Current Protocols in Cell Biology最新文献

Membrane proteins mediate numerous essential cellular functions. Due to the aggregation propensity of hydrophobic transmembrane domains in aqueous environments, the targeting and insertion of membrane proteins pose major challenges to cells. In the Guided Entry of Tail-anchored protein (GET) pathway, an essential class of newly synthesized tail-anchored proteins (TAs) are chaperoned and guided by multiple targeting factors to the endoplasmic reticulum (ER) membrane. Deciphering the molecular mechanism of this cellular process has benefitted from successful in vitro reconstitution of individual molecular events in the GET pathway with purified components. Here we describe recently developed protocols for in vitro reconstitution of functional complexes of TA substrates with their targeting factors, for monitoring the transfer of TAs between targeting factors, and for the insertion of TA into the microsomal membrane. These procedures are generally applicable to the interrogation of other post-translational membrane protein targeting pathways. © 2018 by John Wiley & Sons, Inc.

Sphingolipids are structural components of organelle membranes that also participate in signal transduction pathways. Complex sphingolipids are trafficked from their site of synthesis in organelles of the early secretory pathway to the Golgi apparatus, the plasma membrane, and the endo-lysosomal system. We have developed fluorescence microscopy–based methods to monitor sphingolipid trafficking in coordination with secretory protein sorting. A sphingomyelin binding protein fused to a fluorescent protein, which we term “EQ-SM,” is implemented to monitor sphingomyelin trafficking from the Golgi apparatus to the plasma membrane via secretory vesicles. A protocol is provided to determine if a query protein of interest is secreted from the cell via vesicles enriched in EQ-SM, an indication that the vesicle membrane is enriched in sphingomyelin. A complementary protocol is described that implements a chemically modified form of sphingosine, a metabolic precursor to complex sphingolipids, to visualize ceramide and complex sphingolipids in fixed cells. © 2018 by John Wiley & Sons, Inc.

Brain organoids are an emerging technique for studying human neurodevelopment in vitro, with biomedical implications. However, three-dimensional tissue culture poses several challenges, including lack of nutrient exchange at the organoid core and limited imaging accessibility of whole organoids. Here we present a method for culturing organoids in a micro-fabricated device that enables in situ real-time imaging over weeks with efficient nutrient exchange by diffusion. Our on-chip approach offers a means for studying the dynamics of organoid development, cell differentiation, cell cycle, and motion. © 2018 by John Wiley & Sons, Inc.

Co-cultivation of tumor cells and liver resident immune cells or other non-parenchymal cells (NPCs) from the same donor is important for the study of cancer metastasis. So far, little is known about the mechanism of tumor cell or pathogen clearance, leukocyte infiltration, and immune cell recruitment in the human liver. To investigate these processes in vitro, the use of primary human hepatocytes and non-parenchymal cell, especially immune cell, co-culture systems play essential roles in the establishment of cell–cell and cell–extracellular matrix communications similar to native liver tissues. Hepatic non-parenchymal cells mainly comprise liver sinusoid endothelial cells (LSECs), microvascular endothelial cells, hepatic stellate cells, Kupffer cells (KCs), natural killer T (iNKT) cells, and dendritic cells (DCs). Here we describe procedures for preparation, isolation, and culture of human liver resident immune cells and other non-parenchymal cells. © 2018 by John Wiley & Sons, Inc.

Many cell-adhesive peptides have been identified from extracellular matrix (ECM) proteins, such as collagen, fibronectin, laminin, and vitronectin. ECM proteins have various cell-adhesive sequences. Most peptides demonstrate cell-adhesive activity when simply coated on a tissue culture plate, but solubility, conformation, and coating efficiency of the peptides can significantly alter their biological function. Evaluation of peptide cell-adhesive activity using peptide-conjugated polysaccharide constructs is a useful strategy for overcoming peptide solubility and conformation problems. After a simple modification of the polysaccharides, various polysaccharides (chitosan, alginate, and hyaluronate) can fix the peptides on the tissue culture plate quantitatively. The peptide-polysaccharide strategy can be used to fix different active peptides to the polysaccharide at same time, thus, mimicking the biological functions of the ECM. This paper describes the modification of polysaccharides that are suitable for covalently coupling the peptides and evaluation of the cell-adhesive activity of peptide as a peptide-polysaccharide matrix. © 2018 by John Wiley & Sons, Inc.

Different multicellular organisms undergo cell-cell fusion to form functional syncytia that support specialized functions necessary for proper development and survival. For years, monitoring the structural consequences of this process using live-cell imaging has been challenging due to the unpredictable timing of cell fusion events in tissue systems. Here we present a triggered vesicular stomatitis virus G-protein (VSV-G)-mediated cell-cell fusion assay that can be used to synchronize fusion between cells. This allows the study of cellular changes that occur during cell fusion. The process is induced using a fast wash of low pH isotonic buffer, promoting the fusion of plasma membranes of two or more adjacent cells within seconds. This approach is suitable for studying mixing of small cytoplasmic molecules between fusing cells as well as changes in organelle distribution and dynamics. © 2018 by John Wiley & Sons, Inc.

Many cells possess the ability to repair plasma membrane disruption in physiological conditions. Growing evidence indicates a correlation between membrane repair and many human diseases. For example, a negative correlation is observed in muscle where failure to reseal sarcolemma may contribute to the development of muscular dystrophies. Instead, a positive correlation is observed in cancer cells where membrane repair may be exacerbated during metastasis. Here we describe a protocol that combines laser technology for membrane damage, immunostaining with gold nanoparticles and imaging by fluorescence microscopy and transmission electron microscopy (TEM), which allows the characterization of the molecular machinery involved in membrane repair. Fluorescence microscopy enables to determine the subcellular localization of candidate proteins in damaged cells while TEM offers high-resolution ultrastructural analysis of the µm²-disruption site, which enables to decipher the membrane repair mechanism. Here we focus on the study of human skeletal muscle cells, for obvious clinical interest, but this protocol is also suitable for other cell types. © 2018 by John Wiley & Sons, Inc.

Expansion microscopy (ExM) is a recently developed technique that enables nanoscale-resolution imaging of preserved cells and tissues on conventional diffraction-limited microscopes via isotropic physical expansion of the specimens before imaging. In ExM, biomolecules and/or fluorescent labels in the specimen are linked to a dense, expandable polymer matrix synthesized evenly throughout the specimen, which undergoes 3-dimensional expansion by ∼4.5 fold linearly when immersed in water. Since our first report, versions of ExM optimized for visualization of proteins, RNA, and other biomolecules have emerged. Here we describe best-practice, step-by-step ExM protocols for performing analysis of proteins (protein retention ExM, or proExM) as well as RNAs (expansion fluorescence in situ hybridization, or ExFISH), using chemicals and hardware found in a typical biology lab. Furthermore, a detailed protocol for handling and mounting expanded samples and for imaging them with confocal and light-sheet microscopes is provided. © 2020 The Authors.