Guidelines for preparation and flow cytometry analysis of human nonlymphoid tissue DC

Abstract

This article is part of the Dendritic Cell Guidelines article series, which provides a collection of state-of-the-art protocols for the preparation, phenotype analysis by flow cytometry, generation, fluorescence microscopy, and functional characterization of mouse and human dendritic cells (DC) from lymphoid organs, and various nonlymphoid tissues. Within this article, detailed protocols are presented that allow for the generation of single-cell suspensions from human nonlymphoid tissues including lung, skin, gingiva, intestine as well as from tumors and tumor-draining lymph nodes with a subsequent analysis of dendritic cells by flow cytometry. Further, prepared single-cell suspensions can be subjected to other applications including cellular enrichment procedures, RNA sequencing, functional assays, etc. While all protocols were written by experienced scientists who routinely use them in their work, this article was also peer-reviewed by leading experts and approved by all co-authors, making it an essential resource for basic and clinical DC immunologists.

Keywords: dendritic cells; flow cytometry; nonlymphoid tissues; tumor; tumor‐draining lymph node.

Figure 1

Schematic overview of the lung. (A) Schematic view of the lung showing the trachea and bronchioles (light blue) and lung (pink) tissue. Immune cell populations, such as dendritic cells, monocytes, macrophages, and tissue‐resident T cells can be spread out across the entire lung and are partially found in the airways as well. (B) Schematic view of the bronchoalveolar niche, depicting the alveolar and interstitial niches. DC and monocytes can be found spread across the entire bronchoalveolar niche, while alveolar and interstitial macrophages are refined to their specific niche.

Figure 2

Flow cytometric analyses of human lung DC subsets. As depicted, human lung samples are pregated on single, live, CD45⁺ cells. The CD45⁺ population of the lung then is further gated on CD3⁻ and CD16/CD19/CD20⁻ cells (also called LIN⁻), before gating on all HLA‐DR⁺ cells. From here, monocytes are excluded by gating on CD88⁻ cells only and CD14⁻CD88⁻ and CD14⁺ CD88⁻ are used for subsequent gating on the different DC subsets. Within the CD88⁻ population, pDC can be identified as CD123⁺CD169⁻ cells and pre‐DC as CD123^int/+CD169⁺ cells. Subsequently, the CD123/CD169 double negative population is further gated on CD141⁺ cDC1 (optional also CADM1⁺) or CD1c⁺ cells, which contain a mixture of cDC2 (DC2 and DC3). These CD1c⁺ cDC2 can be subdivided using CD5 and CD14 and by gating on the CD5⁺CD14⁻ DC2, CD5⁻CD14⁻ DC3, and CD5⁻CD14⁺ DC3 subsets. Samples shown here were acquired on a CYTEK Aurora 5L.

Figure 3

Gating strategy for flow cytometry analysis of human skin. The skin was enzymatically digested to generate single‐cell suspensions. (A) Gating strategy for viable CD45⁺ cells after exclusion of cellular debris, doublets, and dead cells. (B) CD3⁺ T cells and CD19⁺ B cells as well as CD14⁺ cells were excluded before DC characterization. (C) HLA‐DR expressing cells comprise DC, which can be further subdivided into CD141⁺ CD1c⁺ cDC1 and CD1c⁺ subsets, consisting of CD207⁺ LC and CD207⁻ cDC2. (D) Surface expression of co‐stimulatory markers CD40, CD80, and CD86 on the indicated DC subsets: cDC1 (orange), cDC2 (grey), and LC (blue). MFI values are shown. cDC, conventional DC; LC, Langerhans cells.

Figure 4

Dissection of the gingiva from the masticatory mucosa. A buccal view of the upper right molars, premolars, and an incision in the gingiva. The gingiva is part of the masticatory mucosa that surrounds the teeth. Using a sharp No. 15c scalpel blade, dissect a sample of fresh gingiva that includes the lamina propria and epithelium. The minimal sample size is 3 × 2 mm (∼15 mg). In this case, a 15 × 2 mm tissue sample was harvested (black dashed line).

Figure 5

Processing the fresh gingival sample. (1) After dissecting the gingival sample, store the sample in 3 mL of FACS buffer (see the section  5 ) in a 15 mL tube. (2) Clean the blood from the fresh sample, dry it carefully, and (3) weigh the tissue. (4) Place the sample in a six‐well plate. If the sample is bigger than 3 × 2 mm, divide it into 3 × 2 mm pieces and place each piece in a separate well. Chop the tissue into tiny pieces (smaller than 1 × 1 mm) using a no. 10 scalpel blade to allow better contact between the tissue and the digestion mix (see the section  5 ). Add 1000 µL of digestion mix to each well.

