal resolution. Chemically induced dimerization (CID) techniques offer a powerful means to manipulate protein function with high temporal resolution and subcellular specificity, enabling direct control over cellular behavior. Here, we present the detailed synthesis and application of dual SLIPT and dual SLIPT^NVOC, which expand the SLIPT (self-localizing ligand-induced protein translocation) platform by incorporating a dual-ligand CID system. Dual SLIPT and dual SLIPT^NVOC independently sort into the inner leaflet of the plasma membrane via a lipid-like anchoring motif, where they present the two headgroup moieties trimethoprim (TMP) and HaloTag ligand (HTL), which recruit and dimerize any two ^iK6eDHFR- and HOB-tagged proteins of interest (POIs). Dual-SLIPT^NVOC furthermore enables this protein dimerization of POIs at the inner leaflet of the plasma membrane in a pre-determined order and light-controlled manner. In this protocol, we detail the synthetic strategy to access dual SLIPT and dual SLIPT^NVOC, while also providing the underlying rationale for key design and synthetic decisions, with the aim of offering a streamlined, accessible, and broadly implementable methodology. In addition to the detailed synthesis, we present representative applications and typical experimental outcomes and recommend strategies for data analysis to support effective use of the system. Notably, dual SLIPT and dual SLIPT^NVOC represent the first CID systems to emulate endogenous lipidation-driven membrane targeting, while retaining hallmark advantages of CID technology—the precision over POI identity, recruitment sequence, high spatiotemporal control, and “plug-and-play” flexibility.

Cell–surface and cell–cell interaction assays are fundamental for studying receptor–ligand interactions and characterizing cellular responses and functions. They play a critical role in diagnostics and in modulating immune system activity for therapeutic applications, notably in cancer immunotherapy. By providing time-lapsed and cell-level direct observation of the sample, optical microscopy offers strong advantages compared to current go-to techniques, which are typically either ensemble methods (e.g., measuring cell populations) or indirect readouts (e.g., impedance for adherent cells). This protocol describes two complementary microscopy-based assays: (1) a cell–surface ligand binding assay to quantify dynamic interactions between human primary Natural Killer (NK) cells and a cancer-mimicking surface, and (2) a cell–cell interaction assay to evaluate antibody-dependent cell cytotoxicity (ADCC) mediated by NK cells targeting tumor cells. Additionally, the protocol uses Celldetective, a new open graphical user interface for quantitative analysis of cell interaction dynamics from 2D time-lapse microscopy datasets. Although applied here to primary immune cells, these methods are adaptable to various cell types, including other immune cells, fibroblasts, and cancer cells. This approach enables direct observation and quantification of cellular morphology, motility, cell–cell interactions, and dynamic behaviors at single-cell resolution over time, facilitating detailed analysis of mechanisms such as cell death, migration, and immune synapse formation.

Proper genome organization is essential for genome function and stability. Disruptions to this organization can lead to detrimental effects and the transformation of cells into diseased states. Individual chromosomes and their subregions can move or rearrange during transcriptional activation, in response to DNA damage, and during terminal differentiation. Techniques such as fluorescence in situ hybridization (FISH) and chromosome conformation capture (e.g., 3C and Hi-C) have provided valuable insights into genome architecture. However, these techniques require cell fixation, limiting studies of the temporal evolution of chromatin organization in detail. Our understanding of the heterogeneity and dynamics of chromatin organization at the single-cell level is still emerging. To address this, clustered regularly interspaced short palindromic repeats (CRISPR)/dead Cas9 (dCas9) systems have been repurposed for precise live-cell imaging of genome dynamics. This protocol uses a system called CRISPRainbow, a powerful tool that allows simultaneous targeting of up to seven genomic loci and tracks their locations over time using spectrally distinct fluorescent markers to study real-time chromatin organization. Multiple single-guide RNA (sgRNA), carrying specific RNA aptamers for labeling, can be cloned into a single vector to improve transfection efficiency in human cells. The precise targeting of CRISPRainbow offers distinct advantages over previous techniques while also complementing them by validating findings in live cells.

