Guidelines for DC preparation and flow cytometric analysis of human lymphohematopoietic tissues

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 non‐lymphoid tissues. Within this article, detailed protocols are presented that allow for the generation of single cell suspensions from human lymphohematopoietic tissues including blood, spleen, thymus, and tonsils with a focus on the subsequent analysis of DC via flow cytometry, as well as flow cytometric cell sorting of primary human DC. Further, prepared single cell suspensions as well as cell sorter‐purified DC can be subjected to other applications including cellular enrichment procedures, RNA sequencing, functional assays, and many more. 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.

Dendritic cells (DC) act as central regulators of T cell immunity orchestrating the induction of T cell responses or the maintenance of peripheral tolerance [1].Much of our knowledge about the specialized functions of DC subpopulations stems from studies using murine BM-derived DC or primary DC [2].A lot of our knowledge about human dendritic cells (DC) is derived from studies using monocyte-derived DC (moDC) as these cells are easily generated in high cell numbers [3,4].However, transcriptomic as well as functional studies show strong differences between moDC and primary DC [5][6][7][8].In order to analyze the function of human primary DC, isolation of steady-state DC subpopulations is necessary.As blood is relatively easily accessible from human donors, we here provide a protocol to isolate peripheral blood mononuclear cells containing all known DC subpopulations [9].As for functional assays, high numbers of DC are necessary, we here use leukocyte reduction system (LRS) cones, which can be obtained from healthy donors undergoing thrombocytapheresis.Thereby, isolation of 1×10 9 cells is routinely possible enabling cell sorting (see Section 3 Cell sorting of primary human DC) and various functional assays with primary human cDC1, cDC2 and pDC [10][11][12].

FCS
Quickly thaw FCS at 37°C in a water bath.Once completely thawed, incubate for 15 min at 42°C in the water bath to destroy complement activity (inactivation for 30 min at 56°C is also possible but will result in lower activity of the growth factors contained in FCS).Directly filter the warm FCS through a sterile 0.22 μm membrane (Corning #431118) into a sterile storage bottle (Corning #430518) and aliquot into 50 ml portions.Use aseptic techniques during the whole procedure.Aliquoted FCS should be stored at −20°C.Avoid freeze-thaw cycles.

Trypan blue
Create a 0.9% (w/v) NaCl solution by dissolving NaCl in double-distilled H 2 O. Dissolve 0.36% (w/v) Trypan blue powder in 0.9% NaCl solution.Sterile filter the solution via a 0.22 μM membrane and store at room temperature.

Equipment Company Purpose
Centrifuge "Allegra X-15R" Beckman-Coulter Centrifugation of 50 ml tubes, 15 ml tubes and V-bottom plates Incubator "HERAEUS BBD6220" Thermo Scientific Cabinet style incubator with 5% CO 2 and 96% relative humidity for the lymphoid tissue digestion Neubauer chamber 0.100 mm; 0.0025 mm 2  Superior Marienfeld Cell counting Sterile bench "Mars Safety Class 2" Scanlaf Performance of all aseptic procedures 50 ml tubes (#352070) Falcon Centrifugation of cell suspensions Serological pipettes (#606180) Greiner bio-one Pipetting . Isolation of peripheral blood mononuclear cells (PBMCs) from leukocyte reduction system cones.(A) Picture of a leukocyte reduction system (LRS) cone of a healthy blood donor undergoing thrombocytapheresis. White arrows mark the pinched-off flexible tubes, which have to be opened using a scissor to transfer the blood product into a 50 ml tube.(B) The diluted blood product was carefully overlaid onto Lymphocyte Separation medium (clear phase).Phases should be completely separated without mixing.(C) After centrifugation of the tube prepared as shown in (B), erythrocytes and granulocytes build a pellet at the bottom of the tube, while mononuclear cells, such as DC, lymphocytes, and monocytes, are contained in the interphase (marked with white arrows) between the Lymphocyte Separation medium and the diluted blood product.The interphase is collected in order to isolate PBMCS.

Freezing media
Add 10% of DMSO (v/v) to FCS.

