Current Protocols in Cell Biology最新文献

Measuring oxygen consumption allows for the role of mitochondrial function in biological phenomena and mitochondrial diseases to be determined. Although respirometry has become a common approach in disease research, current methods are limited by the necessity to process and measure tissue samples within 1 hr of acquisition. Detailed by Acin-Perez and colleagues, a new respirometry approach designed for previously frozen tissue samples eliminates these hurdles for mitochondrial study. This technique allows for the measurement of maximal respiratory capacity in samples frozen for long-term storage before testing. This protocol article describes the optimal tissue isolation methods and the combination of substrates to define electron transport chain function at high resolution in previously frozen tissue samples. © 2020 The Authors.

Basic Protocol 1: Sample collection, storage, and homogenization for previously frozen tissue respirometry

Basic Protocol 2: Running a Seahorse respirometry assay using previously frozen tissue samples

Basic Protocol 3: Normalization to mitochondrial content for previously frozen tissue respirometry

Biochemical methods can reveal stable protein-protein interactions occurring within cells, but the ability to observe transient events and to visualize the subcellular localization of protein-protein interactions in cells and tissues in situ provides important additional information. The Proximity Ligation Assay^® (PLA) offers the opportunity to visualize the subcellular location of such interactions at endogenous protein levels, provided that the probes that recognize the target proteins are within 40 nm. This sensitive technique not only elucidates protein-protein interactions, but also can reveal post-translational protein modifications. The technique is useful even in cases where material is limited, such as when paraffin-embedded clinical specimens are the only available material, as well as after experimental intervention in 2D and 3D model systems. Here we describe the basic protocol for using the commercially available Proximity Ligation Assay™ materials (Sigma-Aldrich, St. Louis, MO), and incorporate details to aid the researcher in successfully performing the experiments. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Proximity ligation assay

Support Protocol 1: Antigen retrieval method for formalin-fixed, paraffin-embedded tissues

Support Protocol 2: Creation of custom PLA probes using the Duolink™ In Situ Probemaker Kit when commercially available probes are not suitable

Basic Protocol 2: Imaging, quantification, and analysis of PLA signals

Science and medicine have become increasingly “human-centric” over the years. A growing shift away from the use of animals in basic research has led to the development of sophisticated in vitro models of various tissues utilizing human-derived cells to study physiology and disease. The human cornea has likewise been modeled in vitro using primary cells derived from corneas obtained from cadavers or post-transplantation. By utilizing a cell's intrinsic ability to maintain its tissue phenotype in a pre-designed microenvironment containing the required growth factors, physiological temperature, and humidity, tissue-engineered corneas can be grown and maintained in culture for relatively long periods of time on the scale of weeks to months. Due to its transparency and avascularity, the cornea is an optimal tissue for studies of extracellular matrix and cell-cell interactions, toxicology and permeability of drugs, and underlying mechanisms of scarring and tissue regeneration. This paper describes methods for the cultivation of corneal keratocytes, fibroblasts, epithelial, and endothelial cells for in vitro applications. We also provide detailed, step-by-step protocols for assembling and culturing 3D constructs of the corneal stroma, epithelial- and endothelial-stromal co-cultures and isolation of extracellular vesicles. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Isolating and culturing human corneal keratocytes and fibroblasts

Basic Protocol 2: Isolating and culturing human corneal epithelial cells

Basic Protocol 3: Isolating and culturing human corneal endothelial cells

Basic Protocol 4: 3D corneal stromal construct assembly

Basic Protocol 5: 3D corneal epithelial-stromal construct assembly

Basic Protocol 6: 3D corneal endothelial-stromal construct assembly

Basic Protocol 7: Isolating extracellular vesicles from corneal cell conditioned medium

Support Protocol: Cryopreserving human corneal fibroblasts, corneal epithelial cells, and corneal endothelial cells

BioID, a proximity biotinylation technique, offers a valuable approach to examine the interactions occurring within protein complexes that complements traditional protein biochemical methods. BioID has various advantages that are beneficial to the study of complexes, including an ability to detect insoluble and transient proteins. We have applied BioID to the study of integrin adhesion complexes (IACs), which are located at the junction between the plasma membrane and actin cytoskeleton. The use of multiple BioID baits enables a complex-wide, spatial annotation of IACs, which in turn facilitates the detection of novel proximal interactors and provides insights into IAC architecture. This article describes the labeling and affinity purification of IAC-proximal proteins and their analysis by label-free quantitative mass spectrometry. The article also outlines steps to identify high-confidence proximity interactors, and to interrogate the topology and functional relevance of proximity interaction networks through bioinformatic analyses. © 2020 The Authors.

