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.