Isolation of peripheral blood mononuclear cells (PBMCs)
from leukocyte reduction system (LRS) cones.
1. Disinfect the LRS cone (Figure 1A) completely with 70% Ethanol before transferring it into the sterile hood.2. Transfer the blood product inside the cone into a 50 ml tube by opening the pinched-off flexible tubes (marked in Figure 1A with white arrows) using a sterile scissor (also buffy coats or full blood products can be processed with this protocol; please refer to section 1.1.6). 3. Dilute the blood product with RPMI-1640 to 40 ml.Prepare two 50 ml tubes and fill them with 14 ml Lymphocyte Separation medium (density of 1.077 g/ml).Carefully overlay 20 ml of the diluted blood product onto the 14 ml of lymphocyte separation media.Pipet slowly and avoid any mixing of the two phases (Figure 1B). 4. Centrifuge for 25 min with 520 × g at RT without brakes (deceleration set to 0).Transfer the interphase (marked with white arrows in Figure 1C) containing the PBMCs into a new 50 ml tube using a pipette.Then, fill up to 50 ml and centrifuge for 5 min with 520 × g at RT. 5. Discard the supernatant and resuspend the cell pellet in 10 ml RPMI-1640.Fill up to 50 ml and centrifuge again for 5 min with 520 × g at RT.

Blood product Live cells per spleen
Leukocyte reduction system cone (after thrombocytapheresis) 8 × 10 8 -12 × 10 8 6. Discard the supernatant and resuspend the cell pellet in 10 ml RPMI-1640.Harvest 10 μl of the cell suspension and dilute it 1:50 in PBS.Fill up the cell suspension in the 50 ml tube to 50 ml with RPMI-1640 and centrifuge again for 5 min with 520 × g at RT. 7. In the meantime, perform cell counting.Therefore, dilute 10 μl of the 1:50 diluted cell suspension with Trypan blue (1:10) and determine the cell number using a Neubauer chamber (see table 3 for expected cell yield).8.After centrifugation, discard the supernatant and resuspend the cells in a buffer of your choice with a final concentration of 5×10 7 -1×10 8 cells/ml.Typically, you should retrieve about 1×10 9 PBMCs from an LRS cone of a healthy donor undergoing thrombocytapheresis.You can now proceed with flow cytometric analysis of blood DC (see section 2.1) or the enrichment of primary blood DC for cell sorting for functional assays (see section 3 Cell sorting of primary human DC).

Data analysis
Examples for flow cytometry data analysis of DC subsets in blood, spleen, thymus, or tonsils using the described single cell preparation are covered in detail in the section 2 Flow cytometric analysis of the human lymphohematopoietic DC compartment.

Pitfalls
Problem: The cell yield is too low

Potential solution:
During preparation of the PBMCs, make sure that you carefully overlay the lymphocyte separation medium with the diluted blood product and collect the whole interphase after the density gradient centrifugation.In addition, verify that the deceleration of the centrifugation was set to zero.

Potential solution:
In case the interphase is blurry, often the overlaying of the lymphocyte separation medium with the diluted blood product was not performed properly.Make sure that the two phases are clearly and sharply separated.If the centrifugation still leads to a blurry interphase, the setup of the centrifugation might be adapted.Dependent on the protocol and the manufacturer of the separation medium, different centrifugal forces are recommended for the centrifugation.Thus, it can be tested whether centrifugation with lower (e.g., 400 × g) or higher (e.g., 800 × g to 1100 × g) centrifugal forces will lead to a clearer interphase.Further, in addition to deceleration also acceleration of the centrifuge can be set to lower values (e.g.test acceleration values of 0 to 6 instead of the maximum value of 10).

Potential solution:
In this protocol, we use 520 × g for the density gradient centrifugation.In our hands, it leads to pure PBMCs without major leftovers of granulocytes.In case you observe contamination with granulocytes, make sure that you only isolated the interphase after the density gradient centrifugation and did not disturb the pellet of red blood cells and granulocytes.Further, avoid collecting too much of the lymphocyte separation medium in order to obtain pure PBMCs.If you still have a contamination with granulocytes, higher centrifugal forces might be tested.Other protocols suggest centrifugal forces up to 1100 × g.However, carefully check viability of the cells, if you test higher centrifugal forces for the density gradient centrifugation.

