细胞生物学

ER-phagy, a selective autophagy process crucial for maintaining cellular homeostasis by targeting the endoplasmic reticulum (ER), has been challenging to study in vivo due to the lack of suitable spatiotemporal quantification tools. Existing methods like electron microscopy, biochemical assays, and in vitro reporters lack resolution, scalability, or physiological relevance. Here, we present a detailed protocol for generating two transgenic mouse models: ER-TRG (constitutively expressing an ER lumen-targeting tandem RFP-GFP tag) and CA-ER-TRG (Cre-recombinase-activated ER-TRG). Additionally, we outline procedures for quantitative imaging of ER-phagy in vivo, covering tissue preparation, confocal microscopy, and signal analysis. This protocol offers a robust and reproducible tool for investigating ER-phagy dynamics across various tissues, developmental stages, and pathophysiological conditions, facilitating both fundamental and translational research.

Primary cilia are evolutionarily conserved organelles that play critical roles in brain development. In the developing cortex, neural progenitors extend their primary cilia into the ventricular surface, where the cilia act as key signaling hubs. However, visualizing these cilia in a systematic and intact manner has been challenging. The commonly used cryostat sectioning only provides a limited snapshot of cilia on individual sections, and this process often disrupts the ciliary morphology. By contrast, the previously established whole-mount technique has been shown to preserve ciliary architecture in the adult mouse cortex. Here, we adapt and optimize the whole-mount approach for embryonic and neonatal brain, allowing robust visualization of ciliary morphology at the ventricular surface during development. This protocol describes step-by-step procedures for whole-mounting and immunostaining delicate embryonic and neonatal mouse cortices, enabling direct visualization of cilia in neural progenitors in the developing brain.

Expansion microscopy (ExM) enables nanoscale imaging of biological structures using standard fluorescence microscopes. Accurate labeling of cytoskeletal filaments, such as microtubules, remains challenging due to structural distortion and labeling inaccuracy during sample preparation. This protocol describes an optimized method combining detergent extraction and NHS-ester labeling for high-precision visualization of microtubules in expanded samples. Cytoplasmic components and membranes are selectively removed, preserving the ultrastructure of the microtubule network. Microtubules are digested into peptides during expansion and subsequently labeled at their N-termini using NHS-ester dyes, eliminating the need for antibodies. Effective fluorophore displacement of ~1 nm or lower is achieved, depending on the applied expansion factor. The protocol is compatible with both in vitro and cellular samples and can be integrated into a wide range of ExM workflows. Labeled microtubules can serve as internal reference standards for correcting expansion factors in ExM datasets.

Quantitative analysis of biological membrane morphology is essential for understanding fundamental cellular processes such as organelle biogenesis and remodeling. While manual annotation has been the standard for complex structures, it is laborious and subjective, and conventional automated methods often fail to accurately delineate overlapping objects in 2D projected microscopy images. This protocol provides a complete, step-by-step workflow for the quantitative analysis of overlapping prospore membranes (PSMs) in sporulating yeast. The procedure details the synchronous induction of sporulation, acquisition of 3D fluorescence images and their conversion to 2D maximum intensity projections (MIPs), and the generation of a custom-annotated dataset using a semi-automated pipeline. Finally, it outlines the training and application of our mask R-CNN-based model, DeMemSeg, for high-fidelity instance segmentation and the subsequent extraction of morphological parameters. The primary advantage of this protocol is its ability to enable accurate and reproducible segmentation of individual, overlapping membrane structures from widely used 2D MIP images. This framework offers an objective, efficient, and scalable solution for the detailed quantitative analysis of complex membrane morphologies.

Protein synthesis and degradation (i.e., turnover) forms an important part of protein homeostasis and has been implicated in many age-associated diseases. Different cellular locations, such as organelles and membraneless compartments, often contain individual protein quality control and degradation machineries. Conventional methods to assess protein turnover across subcellular compartments require targeted genetic manipulation or isolation of specific organelles. Here we describe a protocol for simultaneous proteome localization and turnover (SPLAT) analysis, which combines protein turnover measurements with unbiased subcellular spatial proteomics to measure compartment-specific protein turnover rates on a proteome-wide scale. This protocol utilizes dynamic stable isotope labeling of amino acids in cell culture (dynamic SILAC) to resolve the temporal information of protein turnover and multi-step differential ultracentrifugation to assign proteins to multiple subcellular localizations. We further incorporate 2D liquid chromatography fractionation to greatly increase analytical depth while multiplexing with tandem mass tags (TMT) to reduce acquisition time 10-fold. This protocol resolves the spatial and temporal distributions of proteins and can also reveal temporally distinct spatial localizations within a protein pool.

PIEZO1 is a mechanically activated ion channel essential for mechanotransduction and downstream signaling in almost all organ systems. Western blotting is commonly used to study the expression, stability, and post-translational modifications of proteins. However, as a large transmembrane protein, PIEZO1 contains extensive hydrophobic regions and undergoes post-translational modifications that increase its propensity for nonspecific protein–protein interactions. As a result, conventional sample preparation methods seem unsuitable for PIEZO1. For example, heating and sonicating transmembrane proteins exposes hydrophobic regions, leading to aggregation, improper detergent interactions, and loss of solubility, ultimately compromising their detection in western blots. To address these challenges, we developed a western blot protocol optimized for human PIEZO1 by preparing lysates consistently at lower temperatures and incorporating strong reducing and alkylation reagents into the western blot lysis buffer to ensure proper protein solubilization and minimal cross-linking. Using the same antibody, we also developed an immunoprecipitation protocol with optimized detergents to maintain the solubilization of native human PIEZO1, enabling the discovery of a new family of auxiliary subunits.

