The lung is a structurally and immunologically complex organ, constantly exposed to airborne microbes, allergens, and pollutants. Understanding how diverse pulmonary immune cells respond to these challenges is critical for advancing respiratory disease research and identifying appropriate therapeutic interventions. Flow cytometry remains a cornerstone of immune profiling, and advances in high-parameter spectral cytometry have significantly expanded its analytical capabilities. However, challenges, such as poor tissue dissociation, spectral overlap, and loss of spatial information, can hinder the comprehensive interpretation of the lung environment. To address these limitations, we developed an optimized spectral flow cytometry platform, utilizing 5-laser Cytek Aurora spectral cytometers, for deep immunophenotyping of murine lung tissue. Our approach integrates in vivo CD45 antibody labeling-administered intravenously and oropharyngeally-to distinguish circulating, airway, and interstitial immune populations, preserving spatial context in single-cell suspensions. We utilize complementary 25+ parameter panels targeting myeloid and lymphoid compartments, built on a shared backbone to enable consistent classification across datasets. Refined tissue processing protocols support optimal recovery of representative lung cell populations, and overnight intracellular staining enhances marker resolution. Using this platform, we reliably resolve stromal, endothelial, and epithelial cells alongside immune subsets-including macrophages, monocytes, dendritic cells, eosinophils, neutrophils, T and B cells, innate lymphoid cells (ILCs), and natural killer (NK) cells-subclassified by activation, function, and tissue residency. Validation with an influenza A virus model confirmed expected dynamic immune responses and revealed previously unrecognized populations. This spatially informed approach enables high-resolution interrogation of pulmonary immunity in health and disease. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Differential in vivo labeling for spatial profiling of pulmonary immune cells by spectral cytometry Basic Protocol 2: Preparation of single-cell suspensions from murine lung tissue for spectral cytometric analysis of immune cell populations Basic Protocol 3: Optimized workflow and spectral flow cytometry panels for profiling of pulmonary immune cell populations in a single-cell suspension.
Use of flow cytometry to analyze small particles has been implemented for several decades. More recently, small particle analysis has become increasingly utilized owing to the increased sensitivity of conventional and commercially available flow cytometers along with growing interest in small particles such as extracellular vesicles (EVs). Despite an increase in small particle flow cytometry utilization, a lack of standardization in data reporting has resulted in a growing body of literature regarding EVs that cannot be easily interpreted, validated, or reproduced. Methods for fluorescence and light scatter standardization are well established, and the reagents to perform these analyses are commercially available. Here, we describe FCMPASS , a software package for performing fluorescence and light scatter calibration of small particles while generating standard reports conforming to the MIFlowCyt-EV standard reporting framework. This article covers the workflow of implementing calibration using FCMPASS as follows: acquisition of fluorescence and light scatter calibration materials, cataloguing the reference materials for use in the software, creating cytometer databases and datasets to associate calibration data and fcs files, importing fcs files for calibration, inputting fluorescence calibration parameters, inputting light scatter calibration parameters, and applying the calibration to fcs files. Published 2020. U.S. Government. Basic Protocol 1: Acquisition and gating of light scatter calibration materials Basic Protocol 2: Acquisition and gating of fluorescence calibration materials Alternate Protocol: Cross-calibration of fluorescence reference materials Basic Protocol 3: Cataloguing light scatter calibration materials Basic Protocol 4: Cataloguing fluorescence calibration materials Basic Protocol 5: Creating cytometer databases and datasets Basic Protocol 6: Importing fcs files Basic Protocol 7: Fluorescence calibration Basic Protocol 8: Light scatter calibration Basic Protocol 9: Performing and reporting fcs file calibration.