Top tricks
Using this protocol, also other blood products such as buffy coats or full blood can be processed.Therefore, the dilution of the blood product with RPMI prior to density gradient centrifugation should be adapted (see step 3 in 1.1.3.2).For LRS cones, which are highly enriched for mononuclear cells, the blood product is diluted 1:4 (8 ml of blood product with approximately 32 ml of RPMI).For fresh full blood, dilute it 1:1 with RPMI (20 ml of blood with 20 ml of RPMI).For buffy coats, dilute the blood product 2:3 with RPMI (e.g. 60 ml of blood product with 90 ml of RPMI).After dilution, proceed as described above: overlay 20 ml of blood product onto 14 ml of lymphocyte separation media.
In mice, the spleen is the major source for the isolation and analysis of murine primary DC.Also in humans, functional and transcriptomic analysis on human splenic DC was performed [12,19,[25][26][27].Thereby, it became evident that the microenvironment has an influence on the phenotype of the different DC subpopulations [12].However, also the protocol of organ preparation has an influence on the yield and condition of the cells.For instance, crawl outs of DC from the skin will not lead to digestion of surface markers needed for analysis of DC subpopulations as no enzymes are used, but will have an impact on the activation status of the DC as only semi-mature or activated DC will leave the skin [16, 21-24, 28, 29].On the other hand, enzymatic digestion can lead to digestion of surface markers necessary for subsequent analysis and cell sorting of DC subpopulations [30,31].Therefore, we here provide an optimized protocol for isolation of mononuclear cells from human spleen allowing for the analysis of otherwise digestion-sensitive markers.

Reagents.
A complete list of reagents is provided in Table 4. Human splenic tissue was received under local ethical committee approval (Ethikkommission der Friedrich-Alexander-Universität Erlangen-Nürnberg, #3761).

FCS
Quickly thaw FCS at 37°C in a water bath.Once completely thawed, incubate for 15 min at 42°C in the water bath to destroy complement activity (inactivation for 30 min at 56°C is also possible but will result in a lower activity of the growth factors contained in FCS).Directly filter the warm FCS through a sterile 0.22 μm membrane (Corning #431118) into a sterile storage bottle (Corning #430518) and aliquot into 50 ml portions.Use aseptic techniques during the whole procedure.Aliquoted FCS should be stored at -20°C.Avoid freeze-thaw cycles.

Trypan blue
Create a 0.9% (w/v) NaCl solution by dissolving NaCl in double-distilled H 2 O. Dissolve 0.36% (w/v) Trypan blue powder in 0.9% NaCl solution.Sterile filter the solution via a 0.22 μM membrane and store at room temperature.

Freezing media
Add 10% of DMSO (v/v) to FCS.

Digestion of human spleen.
1. Store the human spleen tissue in sterile RPMI-1640 on ice.
Proceed as soon as possible with the preparation of the tissue to avoid any activation of the DC. 2. Transfer the tissue into a sterile cell culture plate using sterile forceps (Figure 2A).2a.In case immunofluorescence analyses should be performed, remove small cubes (2×2×6 mm) of the tissue using sterile forceps and scalpel covering the whole architecture of the tissue (including the capsule) (for more detailed instructions, please refer to [16,17].2b.In case immunofluorescence analyses should be performed, fill cryomolds with O.C.T. tissue tek and avoid any air bubbles (Figure 2B).Transfer the splenic cubes into the cryomold and store them at -80°C until cutting with a cryotome (for more detailed instructions, please refer to [32,33]).3.For the preparation of single cell suspensions, proceed with mechanical disruption of the remaining tissue using forceps and scalpel.The tissue should be cut into pieces as small as possible as it will facilitate the enzymatic digestion ( Use the sterile part of the plug of a 2 ml syringe to facilitate the passage over the filter by stirring the cell suspension in the filter.10.Wash the filter with 5 ml of washing buffer.Add the second C-tube to the same filter.Again, use the plug of the syringe to facilitate the passage over the filter.11.Wash the filter with further 5 ml of washing buffer.12. Remove the filter from the 50 ml tube and fill up to 50 ml with washing buffer.

