Current Protocols in Cytometry最新文献

Confocal microscopy has been an important imaging tool for life scientists for over 20 years. Early techniques focused on indirect staining processes that involved staining with an unconjugated primary antibody, followed by incubation with a secondary fluorescent antibody that would reveal and amplify the signal of the primary antibody. With more and more directly conjugated fluorescent primary antibodies becoming commercially available, staining with multiple fluorescent primary antibodies is now more frequent. To date, staining with up to three primary antibodies and a nuclear dye is widely practiced.

Here, we describe an important improvement to the standard polychromatic immunofluorescent staining protocol that allows the simultaneous detection of seven fluorescent parameters using a standard confocal laser scanning microscope with four laser lines and four photomultiplier tubes. By incorporating recently available tandem dyes that emit in the blue and violet regions of the visible light spectrum (Brilliant Blue and Brilliant Violet), we were able to differentiate several additional fluorochromes simultaneously. Due to the added complexity of 7-color immunofluorescent imaging, we developed a clear methodology to optimize antibody concentrations and simple guidelines on how to identify and correct non-specific signals. These are detailed in the following protocol. © 2019 by John Wiley & Sons, Inc.

Basic Protocol: 7-Color immunofluorescent staining protocol using directly conjugated antibodies

Support Protocol 1: Antibody titration protocol

Support Protocol 2: Spillover optimization protocol

Recent advances in analytical cytometry have improved diagnostic tools for the study of erythropoiesis in anemic patients and resolution of differential diagnosis in diseases of the erythron. This article presents three applications of red blood cell (RBC) analysis-quantitation of fetal red cells, F-cell enumeration, and F-reticulocyte analysis-which improve diagnostic precision, sensitivity, and specificity, and provide better laboratory indicators of therapeutic efficacy in a variety of hematologic and obstetric disorders. Such advances also include the measurement and quantitation of RBC hemoglobins and their relative ribonucleic acid levels. These advances not only promise to improve diagnostic accuracy and laboratory precision over techniques such as the traditional manual reticulocyte counting method and the Kleihauer-Betke stain method for evaluating fetomaternal hemorrhage (FMH), but also serve as tools for newer assays of anemia diagnosis and improved clinical outcomes. In addition to the primary methods, supporting techniques for preparing spiked controls, automating data analysis, setting up a fetal hemoglobin acquisition protocol, and assaying reticulocytes using thiazole orange are also presented. © 2019 by John Wiley & Sons, Inc.

Cell volume is an important parameter in studying cell adaptation to anisosmotic stress, activation of monovalent ion channels, and cell death. This article describes a method for measurement of the volumes of adherent cells using a standard light microscope. A coverslip with attached cells is placed in a shallow chamber in a medium containing a strongly absorbing and cell-impermeant dye, Acid Blue 9. When such a sample is imaged in transmitted light at a wavelength of maximum dye absorption (630 nm), the resulting contrast quantitatively reflects cell thickness; once the thickness is known at every point, the volume can be computed as well. Technical details, interpretation of data, and possible artifacts are discussed. © 2019 by John Wiley & Sons, Inc.

Cover: In Zhao et al. (https://doi.org/10.1002/cpcy.55), Detection of histone H2AX phosphorylated on Ser-139 (γH2AX) induced by topotecan (Tpt) or etoposide (Etp) in relation to cell position in the cell cycle. Bivariate cellular DNA content versus γH2AX Immunofluorescence (IF) distributions (scatterplots) are shown for TK6 cells left untreated (Ctrl) or treated in culture with 0.15 μM Tpt or 10 μM Etp for 2 hr, as previously described (Tanaka et al., 2007). See e55.

This unit describes immunocytochemical detection of histone H2AX phosphorylated on Ser-139 (γH2AX) to reveal DNA damage, particularly when the damage involves the presence of DNA double-strand breaks (DSBs). These breaks often result from DNA damage induced by ionizing radiation or by treatment with anticancer drugs such as DNA topoisomerase inhibitors. Furthermore, DSBs are generated in the course of DNA fragmentation during apoptosis. The unit presents strategies to distinguish radiation- or drug-induced DNA breaks from those intrinsically formed in untreated cells or associated with apoptosis. The protocol describes immunocytochemical detection of γH2AX combined with measurement of DNA content to identify cells that have DNA damage and concurrently to assess their cell-cycle phase. The detection is based on indirect immunofluorescence using FITC– or Alexa Fluor 488–labeled antibody, with DNA counterstained with propidium iodide and cellular RNA removed with RNase A. © 2019 by John Wiley & Sons, Inc.

Cover: In Eller et al. (https://doi.org/10.1002/cpcy.54), Hierarchical gates to evaluate fluorescent cells. Gates are applied identically to all samples. The P1 gate outlines the uniform population of HeLa cells for evaluation (typically >75% of total events). Under these specific conditions, events in the red region represent cellular debris or mechanically disrupted cells; if >50% of events are in this region it indicates an overall unhealthy sample. From the P1/cell population, single cells are identified by the P2 gate using their size distribution (typically >90% of P1). The P3/fluorescent population is derived from the P2/single cell population. The P3 fluorescent cell population is defined by a gate that excludes non-fluorescent cells in the untransfected sample (gray) and includes transfected cells (green, typically >50% of P2). There should be 10,000 events in P3 for an accurate evaluation. To calibrate the sensor in cells, an additional gate is required to identify the permeabilized and equilibrated cells. This P4 gate (typically >85% of P3) is derived from uniform, single cells in P2, and is delineated by equilibrated intracellular propidium iodide (PI). Fluorescence is then evaluated in P5, which is derived from P4. In this example, a gray density contour plot in the lower left quadrant represents cells that do no express either sensor or cpVenus. Cells that express either sensor or cpVenus populate P5, and the fluorescence of the populations treated with either equilibrated buffer (red) or 500 μM NAD+ (indigo) are overlaid. Data was collected on a BD Fortessa flow cytometer and analyzed on FlowJo V10 software. See e54.

Flow cytometry approaches combined with a genetically encoded targeted fluorescent biosensor are used to determine the subcellular compartmental availability of the oxidized form of nicotinamide adenine dinucleotide (NAD⁺). The availability of free NAD⁺ can affect the activities of NAD⁺-consuming enzymes such as sirtuin, PARP/ARTD, and cyclic ADPR-hydrolase family members. Many methods for measuring the NAD⁺ available to these enzymes are limited because they cannot determine free NAD⁺ as it exists in various subcellular compartments distinctly from bound NAD⁺ or NADH. Here, an approach to express the sensor in mammalian cells, monitor NAD⁺-dependent fluorescence intensity changes using flow cytometry approaches, and analyze data obtained is described. The benefit of flow cytometry approaches with the NAD⁺ sensor is the ability to monitor compartmentalized free NAD⁺ fluctuations simultaneously within many cells, which greatly facilitates analyses and calibration. © 2018 by John Wiley & Sons, Inc.