In light microscopy, illuminating light is passed through the sample as uniformly as possible over the field of view. For thicker samples, where the objective lens does not have sufficient depth of focus, light from sample planes above and below the focal plane will also be detected. The out-of-focus light will add blur to the image, reducing the resolution. In fluorescence microscopy, any dye molecules in the field of view will be stimulated, including those in out-of-focus planes. Confocal microscopy provides a means of rejecting the out-of-focus light from the detector such that it does not contribute blur to the images being collected. This technique allows for high-resolution imaging in thick tissues. In a confocal microscope, the illumination and detection optics are focused on the same diffraction-limited spot in the sample, which is the only spot imaged by the detector during a confocal scan. To generate a complete image, the spot must be moved over the sample and data collected point by point. A significant advantage of the confocal microscope is the optical sectioning provided, which allows for 3D reconstruction of a sample from high-resolution stacks of images. Several types of confocal microscopes have been developed for this purpose, and each has different advantages and disadvantages. This article provides a concise introduction to confocal microscopy. © 2019 by John Wiley & Sons, Inc.
Recent advances in high resolution flow cytometry (HRFC), which show improvements in both light scatter and fluorescence sensitivity have resulted in the development of techniques that better isolate, stain and analyze single EVs(Boing et al., 2014; Groot Kormelink et al., 2016; Morales-Kastresana, Musich, Welsh, Telford, Demberg, Wood, Bigos, Ross, Kachynski, Dean, Feton, et al., 2019; Morales-Kastresana et al., 2017; Stoner et al., 2016; van der Vlist, Nolte-'t Hoen, Stoorvogel, Arkesteijn, & Wauben, 2012). Below, we describe protocols to fluorescently label EVs using CFDA-SE (hereinafter called CFSE), as well as antibodies targeted at specific EV surface proteins. We also provide guidelines for residual dye and antibody removal, appropriate data acquisition by HRFC and EV counting by HRFC. Figure 1summarizes these protocols. The EVs used in this protocol are derived from the DC2.4 cell line, and bone marrow derived dendritic cells (BMDCs). The DC2.4 cell line are immature dendritic cells (DCs) with very low expression of typical DC markers on their surface (unpublished observation and(Hargadon, Forrest, & Reddy, 2012)) and that release a morphologically homogeneous population of EVs (~130 nm in diameter). DC2.4 EVs will be used to demonstrate a CFSE staining in Basic Protocol 1, as well as being used as a negative control for antibody-based staining methods (Basic Protocol 2). Bone marrow dendritic cell (BMDC)-derived EVs are more heterogeneous in diameter (100-200 nm) and composition(Morales-Kastresana, Musich, Welsh, Telford, Demberg, Wood, Bigos, Ross, Kachynski, Dean, Felton, et al., 2019), and express DC markers such as MHC-II. BMDC EVs will be used to demonstrate antigen-specific staining with fluorochrome-conjugated antibodies in Basic Protocol 2. DC2.4 and BMDC-derived EVs were isolated by serial ultracentrifugation, with concentration and diameter distribution characterized by NTA, as described before(Morales-Kastresana, Musich, Welsh, Telford, Demberg, Wood, Bigos, Ross, Kachynski, Dean, Feton, et al., 2019; Morales-Kastresana et al., 2017).
Live imaging is critical to determining the dynamics and spatial interactions of cells within the tissue environment. In the lung, this has proven to be difficult due to the motion brought about by ventilation and cardiac contractions. A previous version of this Current Protocols in Cytometry article reported protocols for imaging ex vivo live lung slices and the intact mouse lung. Here, we update those protocols by adding new methodologies, new approaches for quantitative image analysis, and new areas of potential application. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Live imaging of lung slices Support Protocol 1: Staining lung sections with fluorescent antibodies Basic Protocol 2: Live imaging in the mouse lung Support Protocol 2: Intratracheal instillations Support Protocol 3: Intravascular instillations Support Protocol 4: Monitoring vital signs of the mouse during live lung imaging Support Protocol 5: Antibodies Support Protocol 6: Fluorescent reporter mice Basic Protocol 3: Quantification of neutrophil-platelet aggregation in pulmonary vasculature Basic Protocol 4: Quantification of platelet-dependent pulmonary thrombosis Basic Protocol 5: Quantification of pulmonary vascular permeability.
