Current Protocols in Cell Biology最新文献

The rapid advancement of live-cell imaging technologies has enabled biologists to generate high-dimensional data to follow biological movement at the microscopic level. Yet, the “perceived” ease of use of modern microscopes has led to challenges whereby sub-optimal data are commonly generated that cannot support quantitative tracking and analysis as a result of various ill-advised decisions made during image acquisition. Even optimally acquired images often require further optimization through digital processing before they can be analyzed. In writing this article, we presume our target audience to be biologists with a foundational understanding of digital image acquisition and processing, who are seeking to understand the essential steps for particle/object tracking experiments. It is with this targeted readership in mind that we review the basic principles of image-processing techniques as well as analysis strategies commonly used for tracking experiments. We conclude this technical survey with a discussion of how movement behavior can be mathematically modeled and described. © 2019 by John Wiley & Sons, Inc.

Although single-cell RNA sequencing (scRNA-seq) has become one of the most powerful methods available for transcriptome analysis, the quality of scRNA-seq data largely depends on cell preparation. Cell preparation from cultured cells and tissues requires different methods because of the inherent differences between these two categories of cells. Compared to cultured cells, tissues have more extracellular matrix, and the cells are generally more adherent and thus difficult to dissociate. The challenge is to achieve sufficient dissociation, cell counts, and viability all at the same time. This protocol describes approaches that help achieve these goals. These include a cold dissociation technique using cryophilic proteases active at cold temperature, timing of trituration during protease digestion, as well as filtration and washing methods that optimize cell viability and retention. Materials and equipment that optimize the process will also be discussed. © 2019 by John Wiley & Sons, Inc.

Genetically encoded Förster resonance energy transfer (FRET)-based tension sensors measure piconewton-scale forces across individual molecules in living cells or whole organisms. These biosensors show comparably high FRET efficiencies in the absence of tension, but FRET quickly decreases when forces are applied. In this article, we describe how such biosensors can be generated for a specific protein of interest, and we discuss controls to confirm that the observed differences in FRET efficiency reflect changes in molecular tension. These FRET efficiency changes can be related to mechanical forces as the FRET–force relationship of the employed tension sensor modules are calibrated. We provide information on construct generation, expression in cells, and image acquisition using live-cell fluorescence lifetime imaging microscopy (FLIM). Moreover, we describe how to analyze, statistically evaluate, and interpret the resulting data sets. Together, these protocols should enable the reader to plan, execute, and interpret FRET-based tension sensor experiments. © 2019 by John Wiley & Sons, Inc.

Protein-protein interactions (PPIs) are principle biological processes that control normal cell growth, differentiation, and homeostasis but are also crucial in diseases such as malignancy, neuropathy, and infection. Despite the importance of PPIs in biology, this target class has been very challenging to convert to therapeutics. In the last decade, much progress has been made in the inhibition of PPIs involved in diseases, but many remain difficult such as RAS-effector interactions in cancers. We describe here a protocol for using Bioluminescence Resonance Energy Transfer 2 (BRET2)-based RAS biosensors to detect and characterize RAS PPI inhibition by macromolecules and small molecules. This method could be extended to any other small GTPases or any other PPIs of interest. © 2019 by John Wiley & Sons, Inc.

Visualization of dynamic cellular activity has greatly expanded our understanding of brain function. Recently, there has been an increasing number of studies imaging rodent brain activity in real time. However, traditional in vivo calcium imaging technology has been limited to superficial brain structures. Because the trigeminal ganglion (TG) is located deep within the cranial cavity of mice, few studies have been able to access to it. To circumvent this limitation, overlying brain tissue must be removed to expose the TG so that optical recording can access deep brain neural ensembles. This unit describes a procedure for conducting non-survival surgery to visualize the TG in live mice. Obtaining large ensembles of direct, real-time readouts of sensory neuron signaling, providing temporal and spatial information across the TG, will help to define the cellular basis of orofacial somatic sensing and pain perception. © 2019 by John Wiley & Sons, Inc.

Traditionally, studies of cells and tissues have been performed on isolated primary cells or immortalized cell lines by culturing them in 2D culture dishes or flasks. However, a caveat regarding 2D culture is that the cells poorly recapitulate their in vivo counterparts, mainly due to a lack of 3D cell-cell and cell–extracellular matrix interactions. In recent years, the development of in vitro organoids as 3D culture has gained substantial attention as a model to study different tissues. In adults, pancreatic ductal cells are considered as a source of stem or progenitor cells, so developing new methods to study ductal cells would be useful. Here, we provide a protocol to isolate mouse pancreatic ductal cells and a cost-effective protocol to generate 3D organoid structures from such ductal cells. Additionally, we have devised a protocol for immunostaining of intact whole organoids without sectioning. © 2018 by John Wiley & Sons, Inc.

Eukaryotic cells are highly compartmentalized and protein subcellular localization critically influences protein function. Identification of the subcellular localizations of proteins and their translocation events upon perturbation has mostly been confined to targeted studies or laborious microscopy-based methods. Here we describe a systematic mass spectrometry-based method for spatial proteomics. The approach uses simple fractionation profiling and has two applications: Firstly it can be used to infer subcellular protein localization on a proteome-wide scale, resulting in a protein map of the cell. Secondly, the method permits identification of changes in protein localization, by comparing maps made under different conditions, as a tool for unbiased systems cell biology. © 2018 by John Wiley & Sons, Inc.

The consequences of alterations in the distribution of intracellular organelles, observed in many diseases, are often not clear. Intracellular organelles alter their morphology and positioning to regulate cell homeostasis and function. We outline how organelle positioning can be studied employing a density-based analysis of 3D images applied to cells that show similar cellular geometries. Quantification is facilitated by the use of single cells seeded on micropatterned substrates that provide cues for controlled cell spreading. This minimal system mimics the reproducible distribution of organelles typically observed in tissues, simplifying image analysis and minimizing the number of cells required for the observation of robust phenotypes. Here we provide guidelines for how the majority of organelles can be efficiently analyzed in cells seeded on adhesive micropatterns. We exemplify how alterations in the positioning of different organelles as a result of the perturbation of the cytoskeleton or associated motor proteins can be efficiently quantified. © 2018 by John Wiley & Sons, Inc.