细胞生物学

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.

The organ of Corti, located in the inner ear, is the primary organ responsible for animal hearing. Each hair cell has a V-shaped or U-shaped hair bundle composed of actin-filled stereocilia and a kinocilium supported by true transport microtubules. Damage to these structures due to noise exposure, drug toxicity, aging, or environmental factors can lead to hearing loss and other disorders. The challenge when examining auditory organs is their location within the bony labyrinth and their small and fragile nature. This protocol describes the dissection procedure for the cochlear organ, followed by confocal imaging of immunostained endogenous and fluorescent proteins. This approach can be used to understand hair cell physiology and the molecular mechanisms required for normal hearing.

The motile parameters of kinesin superfamily proteins are fundamental to intracellular transport. Single-molecule motility assays using total internal reflection fluorescence (TIRF) microscopy are a gold standard technique for measuring the motile parameters of kinesin motors. With this technique, one can evaluate the velocity, run length, and binding frequency of kinesins on microtubules by directly observing their motility. This protocol provides a comprehensive procedure for single molecule assays of kinesins, including the preparation of labeled microtubules, the measurement of kinesin motility via TIRF microscopy, and the quantification of kinesin motor parameters.

The eukaryotic cytoskeleton is formed in part by microtubules, which are relatively rigid filaments with inherent structural polarity. One consequence of this polarity is that the two ends of a microtubule have different properties with important consequences for their cellular roles. These differences are often challenging to probe within the crowded environment of the cell. Fluorescence microscopy–based in vitro assays with purified proteins and stabilized microtubules have been used to characterize polarity-dependent and end-specific behaviors. These assays require ways to visualize the polarity of the microtubules, which has previously been achieved either by the addition of fluorescently tagged motor proteins with known directionality or by fluorescently polarity marking the microtubules themselves. However, classical polarity-marking protocols require a particular chemically modified tubulin and generate microtubules with chemically different plus and minus segments. These chemical differences in the segments may affect the behavior of interacting proteins of interest in an undesirable manner. We present here a new protocol that uses a previously characterized, reversibly binding microtubule plus-end capping protein, a designed ankyrin repeat protein (DARPin), to efficiently produce polarity-marked microtubules with different fluorescently labeled, but otherwise biochemically identical, plus- and minus-end segments.

MreB is a prokaryotic actin homolog. It is essential for cell shape in the majority of rod-shaped cell-walled bacteria. Structural and functional characterization of MreB protein is important to understand the mechanism of ATP-dependent filament dynamics and membrane interaction. In vitro studies on MreBs have been limited due to the difficulty in purifying the homogenous monomeric protein. We have purified MreB from the cell-wall-less bacteria Spiroplasma citri, ScMreB5, using heterologous expression in Escherichia coli. This protocol provides a detailed description of purification condition optimization that led us to obtain high concentrations of stable ScMreB5. Additionally, we have provided a protocol for detecting the presence of monovalent ions in the ScMreB5 AMP-PNP-bound crystal structure. This protocol can be used to obtain a high yield of ScMreB5 for carrying out biochemical and reconstitution studies. The strategies used for ScMreB5 show how optimizing buffer components can enhance the yield and stability of purified protein.