Direct visualization of higher-order chromatin structure at the molecular scale is of great importance for understanding the impact of chromatin organization on gene expression in many biological processes. Understanding the changes in chromatin structure during pathological processes requires the use of in vivo models and clinical samples, and formalin-fixed, paraffin-embedded (FFPE) tissue is the most widespread form of preservation. Here we describe the details of PathSTORM, an optimized stochastic optical reconstruction microscopy (STORM) protocol for high-quality super-resolution imaging of densely packed higher-order chromatin organization in pathological tissue. We discuss detailed methods for fluorescence staining of DNA and histone proteins, as well as the key technical factors for obtaining high-quality STORM images in pathological tissue samples. © 2020 Wiley Periodicals LLC Basic Protocol 1: Fluorescence staining of chromatin in pathological tissue Basic Protocol 2: STORM data processing Support Protocol 1: Drift correction Support Protocol 2: Image reconstruction Support Protocol 3: Hematoxylin & eosin (H&E) staining.
Translational research has improved the diagnosis and follow-up of hematological diseases and malignancies. However, some classical diagnostics used for research and clinical practice that have remain practically unchanged for decades may be better addressed through advances in flow cytometry technology, whereby more precise measurements may be implemented in a straightforward manner. The current method for semiquantitative analysis of granulocytic alkaline phosphatase (GAP) activity is still based on observer-dependent color-intensity classification. Here, we describe a novel strategy for flow cytometric quantification of GAP activity in which staining and analytical flow cytometry facilitate the detection and quantification of subpopulations of leukocytes with different GAP activities. Our experiments demonstrate the potential of flow cytometry as a simple and highly sensitive approach for measuring GAP activity in unlysed whole blood. Notably, a comparison of flow cytometry and enzyme cytochemistry techniques showed that enzyme activity scores were not similar, indicating that results needs to be interpreted with caution, given that the enzyme-substrate binding affinities may differ, as well as the subjective evaluation of the intensity of the precipitated dye. © 2020 Wiley Periodicals LLC. Basic Protocol: Protocol preparation, sample acquisition, and gating strategy for flow cytometric identification of alkaline phosphatase activity in granulocytes from whole blood samples Support Protocol 1: Sample preparation for granulocyte alkaline phosphatase determination by flow cytometry using no-lyse no-wash methods Support Protocol 2: Data analysis and formula to calculate the GAP score.
SARS-CoV-2 is a novel coronavirus that causes the acute respiratory disease-Coronavirus disease 2019 (COVID-19)-which has led to a global health crisis. Currently, no prophylactics or therapies exist to control virus spread or mitigate the disease. Thus, the risk of infection for physicians and scientists is high, requiring work to be conducted in Biosafety Level-3 (BSL-3) facilities if virus will be isolated or propagated. However, inactivation of the virus can enable safe handling at a reduced biosafety level, making samples accessible to a diverse array of institutions and investigators. Institutions of all types have an immediate need for guidelines that outline safe collection, handling, and inactivation of samples suspected to contain active virus. Here we provide a practical guide for physicians and researchers wishing to work with materials from patients who are COVID-19 positive or suspected positive. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Practical guidelines for the safe collection and handling of specimens collected from COVID-19 and suspected COVID-19 patients Basic Protocol 2: Inactivating SARS-CoV-2.
Flow cytometry allows the visualization of physical, functional, and/or biological properties of cells including antigens, cytokines, size, and complexity. With increasingly large flow cytometry panels able to analyze up to 50 parameters, there is a need to standardize flow cytometry protocols to achieve high-quality data that can be input into analysis algorithms. Without this clean data, algorithms may incorrectly categorize the cell populations present in the samples. In this protocol, we outline a comprehensive methodology to prepare samples for polychromatic flow cytometry. The use of multiple washing steps and rigorous controls creates high-quality data with good separation between cell populations. Experimental data acquired using this protocol can be analyzed via computational algorithms that perform end-to-end analysis. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Preparation of single-cell suspension for flow cytometry Support Protocol 1: Lung preparation Support Protocol 2: Counting cells on a flow cytometer Basic Protocol 2: Surface and intracellular flow cytometry staining Support Protocol 3: Single-color bead controls.