In vitro systems based on Xenopus egg extracts have elucidated many aspects of spindle assembly. Still, numerous unknowns remain, particularly concerning the variation in spindle morphologies. The X. laevis and X. tropicalis egg extract systems, which recapitulate diverse spindle sizes and architectures, serve as ideal tools to investigate the regulation of spindle morphometrics. However, fully understanding spindle architectural differences is hindered by the spindle's size and high microtubule density. Indeed, classical fluorescence microscopy lacks the resolution to detail the organization of spindle microtubules, and although electron tomography can distinguish individual microtubules, segmenting thousands of microtubules and tracking them across dozens of sections remains an unachieved challenge. Therefore, we set out to apply expansion microscopy to the study of Xenopus egg extract spindles. During this process, we realized that optimizing spindle fixation as well was crucial to preserve microtubule integrity. Here, we present an optimized fixation and expansion microscopy protocol that enables the study of spindle architecture in egg extracts of both X. laevis and X. tropicalis. Our method retains the fluorescence of rhodamine tubulins added to the extracts and allows for both pre- and post-expansion immunofluorescence analysis.

Centrosomes are vital eukaryotic organelles involved in regulating cell adhesion, polarity, mobility, and microtubule (MT) spindle assembly during mitosis. Composed of two centrioles surrounded by the pericentriolar material (PCM), centrosomes serve as the primary microtubule-organizing centers (MTOCs) in proliferating cells. The PCM is crucial for MT nucleation and centriole biogenesis. Centrosome numbers are tightly regulated, typically duplicating once per cell cycle, during the S phase. Deregulation of centrosome components can lead to severe diseases. While traditionally viewed as stable structures, centrosomes can be inactivated or disappear in differentiating cells, such as epithelial cells, muscle cells, neurons, and oocytes. Despite advances in understanding centrosome biogenesis and function, the mechanisms maintaining mature centrosomes or centrioles, as well as the pathways regulating their inactivation or elimination, remain less explored. Studying centrosome maintenance is challenging as it requires the uncoupling of centrosome biogenesis from maintenance. Tools for acute spatial-temporal manipulation are often unavailable, and manipulating multiple components in vivo is complex and time-consuming. This study presents a protocol that decouples centrosome biogenesis from maintenance, allowing the study of critical factors and pathways involved in the maintenance of the integrity of these important cellular structures.

Phosphatidylinositol 4,5-bisphosphate [PI(4,5)P₂] is a phospholipid enriched on the cytoplasmic leaflet of the plasma membrane, where it plays important roles in membrane trafficking and cytoskeletal dynamics through proteins that directly bind to it. PI(4,5)P₂ can be metabolized to other phosphorylated forms of phosphatidylinositol to regulate numerous processes such as cell growth and development. PI(4,5)P₂ can also be hydrolyzed to generate the second messengers diacylglycerol (DAG) and inositol triphosphate (IP₃). Altered metabolism or mislocalization of PI(4,5)P₂ can perturb one or more of its functions and contribute to disease states. Here, we present a protocol to visualize and quantify the localization of PI(4,5)P₂ in live cells. The protocol uses a highly specific PI(4,5)P₂ protein binding domain coupled to enhanced green fluorescence protein (PH-PLCD1-GFP), enabling localization and quantification of cytosol-facing PI(4,5)P₂ to be determined. Localization and quantification of the PH-PLCD1-GFP, PI(4,5)P₂ specific probe, is enabled by fluorescence imaging and confocal microscopy. This approach can be used to study the dynamics of PI(4,5)P₂ localization temporally in live cells under both physiological and pathological conditions.

The osteocyte lacuno-canalicular system (LCS) plays a crucial role in maintaining bone homeostasis and mediating cellular mechanotransduction. Current histological techniques, particularly the Ploton silver nitrate staining method, face challenges such as variations in solution concentrations and types as well as a lack of standardization, which limits their broader application in osteocyte research. In this study, we present a simplified and more effective silver nitrate staining protocol designed to address these issues. Our method utilizes a 1 mol/L silver nitrate solution combined with optimized gelatin-formic acid solutions at varying concentrations (0.05%–0.5% type-B gelatin and 0.05%–5% formic acid, or 1%–2% type-B gelatin and 0.1%–2% formic acid). Staining is performed for 1 h under 254 nm ultraviolet light or 90 min under room light, followed by washing with Milli-Q water to terminate staining. This novel optimized method yields consistent and distinct staining of the osteocyte LCS across multiple species, demonstrating superior efficiency and reliability compared to the Ploton method. It will significantly advance research in osteocyte biology and provide a valuable tool for exploring the adaptive evolution of osteocyte LCS morphology and function across various taxa.