Visualizing fluorescence‐tagged molecules is a powerful strategy that can reveal the complex dynamics of the cell. One robust and broadly applicable method is immunofluorescence microscopy, in which a fluorescence‐labeled antibody binds the molecule of interest and then the location of the antibody is determined by fluorescence microscopy. The effective application of this technique includes several considerations, such as the nature of the antigen, specificity of the antibody, permeabilization and fixation of the specimen, and fluorescence imaging of the cell. Although each protocol will require fine‐tuning depending on the cell type, antibody, and antigen, there are steps common to nearly all applications. This article provides protocols for staining the cytoskeleton and organelles in two very different kinds of cells: flat, adherent fibroblasts and thick, free‐swimming
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
- NSF-PAR ID:
- 10175118
- Publisher / Repository:
- Wiley Blackwell (John Wiley & Sons)
- Date Published:
- Journal Name:
- Current Protocols in Cell Biology
- Volume:
- 88
- Issue:
- 1
- ISSN:
- 1934-2500
- Format(s):
- Medium: X
- Sponsoring Org:
- National Science Foundation
More Like this
-
Abstract Tetrahymena cells. Additional protocols enable visualization with widefield, laser scanning confocal, and eSRRF super‐resolution fluorescence microscopy. © 2023 Wiley Periodicals LLC.Basic Protocol 1 : Immunofluorescence staining of adherent cells such as fibroblastsBasic Protocol 2 : Immunofluorescence of suspension cells such asTetrahymena Basic Protocol 3 : Visualizing samples with a widefield fluorescence microscopeAlternate Protocol 1 : Staining suspension cells adhered to poly‐l ‐lysine‐coated coverslipsAlternate Protocol 2 : Visualizing samples with a laser scanning confocal microscopeAlternate Protocol 3 : Generating super‐resolution images with SRRF microscopy -
Abstract Intracellular signaling processes are frequently based on direct interactions between proteins and organelles. A fundamental strategy to elucidate the physiological significance of such interactions is to utilize optical dimerization tools. These tools are based on the use of small proteins or domains that interact with each other upon light illumination. Optical dimerizers are particularly suitable for reproducing and interrogating a given protein‐protein interaction and for investigating a protein's intracellular role in a spatially and temporally precise manner. Described in this article are genetic engineering strategies for the generation of modular light‐activatable protein dimerization units and instructions for the preparation of optogenetic applications in mammalian cells. Detailed protocols are provided for the use of light‐tunable switches to regulate protein recruitment to intracellular compartments, induce intracellular organellar membrane tethering, and reconstitute protein function using enhanced Magnets (eMags), a recently engineered optical dimerization system. © 2021 Wiley Periodicals LLC.
This article was corrected on 25 July 2022. See the end of the full text for details.
Basic Protocol 1 : Genetic engineering strategy for the generation of modular light‐activated protein dimerization unitsSupport Protocol 1 : Molecular cloningBasic Protocol 2 : Cell culture and transfectionSupport Protocol 2 : Production of dark containers for optogenetic samplesBasic Protocol 3 : Confocal microscopy and light‐dependent activation of the dimerization systemAlternate Protocol 1 : Protein recruitment to intracellular compartmentsAlternate Protocol 2 : Induction of organelles’ membrane tetheringAlternate Protocol 3 : Optogenetic reconstitution of protein functionBasic Protocol 4 : Image analysisSupport Protocol 3 : Analysis of apparent on‐ and off‐kineticsSupport Protocol 4 : Analysis of changes in organelle overlap over time -
Abstract Immune cell signaling is largely regulated by protein phosphorylation. Stimulation of toll‐like receptors (TLRs) by pathogen‐associated ligands drives the cascade of immune response, which can be influenced by differences in phosphoprotein abundance. Therefore, the analysis of phosphorylation signatures at a global level is central to understanding the complex and integrated signaling in macrophages upon pathogen attack. Here, we describe a mass spectrometry‐based approach to identify and quantify phosphoproteome changes in response to the stimulation of TLR2, TLR4, and TLR7 with immune‐response inducing ligands in cultured immune cells. This review will focus on the TLR stimulation of mouse macrophages as an example; however, the technique is applicable to any immortalized immune cell and any soluble stimuli. The methodology includes protocols for metabolic labeling of immune cells (stable isotope labeling of amino acids in cell culture, i.e., SILAC); ligand‐initiated stimulation of immune receptors followed by cell lysis; in‐solution trypsin digestion of proteins and enrichment of the resulting peptide mix for collecting phosphopeptides, which are then analyzed by high‐resolution LC‐MS/MS (liquid‐chromatography tandem mass spectrometry). Published 2020. U.S. Government.