Tissue Live cells per g of spleen tissue
Spleen 1.5×10 8 -2.5×10 8   13.Centrifuge for 5 min at 520 × g at 4°C.Carefully pour off the supernatant and resuspend the cell pellet in 10 ml of washing buffer.14.Prepare for each 50 ml tube a new 50 ml tube and insert a 70 μm filter.Wet the filter with 5 ml washing buffer and pipet or pour the resuspended cell suspension onto the filter.A new plug of a syringe can be used in case the filter is clogging.15.Wash the filter with 5 ml of washing buffer.Remove the filter and fill up to 50 ml with washing buffer.16.Centrifuge for 5 min at 520 × g at 4°C.Pour off the supernatant and resuspend the cell pellet in 10 ml of RPMI-1640.
Fill up to 40 ml with RPMI-1640.17. Prepare for each 50 ml tube two new 50 ml tubes.Pipet 14 ml of lymphocyte separation media (ρ = 1.077 g/ml, RT) into each of the new 50 ml Tubes.18. Carefully overlay 20 ml of the cell suspension onto 14 ml of lymphocyte separation media.Avoid any mixing of the liquids as it will reduce the final cell yield.19.To perform a density gradient centrifugation, centrifuge for 30 min at 520 × g at RT and set deceleration to 0. 20.Harvest the interphase between the lymphocyte separation media and the RPMI-1640 and transfer into a new 50 ml tube.Pool up to four interphases into one 50 ml tube.Fill up to 50 ml with RPMI-1640 and centrifuge for 5 min at 520 × g at 4°C. 21.Pour off the supernatant, resuspend in 10 ml of RPMI-1640, and pool all 50 ml tubes into one.Fill up to 50 ml and centrifuge again at 520 × g at 4°C for 5 min.22.To determine the cell number, resuspend the cell pellet in 10 ml of RPMI-1640.Pipet 10 μl of the cell suspension into a 1.5 ml microtube and dilute it 1:200 to 1:1,000 depending on the size of the pellet.23.Dilute 10 μl of the diluted cell suspension 1:10 with Trypan blue and determine the cell number using a Neubauer chamber.Typically, 1.7 × 10 8 cells/g of tissue (±9.8 × 10 7 ) are retrieved (see Table 6).24.Cells can be either used directly for flow cytometric analysis (see Section 2 Flow cytometric analysis of the human DC compartment in lymphohematopoietic tissues) or cell sorting of DC subpopulations after further enrichment (see Section 3 Cell sorting of primary human DC).
Unused cells can be stored in liquid nitrogen.In case tanks with liquid nitrogen for storage are not available, storage at −80°C is also possible.Therefore, resuspend the cells in freezing media to reach a final concentration of 5 × 10 7 cells/ml and transfer the cell suspension into 1.8 ml or 4 ml CryoTubes.Transfer the CryoTubes into isopropanol-containing Mr. Frosty TM Freezing Container and freeze/store at −80°C.

Data analysis
Examples for flow cytometry data analysis of DC subsets in human spleen using the described single cell preparation are covered in detail in Section 2.2 Flow cytometric analysis of human splenic DC.

Potential solution:
In case a Gentle MACS Dissociator is not available, digestion can be performed in 50 ml tubes instead of C-tubes.Therefore, tissue should be cut into pieces as small as possible.Further, repetitively invert the 50 ml tubes during the incubation step at 37°C.During the first filter step (see step 9), the tissue has to be ground over the filter using the sterile plug of a syringe.As the filters will clog faster without the dissociation using the Gentle MACS Dissociator, for each 50 ml tube filled with 2 g of tissue one 100 μm filter should be used instead of pooling two tubes over one filter.

Potential solutions:
Potentially, the tissue was not completely digested.Therefore, it should be tested whether the concentrations of the enzymes have to be increased as the efficiency of the enzymes might vary from Lot to Lot.Additionally, an increase of the incubation time at 37°C might be tested.During filtering, incompletely digested tissue might be ground over the filter to release further cells from the tissue.In order to ease the digestion, the tissue should be mechanically reduced to the smallest pieces possible using scalpels.The Gentle MACS program "m_liver_02.1"should be used in addition before the incubation at 37°C.

Potential solution:
Either the tissue was not completely digested or too much cell death occurred during incubation at 37°C.Please refer to potential solution of low cell yield (see above) for advice how to improve the digestion of the tissue.In addition, the capsule of the spleen is hardly digested with the enzymes used in this protocol.Therefore, it might be beneficial to remove the capsule before transferring the tissue into the C-tubes to reduce the amount of tissue, which can potentially clog the cell strainer.However, be careful to not lose too much tissue.Further, to reduce the amount of tissue per cell strainer, for each C-tube a separate 100 μm cell strainer can be used.