Figure 6

Gating strategy for the characterization of dendritic cell subsets in human gingiva. (i) Cells are pregated according to their size and granularity (FSC‐A/SSC‐A) to exclude debris and dead cells. (ii, iii) Doublets are excluded by gating on the FSC and SSC area (A) and height (H). (iv) Red blood cells are excluded by using SSC‐B‐H and SSC‐H. (v) Dead cells are excluded using Zombie staining. (vi) CD45⁺ hematopoietic cells are selected. (vii) CD3⁺ T cells and CD19⁺ B cells are excluded. (viii) CD66b⁺ neutrophils and CD56⁺ NK cells are excluded. (ix) HLA‐DR⁺ cells are selected. (x) CD123⁺, CD45RA⁺, and CD123‐CD45RA^+∖− cells are further gated, which represent the pre‐DC, pDC, and DC subpopulations, respectively. (xi) CD5⁺AXL⁺ cells represent the pre‐DC subpopulation and CD5‐AXL‐ cells constitute the pDC subpopulation. (xii) LC cells are isolated from other DC cells by their expression of both CD207 and Epcam. (xiii) Conventional DC (cDC) are divided into cDC1 and cDC2 according to their CD141 and CD1c expression. While cDC1 are positive for CD141 and negative for CD1c, cDC2 are positive for CD1a and negative for CD141. Data acquisition was performed on a Cytek Aurora Flow Cytometry System and data was analyzed using FlowJo V10.8.1 software.

Figure 7

Illustration of immune compartments that can be isolated from the gut wall of small and large human intestines. The human gut wall can be processed and then peeled apart under a dissecting microscope to isolate the mucosa from the submucosa. From the mucosa, it is then possible to isolate the epithelium, lamina propria, and mucosal ILF. Small intestinal Peyer's patches can be macroscopically identified and removed from surrounding tissue, after which individual follicles can be stained and isolated using methylene blue. By staining the submucosa of the large intestine with methylene blue, individual submucosal ILF can be isolated.

Figure 8

Flow cytometry analysis of cDC subsets isolated from different compartments of the human intestine, showing (A) pregating used for all compartments and (B) plasmacytoid DC and conventional DC subset gating, with slight gating variations for each compartment. Numbers in histograms represent the median fluorescence intensity of CD103 for DC1 (red) and DC2 (blue), dashed line delineates positive from negative CD103 signal. LP, lamina propria; MLN, mesenteric lymph nodes; PP, Peyer's patch; SM‐ILF, submucosal isolated lymphoid follicles. Representative data of 3–10 colorectal cancer patients per compartment, with tissue taken from unaffected areas.

Figure 9

cDC gating strategy in the flow cytometric analysis of a single‐cell suspension derived from a human colorectal primary carcinoma. (A) Gating strategy for the quantitation and phenotypic analysis of myeloid APC/cDC subsets among live CD45⁺ leukocytes. (B) Expression of subset‐defining and co‐stimulatory markers as well as PD‐L1 on the identified APC/cDC subsets. Histogram overlays with fluorescence‐minus‐one controls are shown.

Figure 10

cDC gating strategy in the flow cytometric analysis of a single‐cell suspension derived from non‐small‐cell lung carcinoma. (A) Gating strategy for the quantitation and phenotypic analysis of myeloid APC/cDC subsets among CD45⁺ leukocytes. (B) Expression of subset‐defining and co‐stimulatory markers as well as PD‐L1 on the identified APC/cDC subsets. Histogram overlays with fluorescence‐minus‐one controls are shown.

Figure 11

cDC gating strategy in the flow cytometric analysis of a single‐cell suspension derived from melanoma metastasis. (A) Gating strategy for the quantitation and phenotypic analysis of myeloid APC/cDC subsets among live CD45⁺ leukocytes. (B) Expression of subset‐defining and co‐stimulatory markers as well as PD‐L1 on the identified APC/cDC subsets. Histogram overlays with fluorescence‐minus‐one controls are shown.

Figure 12

cDC subset gating strategy in Melanoma TDLN. (A) Gating strategy for the quantitation and phenotypic analysis of two migratory and two LNR‐cDC subsets among live CD45⁺ leukocytes. (B) Expression of activation and co‐stimulatory markers as well as immune checkpoints on the identified cDC subsets. Histogram overlays with fluorescence‐minus‐one controls are shown.

Figure 13

cDC subset gating strategy in mammary TDLN. (A) Gating strategy for the quantitation and phenotypic analysis of two migratory and two LNR‐cDC subsets among live CD45⁺ leukocytes. (B) Expression of activation and co‐stimulatory markers as well as immune checkpoints on the identified cDC subsets. Histogram overlays with fluorescence‐minus‐one controls are shown.

Figure 14

cDC subset gating strategy in non‐small‐cell lung cancer TDLN. (A) Gating strategy for the quantitation and phenotypic analysis of cDC subsets among live CD45⁺ leukocytes. Note the absence of detectable CD1a⁺ migratory subsets. (B) Expression of activation and co‐stimulatory markers as well as immune checkpoints on the identified cDC subsets. Shown are histogram overlays with fluorescence‐minus‐one controls.