An improved correlative light and electron microscopy (CLEM) method has recently been introduced and successfully employed to identify and analyze protein inclusions in cultured cells as well as pathological proteinaceous deposits in postmortem human brain tissues from individuals with diverse neurodegenerative diseases. This method significantly enhances antigen preservation and target registration by replacing conventional dehydration and embedding reagents. It achieves an optimal balance of sensitivity, accuracy, efficiency, and cost-effectiveness compared to other current CLEM approaches. However, due to space constraints, only a brief overview of this method was provided in the initial publication. To ensure reproducibility and facilitate widespread adoption, the author now presents a detailed, step-by-step protocol of this optimized CLEM technique. By enhancing usability and accessibility, this protocol aims to promote broader application of CLEM in neurodegenerative disease research.

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.

Accurate identification of cell cycle stages is essential for investigating fundamental biological processes such as proliferation, differentiation, and tumorigenesis. While flow cytometry remains a widely used technique for such analyses, it is limited by its lack of single-cell resolution and its requirement for large sample sizes due to its population-based approach. These limitations underscore the need for alternative or complementary methods that offer single-cell precision with compatibility for small-scale applications. We present ImmunoCellCycle-ID, an immunofluorescence-based method that leverages the spatial distribution of endogenous markers, such as DNA, proliferating cell nuclear antigen (PCNA), centromere protein F (CENP-F), and centromere protein C (CENP-C), to reliably distinguish G1, early S, late S, early G2, late G2, and all mitotic sub-stages. This technique does not rely on precise signal quantification and utilizes standard immunofluorescence protocols alongside conventional laboratory microscopes, ensuring broad accessibility. Importantly, ImmunoCellCycle-ID detects endogenous proteins without the need for genetic modification, making it readily applicable to a wide range of human cell lines. Beyond its utility for single-cell resolution, the method can be scaled for population-level analyses, similar to flow cytometry. With its precision, versatility, and ease of implementation, ImmunoCellCycle-ID offers a powerful tool for high-resolution cell cycle profiling across diverse experimental platforms.

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.

Cryo-electron tomography (cryo-ET) is the main technique to image the structure of biological macromolecules inside their cellular environment. The samples for cryo-ET must be thinner than 200 nm, which is not compatible with micron-sized cells. A focused ion beam (FIB), in conjunction with a scanning electron microscope (SEM) to navigate the sample, can be used to ablate material from vitrified cells such that a thin lamella remains. However, the preparation of lamellae with a FIB-SEM is blind to the location of specific cellular structures and biomolecules. Furthermore, the thickness and uniformity of lamella, while crucial for high-quality tomograms, cannot be established accurately with the FIB-SEM. These limitations strongly affect the success rate for cryo-ET on FIB-milled lamellae and thereby the total throughput of the workflow. To mitigate these problems, a coincident light, electron, and ion beam cryo-microscope was developed by retrofitting a fluorescence microscope, cryogenic microcooler, and piezo stage on a FIB-SEM. The fluorescence of molecules of interest can be monitored in real time while milling to ensure the final lamella contains the structure of interest. In addition, reflected light microscopy can be used for thickness and quality control of the lamella. In this protocol, we will describe how the coincident microscope can be used to prepare lamellae from vitrified cells.

Counting protein molecules helps reveal the organization of components within cellular structures and the stoichiometries of protein complexes. Existing protein and peptide quantitation methods vary in their complexity. Here, we report a straightforward workflow to measure the absolute number of HaloTag-labeled myosin 10 (Myo10) molecules in U2OS cells. Myo10 is a motor protein that plays a prominent role in cellular protrusion formation. Various biochemical and biological properties of Myo10 are established, but it is not well-defined how many molecules of Myo10 pack into narrow cellular structures called filopodia. We present a workflow for using SDS-PAGE to calibrate Myo10 signal with a reference protein, segmenting epifluorescence microscopy images to map Myo10 intracellular distribution, and interpreting the results to derive biological and functional insights. Our protocol is simple to employ and not only applicable for Myo10 research but also easily adaptable for other biological systems that use HaloTag.