Potential solution:
Make sure that the lymphocyte separation medium was carefully overlaid with the cell suspension.In case the two liquids are mixing at the interphase even to a low amount, it will lead to a blurry interphase.In addition, verify that the deceleration of the centrifugation was set to zero.If the liquids were separated by a clear edge and the deceleration was set to zero, but still only a few cells were located in the interphase, the digestion might be not efficient leading to fewer cells in the cell suspension.Refer to the suggestion on how to improve the cell yield above.

Potential solution:
In case one received appropriate amounts of total cells from the digestion of the spleen, but it was not possible to detect a clear DC compartment by flow cytometric analysis of the single cell suspension (see Section 2 Flow cytometric analysis of the human DC compartment in lymphohematopoietic tissues), the condition of the donor might be a problem.In our experience, spleen sample from patients suffering from splenomegaly often contain huge numbers of B cells further diluting the DC compartment.Thereby, hardly any DC are detectable, if performing standard flow cytometry.
Further, it is very important to process the tissue as soon as possible after surgical removal.The tissue should be kept in PBS or RPMI-1640 at 4°C until preparation of the single cell suspension to avoid cell death as well as activation of the cells.

Top tricks
This protocol can be easily used for other lymphoid tissues, such as thymus, lymph nodes, or tonsils, by adapting the concentration of the enzymes dependent on the tissue (see Table 7) [10][11][12].In the thymus, it is important to increase the concentration of DNase I as during processing of the tissue immature thymocytes undergo cell death.Thereby, more potentially immune-stimulatory DNA is released into suspension, which has to be digested.
Depending on the source of lymph nodes, the amount of tissue is very low (< 1 g).Here, it might be better to digest the tissue directly in 50 ml FACS tubes instead of using C-tubes and grind the tissue over the cell strainer as explained in section 1.2.5 Pitfalls (see above).

Introduction
DC are not only involved in peripheral T cell tolerance [34,35], but also have a role in central tolerance [36][37][38].Therefore, analysis of thymic DC subpopulations is important in order to unravel how they contribute to negative and positive selection in the thymus.Several reports showed that conventional as well as plasmacytoid DC are present in human thymus [12,[39][40][41][42].As DC are a rare cell population in the thymus, efficient isolation of the cells is necessary.We here provide a protocol leading to high numbers of cells isolated from human thymus without the induction of maturation.Further, digestion of the tissue preserves expression of otherwise digestion-sensitive markers such as CLEC9A.Thereby, flow cytometric analysis and cell sorting of DC subpopulations is possible (see section 2.3 Flow cytometric analysis of human thymic DC).

FCS
Quickly thaw FCS at 37°C in a water bath.Once completely thawed, incubate for 15 min at 42°C in the water bath to destroy complement activity (inactivation for 30 min at 56°C is also possible but will result in lower activity of the growth factors contained in FCS).Directly filter the warm FCS through a sterile

Freezing media
Add 10% of DMSO (v/v) to FCS.