Basic Protocol 1: Proximity biotinylation of integrin adhesion complex components

Basic Protocol 2: Mass spectrometry data processing by MaxQuant and detection of high-confidence proximal interactors

Basic Protocol 3: Bioinformatic analysis and data visualization

Construction of organized three-dimensional (3D) tissue with extracellular matrix (ECM) and multiple types of cells is important for tissue engineering to enable tissue function and enhance cellular function. However, the concentration of ECM and the thickness of the 3D tissue have been limited in previous methods due to a lack of permeability to nutrients and oxygen. Besides, it is difficult to use matured natural ECM as a cell scaffold without chemical modification due to its insolubility. In this article, we focus on multi-layered structure, which is commonly found in living tissue such as skin, blood vessels, and other organs. Here, we describe the preparation of a paper-like scaffold (ECM paper) from micro-fibered natural ECM and the construction of 3D multi-layered tissue composed of cell layers and ECM layers by stacking cell-seeded ECM papers. The thickness and components of the ECM layers are easily controllable by changing the composition of the ECM papers, and the fibrous structure of ECM paper shows high permeability and permits cell migration. Additionally, the ECM microfiber, which is physically defiberized from natural ECM, has a high ECM concentration equal to that of living tissue and high stability under physiological conditions. Therefore, this set of protocols enables construction of multi-layered 3D tissue composed of precisely controlled ECM layers and cell layers that may contribute to the assembly of tissue models. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Preparation of extracellular matrix paper

Basic Protocol 2: Evaluation of cellular function of cells on extracellular matrix paper

Basic Protocol 3: Construction of multi-layered 3D tissue

Incorporation of a stable-isotope metabolic tracer into cells or tissue can be followed at submicron resolution by multi-isotope imaging mass spectrometry (MIMS), a form of imaging secondary ion microscopy optimized for accurate isotope ratio measurement from microvolumes of sample (as small as ∼30 nm across). In a metabolic MIMS experiment, a cell or animal is metabolically labeled with a tracer containing a stable isotope. Relative accumulation of the heavy isotope in the fixed sample is then measured as an increase over its natural abundance by MIMS. MIMS has been used to measure protein turnover in single organelles, track cellular division in vivo, visualize sphingolipid rafts on the plasma membrane, and measure dopamine incorporation into dense-core vesicles, among other biological applications. In this article, we introduce metabolic analysis using NanoSIMS by focusing on two specific applications: quantifying protein turnover in single organelles of cultured cells and tracking cell replication in mouse tissues in vivo. These examples illustrate the versatility of metabolic analysis with MIMS. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Metabolic labeling for MIMS

Basic Protocol 2: Embedding of samples for correlative transmission electron microscopy and MIMS with a genetically encoded reporter

Alternate Protocol: Embedding of samples for correlative light microscopy and MIMS

Support Protocol: Preparation of silicon wafers as sample supports for MIMS

Basic Protocol 3: Analysis of MIMS data

Exosomes are 50- to 150-nm-diameter extracellular vesicles secreted by all mammalian cells except mature red blood cells and contribute to diverse physiological and pathological functions within the body. Many methods have been used to isolate and analyze exosomes, resulting in inconsistencies across experiments and raising questions about how to compare results obtained using different approaches. Questions have also been raised regarding the purity of the various preparations with regard to the sizes and types of vesicles and to the presence of lipoproteins. Thus, investigators often find it challenging to identify the optimal exosome isolation protocol for their experimental needs. Our laboratories have compared ultracentrifugation and commercial precipitation- and column-based exosome isolation kits for exosome preparation. Here, we present protocols for exosome isolation using two of the most commonly used methods, ultracentrifugation and precipitation, followed by downstream analyses. We use NanoSight nanoparticle tracking analysis and flow cytometry (Cytek^®) to determine exosome concentrations and sizes. Imaging flow cytometry can be utilized to both size exosomes and immunophenotype surface markers on exosomes (ImageStream^®). High-performance liquid chromatography followed by nano-flow liquid chromatography–mass spectrometry (LCMS) of the exosome fractions can be used to determine the presence of lipoproteins, with LCMS able to provide a proteomic profile of the exosome preparations. We found that the precipitation method was six times faster and resulted in a ∼2.5-fold higher concentration of exosomes per milliliter compared to ultracentrifugation. Both methods yielded extracellular vesicles in the size range of exosomes, and both preparations included apoproteins. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Pre-analytic fluid collection and processing