Red blood cell biomechanics can provide us with a deeper understanding of macroscopic physiology and have the potential of being used for diagnostic purposes. In diseases like sickle cell anemia and malaria, reduced red blood cell deformability can be used as a biomarker, leading to further assays and diagnoses. A microfluidic system is useful for studying these biomechanical properties. We can observe detailed red blood cell mechanical behavior as they flow through microcapillaries using high‐speed imaging and microscopy. Microfluidic devices are advantageous over traditional methods because they can serve as high‐throughput tests. However, to rapidly analyze thousands of cells, there is a need for powerful image processing tools and software automation. We describe a workflow process using Image‐Pro to identify and track red blood cells in a video, take measurements, and export the data for use in statistical analysis tools. The information in this protocol can be applied to large‐scale blood studies where entire cell populations need to be analyzed from many cohorts of donors. © 2020 The Authors.
The identification of residual leukemia following therapy, termed minimal or measurable residual disease (MRD), has emerged as one of the most important prognostic factors for patients with acute leukemia, including acute myeloid leukemia (AML). Flow cytometry is a preferred method for MRD detection due to its general applicability and the rapid results that it makes available. In this article, the basic protocol outlines a simple and efficient method for the labeling of hematopoietic cells from bone marrow or peripheral blood with a panel of monoclonal antibodies designed both to highlight patterns of normal maturation and allow identification of neoplastic hematopoietic progenitor populations with a high degree of sensitivity and specificity. The method was developed in a clinical laboratory setting for the diagnosis of myeloid stem cell disorders and neoplasms, and has been extensively validated both technically and clinically for the detection of MRD in AML. © 2020 The Authors. Basic Protocol: Staining and flow cytometry for AML minimal residual disease detection Support Protocol: Analysis and interpretation of data for AML minimal residual disease detection.
This article presents a single experiment designed to introduce a trainee to multiple advanced bench and analysis techniques, including high-dimensional cytometry, profiling cell signaling networks, functional assays with primary human tissue, and single-cell analysis with machine learning tools. The trainee is expected to have only minimal laboratory experience and is not required to have any prior training in flow cytometry, immunology, or data science. This article aims to introduce the advanced research areas with a design that is robust enough that novice trainees will succeed, flexible enough to allow some project customization, and fundamental enough that the skills and knowledge gained will provide a template for future experiments. For advanced users, the updated phospho-flow protocol and the established controls, best practices, and expected outcomes presented here also provide a framework for adapting these tools in new areas with unexplored biology. © 2020 by John Wiley & Sons, Inc. Basic Protocol: Phospho-protein stimulation and mass cytometry data collection Support Protocol: Analysis of signaling mass cytometry data.
Technological advances in fluorescence flow cytometry and an ever-expanding understanding of the complexity of the immune system has led to the development of large 20+ flow cytometry panels. Yet, as panel complexity and size increases, so does the difficulty involved in designing a high-quality panel, accessing the instrumentation capable of accommodating large numbers of parameters, and in analysing such high-dimensional data. A recent advancement is spectral flow cytometry, which in contrast to conventional flow cytometry distinguishes the full emission spectrum of each fluorochrome across all lasers, rather than identifying only the peak of emission. Fluorochromes with a similar emission maximum but distinct off-peak signatures can therefore be accommodated within the same flow cytometry panel, allowing greater flexibility in terms of panel design and fluorophore detection. Here, we highlight the specific characteristics regarding spectral flow cytometry and aim to guide users through the process of building, designing and optimising high-dimensional spectral flow cytometry panels using a comprehensive step-by-step protocol. Special considerations are also given for using highly-overlapping dyes and a logical selection process an optimal marker-fluorophore assignment is provided.
Understanding how immune cells respond to external stimuli such as pathogens or drugs is a key component of biomedical research. Critical to the immune response are the expression of cell-surface receptors and the secretion of cytokines, which are tightly regulated by gene expression and protein synthesis. Previously, cytokine mRNA expression levels have been measured from bulk analysis of heterogeneous or sorted cell populations, and the correlation between cytokine mRNA expression and protein levels using these techniques can be highly variable. Flow cytometry is used to monitor changes in cell-surface and intracellular proteins, but some proteins such as cytokines may be transient and difficult to measure. Thus, a flow cytometry method that can simultaneously measure cytokine mRNA and protein levels in single cells is a very powerful tool. We defined a flow cytometry method that combines the conventional measurement of T cell surface proteins (CD45, CD3, CD4, CD8) and intracellular cytokines (IL-2, INF-γ) with fluorescent in situ hybridization and branched DNA technology for amplification and detection of IL-2 and INF-γ mRNA transcripts in activated T cells. This method has been applied to frozen peripheral mononuclear blood cells (PBMCs) and frozen blood samples, making it applicable to clinical trial specimens that require shipment to the test site. In CD4+ cells from activated PBMCs, the concordance between mRNA and protein levels was 41% for IL-2 and 21% for and INF-γ. In CD8+ cells from activated PBMCs, the concordance was 15% for IL-2 and 32% for INF-γ. © 2020 by John Wiley & Sons, Inc. Basic Protocol: Detection of IL-2 and IFN-γ mRNA and protein expression in frozen PBMCs Alternate Protocol: Detection of IL-2 and IFN-γ mRNA and protein expression in frozen blood.