Basic Protocol 1 : SILAC labeling of mouse macrophagesBasic Protocol 2 : Stimulation, cell lysis and Western BlottingBasic Protocol 3 : Trypsin digestion, fractionation and phosphopeptide enrichmentBasic Protocol 4 : Quantitative mass spectrometryAlternate Protocol : Culturing SILAC‐labeled cells from frozen mouse macrophages cells -
Abstract Microscopic and spectroscopic techniques are commonly applied to study microbial cells but are typically used on separate samples, resulting in population-level datasets that are integrated across different cells with little spatial resolution. To address this shortcoming, we developed a workflow that correlates several microscopic and spectroscopic techniques to generate an in-depth analysis of individual cells. By combining stable isotope probing (SIP), fluorescence in situ hybridization (FISH), scanning electron microscopy (SEM), confocal Raman microspectroscopy (Raman), and nano-scale secondary ion mass spectrometry (NanoSIMS), we illustrate how individual cells can be thoroughly interrogated to obtain information about their taxonomic identity, structure, physiology, and metabolic activity. Analysis of an artificial microbial community demonstrated that our correlative approach was able to resolve the activity of single cells using heavy water SIP in conjunction with Raman and/or NanoSIMS and establish their taxonomy and morphology using FISH and SEM. This workflow was then applied to a sample of yet uncultured multicellular magnetotactic bacteria (MMB). In addition to establishing their identity and activity, backscatter electron microscopy (BSE), NanoSIMS, and energy-dispersive X-ray spectroscopy (EDS) were employed to characterize the magnetosomes within the cells. By integrating these techniques, we demonstrate a cohesive approach to thoroughly study environmental microbes on a single-cell level.
-
Abstract Characterizing the mechanical properties of single cells is important for developing descriptive models of tissue mechanics and improving the understanding of mechanically driven cell processes. Standard methods for measuring single‐cell mechanical properties typically provide isotropic mechanical descriptions. However, many cells exhibit specialized geometries
in vivo , with anisotropic cytoskeletal architectures reflective of their function, and are exposed to dynamic multiaxial loads, raising the need for more complete descriptions of their anisotropic mechanical properties under complex deformations. Here, we describe the cellular microbiaxial stretching (CμBS) assay in which controlled deformations are applied to micropatterned cells while simultaneously measuring cell stress. CμBS utilizes a set of linear actuators to apply tensile or compressive, short‐ or long‐term deformations to cells micropatterned on a fluorescent bead‐doped polyacrylamide gel. Using traction force microscopy principles and the known geometry of the cell and the mechanical properties of the underlying gel, we calculate the stress within the cell to formulate stress‐strain curves that can be further used to create mechanical descriptions of the cells, such as strain energy density functions. © 2022 Wiley Periodicals LLC.Basic Protocol 1 : Assembly of CμBS stretching constructsBasic Protocol 2 : Polymerization of micropatterned, bead‐doped polyacrylamide gel on an elastomer membraneSupport Protocol : Cell culture and seeding onto CμBS constructsBasic Protocol 3 : Implementing CμBS stretching protocols and traction force microscopyBasic Protocol 4 : Data analysis and cell stress measurements