细胞生物学

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.

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.

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.