AbstractFor microorganisms in particular, viability is a term that is difficult to define and a state consequently difficult to measure. The traditional (and gold‐standard) usage equates viability and culturability (i.e., the ability to multiply), but the process of determining culturability is often too slow. Flow cytometry provides the opportunity to make rapid and quantitative measurements of dye uptake in large numbers of cells, and we can therefore exploit the flow cytometric approach to evaluate so‐called viability stains and to develop protocols for more routine assessments of microbial viability. This unit is primarily commentary, but several basic protocols have been included to ensure that users have a firm basis for attempting these reasonably difficult assays on traditional flow cytometer instruments. What is clear is that each assay must be carefully validated with the particular microorganism of interest before being applied in any research, clinical, or service form.
Cell volume is an important parameter in cell adaptation to anisosmotic stress, in the development of apoptosis and necrosis, and in the pathogenesis of several diseases. This unit describes a method for measuring the volume 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. Measurements in absolute units require knowledge of the absorption coefficient, and a similar procedure for the measurement of absorption coefficient is described.
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.
The susceptibility of DNA in situ to denaturation is modulated by its interactions with histone and nonhistone proteins, as well as with other chromatin components related to the maintenance of the 3D nuclear structure. Measurement of DNA proclivity to denature by cytometry provides insight into chromatin structure and thus can be used to recognize cells in different phases of the cell cycle, including mitosis, quiescence (G0 ), and apoptosis, as well as to identify the effects of drugs that modify chromatin structure. Particularly useful is the method's ability to detect chromatin changes in sperm cells related to DNA fragmentation and infertility. This article presents a flow cytometric procedure for assessing DNA denaturation based on application of the metachromatic property of acridine orange (AO) to differentially stain single- versus double-stranded DNA. This approach circumvents limitations of biochemical methods of examining DNA denaturation, in particular the fact that the latter destroy higher orders of chromatin structure and that, being applied to bulk cell populations, they cannot detect heterogeneity of individual cells. Because the metachromatic properties of AO have also found application in other cytometric procedures, such as differential staining of RNA versus DNA and assessment of lysosomal proton pump including autophagy, to avert confusion between these approaches and the use of this dye in the DNA denaturation assay, these AO applications are briefly outlined in this unit as well. © 2019 by John Wiley & Sons, Inc. Basic Protocol: Differential staining of single- versus double-stranded DNA with acridine orange.
Half of the patients with acute myeloid leukemia (AML), who achieve complete remission after chemotherapy treatment, will ultimately experience a relapse. Measurable residual disease (MRD) is an important post-treatment risk factor in AML, because it gives additional information about the depth of the remission. Within MRD, the small population of leukemic stem cells (LSCs) is thought to be at the base of the actual relapse. In this protocol, the flow cytometric detection of MRD and LSCs herein is outlined. We give a detailed overview of the sampling procedures for optimal multiparameter flow cytometry assessment of both MRD and LSC, using leukemia associated immunophenotypes (LAIPs) and LSC markers. Moreover, an overview of the gating strategies to detect LAIPs and LSC markers is provided. This protocol serves as guidance for flow cytometric detection of measurable residual (stem cell) disease necessary for proper therapeutic decision making in AML patients. © 2019 The Authors. Basic Protocol 1: Immunophenotypic LAIP detection for measurable residual disease monitoring Basic Protocol 2: Immunophenotypic detection of CD34+CD38- leukemic stem cells.
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.