Basic Protocol 2: Exosome isolation by ultracentrifugation

Alternate Protocol 1: Exosome isolation by precipitation

Basic Protocol 3: Analysis of exosomes by NanoSight nanoparticle tracking analysis

Alternate Protocol 2: Analysis of exosomes by flow cytometry and imaging flow cytometry

Basic Protocol 4: Downstream analysis of exosomes using high-performance liquid chromatography

Basic Protocol 5: Downstream analysis of the exosome proteome using nano-flow liquid chromatography–mass spectrometry

Neural crest cells constitute a unique population of progenitor cells with extensive stem cell capacities able to navigate throughout various environments in the embryo and are a source of multiple cell types, including neurons, glia, melanocytes, smooth muscles, endocrine cells, cardiac cells, and also skeletal and supportive tissues in the head. Neural crest cells are not restricted to the embryo but persist as well in adult tissues where they provide a reservoir of stem cells with great therapeutic promise. Many fundamental questions in cell, developmental, and stem cell biology can be addressed using this system. During the last decades there has been an increased availability of elaborated techniques, animal models, and molecular tools to tackle neural crest cell development. However, these approaches are often very challenging and difficult to establish and they are not adapted for rapid functional investigations of mechanisms driving cell migration and differentiation. In addition, they are not adequate for collecting pure populations of neural crest cells usable in large scale analyses and for stem cell studies. Transferring and adapting the neural crest system in tissue culture may then represent an attractive alternative, opening up numerous prospects. Here we describe a simple method for establishing primary cultures of neural crest cells derived from trunk neural tubes using the avian embryo as a source of cells. This protocol is suited for producing pure populations of neural crest cells that can be processed for cytological, cellular, and functional approaches aimed at characterizing their phenotype, behavior, and potential. © 2020 Wiley Periodicals LLC.

Basic Protocol: Primary cultures of avian trunk neural crest cells

Support Protocol: Adaptations for immunofluorescence labeling and videomicroscopy

Trafficking of intracellular cargo is essential to cellular function and can be defective in pathological states including cancer and neurodegeneration. Tools to quantify intracellular traffic are thus necessary for understanding this fundamental cellular process, studying disease mechanisms, and testing the effects of therapeutic pharmaceuticals. In this article we introduce an algorithm called QuoVadoPro that autonomously quantifies the movement of fluorescently tagged intracellular cargo. QuoVadoPro infers the extent of intracellular motility based on the variance of pixel illumination in a series of time-lapse images. The algorithm is an unconventional approach to the automatic measurement of intracellular traffic and is suitable for quantifying movements of intracellular cargo under diverse experimental paradigms. QuoVadoPro is particularly useful to measure intracellular cargo movement in non-neuronal cells, where cargo trafficking occurs as short movements in mixed directions. The algorithm can be applied to images with low temporal or spatial resolutions and to intracellular cargo with varying shapes or sizes, like mitochondria or endoplasmic reticulum: situations in which conventional methods such as kymography and particle tracking cannot be applied. In this article we present a stepwise protocol for using the QuoVadoPro software, illustrate its methodology with common examples, discuss critical parameters for reliable data analysis, and demonstrate its use with a previously published example. © 2020 Wiley Periodicals LLC.

Basic Protocol: QuoVadoPro, an autonomous tool for measuring intracellular dynamics using temporal variance

The movement of intracellular cargo, such as transcripts, proteins, and organelles, is fundamental to cellular function. Neurons, due to their long axons and dendrites, are particularly dependent on proper intracellular trafficking and vulnerable to defects in the movement of intracellular cargo that are noted in neurodegenerative and neurodevelopmental disorders. Accurate quantification of intracellular transport is therefore needed for studying the mechanisms of cargo trafficking, the influence of mutations, and the effects of potentially therapeutic pharmaceuticals. In this article, we introduce an algorithm called “Kymolyzer.” The algorithm can quantify intracellular trafficking along a defined path, such as that formed by the aligned microtubules of axons and dendrites. Kymolyzer works as a semi-autonomous kymography software application. It constructs and analyzes kymographs to measure the movement and distribution of fluorescently tagged objects along a user-defined path. The algorithm can be used under a wide variety of experimental conditions and can extract a diverse array of motility parameters describing intracellular movement, including time spent in motion, percentage of objects in motion, percentage of objects that are stationary, and velocities of motile objects. This article serves as a user manual describing the design of Kymolyzer, providing a stepwise protocol for its use and illustrating its functions with common examples. © 2020 Wiley Periodicals LLC

Basic Protocol: Kymolyzer, a semi-autonomous kymography tool to analyze intracellular motility