Current Protocols in Cell Biology最新文献

Fluorescent proteins and vital dyes are invaluable tools for studying dynamic processes within living cells. However, the ability to distinguish more than a few different fluorescent reporters in a single sample is limited by the spectral overlap of available fluorophores. Here, we present a protocol for imaging live cells labeled with six fluorophores simultaneously. A confocal microscope with a spectral detector is used to acquire images, and linear unmixing algorithms are applied to identify the fluorophores present in each pixel of the image. We describe the application of this method to visualize the dynamics of six different organelles, and to quantify the contacts between organelles. However, this method can be used to image any molecule amenable to tagging with a fluorescent probe. Thus, multispectral live-cell imaging is a powerful tool for systems-level analysis of cellular organization and dynamics. © 2018 by John Wiley & Sons, Inc.

Reduction or complete prevention of ice crystal formation during freezing of biological specimens is mandatory for two important biological applications: (1) cryopreservation of living cells or tissues for long-term storage, and (2) cryo-fixation for ultrastructural investigations by electron microscopy. Here, a protocol that is fast, easy-to-use, and suitable for both cryo-fixation and cryopreservation is described. Samples are rapidly cooled in tightly sealed metal tubes of high thermal diffusivity and then plunged into a liquid cryogen. Due to the fast cooling speed and high-pressure buildup internally in the confined volume of the metal tubes, ice crystal formation is reduced or completely prevented, resulting in vitrification of the sample. For cryopreservation, however, a similar principle applies to prevent ice crystal formation during re-warming. A detailed description of procedures for cooling (and re-warming) of biological samples using this technique is provided. © 2018 by John Wiley & Sons, Inc.

This unit describes the purification of the multifunctional adhesive glycoprotein fibronectin from plasma or of cell-derived fibronectin from cell surfaces and from conditioned medium. Fibronectin can be used in cell adhesion and migration assays, and can be obtained in relatively high purity using simple affinity chromatography techniques. Curr. Protoc. Cell Biol. 60:10.5.1-10.5.13. © 2013 by John Wiley & Sons, Inc.

This unit describes the purification of extracellular vitronectin from plasma or serum by using heparin-affinity chromatography. First, the plasma is depleted of fibronectin plus other heparin- and Sepharose-binding proteins and treated with urea to activate the heparin-binding activity of vitronectin, which is subsequently bound to a heparin affinity column and eluted. The resulting vitronectin should be ∼98% pure. Curr. Protoc. Cell Biol. 60:10.6.1-10.6.7. © 2013 by John Wiley & Sons, Inc.

For more than 100 years, the ultimate resolution of a light microscope (∼200 nm) has been constrained by the fundamental physical phenomenon of diffraction, as described by Ernst Abbe in 1873. While this limitation is just as applicable to today's light microscopes, it is the combination of high-end optics, clever methods of sample illumination, and computational techniques that has enabled researchers to access information at an order of magnitude greater resolution than once thought possible. This combination, broadly termed superresolution microscopy, has been increasingly practical for many labs to implement from both a hardware and software standpoint, but, as with many cutting-edge techniques, it also comes with limitations. One of the current drawbacks to superresolution microscopy is the limited number of probes and conditions that have been suitable for imaging. Here, a technique termed bleaching/blinking-assisted localization microscopy (BaLM) makes use of the inherent blinking and bleaching properties of almost all fluorophores as a means to generate superresolution images. Curr. Protoc. Cell Biol. 60:21.8.1-21.8.17. © 2013 by John Wiley & Sons, Inc.

Autophagy is a lysosome-based degradation pathway. Autophagic lysosome reformation (ALR) is a lysosomal membrane recycling process that marks the terminal step of autophagy. During ALR, LAMP1-positive tubules, named reformation tubules, are extruded from autolysosomes, and nascent lysosomes are generated from these tubules. By combining proteomic analysis of purified autolysosomes and RNA interference screening of identified candidates, we systematically elucidated the ALR pathway at the molecular level. Based on the key components clathrin, PtdIns(4,5)P₂, and the motor protein KIF5B, among others, we reconstituted this process in vitro. This unit describes a detailed method for visualizing ALR in cells during the autophagy process. This unit also present a protocol for reconstituting the ALR tubular protrusion and elongation process in vitro and three methods for preparing materials for in vitro reconstitution: (1) autolysosome purification from cultured cells, (2) liposome preparation, and (3) KIF5B purification and quality testing. © 2018 by John Wiley & Sons, Inc.

Understanding the nature of fluorescently-labelled molecules requires proper quantification of their colocalization. We developed a new approach to enable quantification of colocalization of markers in fluorescence microscopy images using mobile computers. It consists of three interacting components: desktop computer – cloud – mobile device. After an image is opened on a desktop computer, it is then saved to the cloud and becomes available to a mobile device. Functionality of the desktop and mobile software consists of the same steps and therefore allows using any of the platforms when performing analysis. Changes of the state of the image between devices are synchronized via the cloud, so that nothing is lost when switching them. All interactions are performed seamlessly in the background and do not require any input from the researchers’ side. This approach augments access to analytical imaging by allowing to work in completely new ways with superior levels of control, simplicity, and convenience. © 2018 by John Wiley & Sons, Inc.

To date, the permeability of epithelia to larger solutes (greater than ∼4 Å in diameter) has been analyzed by flux measurements using various tracers that cannot spatially resolve the permeation sites. This unit describes a method for localizing such sites of passage in epithelial sheets with subcellular resolution. The method makes use of avidin as a basolateral capture probe in epithelial monolayers or mucosae to unmask the passage of biotinylated and fluorophore-labeled tracer molecules as they go through the junctional barrier. Once bound to avidin, the tracers are immobilized at the site of a barrier leak. The localization, the distribution, and the extent of passage are eventually evaluated by imaging. The assay detects single leaks and is hence able to spatially resolve rarely occurring changes. It is also modular and flexible to use with various macromolecular tracers, and its sensitivity is adjustable. If designed as a chase experiment, the method allows for analysis of temporal barrier openings. If performed at low temperatures, this assay will block transcellular passage and, combined with global flux measurement, unambiguously determine paracellular passage. © 2018 by John Wiley & Sons, Inc.

Genetically encoded live-cell reporters measure signaling pathway activity at the cellular level with high temporal resolution, often revealing a high degree of cell-to-cell heterogeneity. By using multiple spectrally distinct reporters within the same cell, signal transmission from one node to another within a signaling pathway can be analyzed to quantify factors such as signaling efficiency and delay. With other reporter configurations, correlation between different signaling pathways can be quantified. Such analyses can be used to establish the mechanisms and consequences of cell-to-cell heterogeneity and can inform new models of the functional properties of signaling pathways. In this unit, we describe an approach for designing and executing live-cell multiplexed reporter experiments. We also describe approaches for analyzing the resulting time-course data to quantify correlations and trends between the measured parameters at the single-cell level. © 2018 by John Wiley & Sons, Inc.