Digestion of human thymus.
1. Store the human thymus tissue in sterile RPMI-1640 on ice.
Proceed as soon as possible with the preparation of the tissue to avoid any activation of the DC. 2. Transfer the tissue into a sterile cell culture plate using sterile forceps (Figure 3A).2a.In case immunofluorescence analyses should be performed, remove small cubes (2×2×6 mm) of the tissue using sterile forceps and scalpel covering the whole architecture of the tissue (including the capsule) (for more detailed instructions, please refer to [32,33]).2b.In case immunofluorescence analyses should be performed, fill cryomolds with O.C.T. tissue tek and avoid any air bubbles (Figure 2B).Transfer the thymic cubes into the cryomold and store them at -80°C until cutting with a cryotome (for more detailed instructions, please refer to [32,33]).3.For preparation of single cell suspensions, proceed with mechanical disruption of the remaining tissue using forceps and scalpel.The tissue should be cut into pieces as small as possible as it will facilitate the enzymatic digestion (Figure 3B). 4. For digestion of the tissue, use C-tubes and a Gentle MACS Dissociator.Use one C-tube per ∼2 g thymic tissue.Thus, if preparing 10 g of total tissue, use five C-tubes.More than 2 g of tissue can lead to overloading of the C-tubes.In case a Gentle MACS Dissociator is not available in the lab, please refer to the Pitfalls sections.5. Add 5 ml of RPMI-1640 to each C-tube as well as 500 μl Collagenase IV and 300 μl DNase I (Figure 3C).Tightly close the C-tubes and proceed with mechanical disruption using the Gentle MACS Dissociator.6. Run the program 'm_liver_01.1'and incubate the C-tubes at 37°C for 45 min.After 30 min, run the program 'm_liver_02.1' to facilitate the digestion of the tissue.7.After the end of the incubation time, run the program 'm_liver_02.1'again, add 5 ml of washing buffer and put the C-tubes on ice to stop the digestion of the tissue.8. Prepare one 50 ml tube for each two C-tubes.Insert a 100 μm filter into each 50 ml tube and wet it with 5 ml washing buffer.9. Pipet or pour the cell suspension of one C-tube onto the filter.
Use the sterile part of the plug of a 2 ml syringe to facilitate the passage over the filter by stirring the cell suspension in the filter.10.Wash the filter with 5 ml of washing buffer.Add the second C-tube to the same filter.Again, use the plug of the syringe to facilitate the passage over the filter.11.Wash the filter with further 5 ml of washing buffer.12. Remove the filter from the 50 ml tube and fill up to 50 ml with washing buffer.13.Centrifuge for 5 min at 520 × g at 4°C.Carefully pour off the supernatant and resuspend the cell pellet in 10 ml of washing buffer.14.Prepare for each 50 ml tube a new 50 ml tube and insert a 70 μm filter.Wet the filter with 5 ml washing buffer and pipet or pour the resuspended cell suspension onto the filter.A new plug of a syringe can be used in case the filter is clogging.15.Wash the filter with 5 ml of washing buffer.Remove the filter and fill up to 50 ml.16.Centrifuge for 5 min at 520 × g at 4°C.Pour off the supernatant and resuspend the cell pellet in 10 ml of RPMI-1640.Fill up to 40 ml.17.Prepare for each 50 ml tube filled with single cell suspension two new 50 ml tubes.Pipet 14 ml of lymphocyte separation media (ρ = 1.077 g/ml, RT) into each of the new 50 ml Tubes.18. Carefully overlay 20 ml of the cell suspension onto 14 ml of lymphocyte separation media.Avoid any mixing of the liquids as it will reduce the final cell yield (Figure 3D).

Tissue Live cells per g of human thymus tissue
Thymus 3×10 8 -4×10 8   19.To perform a density gradient centrifugation, centrifuge for 30 min at 520 × g at RT and set deceleration to 0. 20.Harvest the interphase between the lymphocyte separation media and the RPMI-1640 and transfer into a new 50 ml tube (marked with a black rectangle in Figure 3E).Pool up to four interphases into one 50 ml tube.Fill up to 50 ml with RPMI-1640 and centrifuge for 5 min at 520 × g at 4°C. 21.Pour off the supernatant, resuspend in 10 ml of RPMI-1640, and pool all 50 ml tubes into one.Fill up to 50 ml and centrifuge again at 520 × g at 4°C for 5 min.22.To determine the cell number, resuspend the cell pellet in 10 ml of RPMI-1640.Pipet 10 μl of the cell suspension into a 1.5 ml microtube and dilute it 1:200 to 1:1000 dependent on the size of the pellet.23.Dilute 10 μl of the diluted cell suspension 1:10 with Trypan blue and determine the cell number using a Neubauer chamber.Typically, 3.6×10 8 cells/g of tissue (±2.7 × 10 8 ) are retrieved (see Table 10).24.Cells can be either used directly for flow cytometric analysis (see Section 2 Flow cytometric analysis of the human DC compartment in lymphohematopoietic tissues) or cell sorting of DC subpopulations after further enrichment (see Section 3 Cell sorting of primary human DC).25.Unused cells can be stored in liquid nitrogen.In case, tanks with liquid nitrogen for storage are not available, storage at −80°C is also possible.Therefore, resuspend the cells in freezing media to reach a final concentration of 5 × 10 7 cells/ml and transfer the cell suspension into 1.8 ml or 4 ml CryoTubes.Transfer the CryoTubes into isopropanolcontaining Mr. Frosty TM Freezing Container and freeze/store at −80°C.

Data analysis
Examples of flow cytometry data analysis of DC subsets in human thymus using the described single cell preparation are covered in detail in Section 2.3 Flow cytometric analysis of human thymic DC.

Potential solution:
In case a Gentle MACS Dissociator is not available, digestion can be performed in 50 ml tubes instead of C-tubes.Therefore, tissue should be cut into pieces as small as possible.Further, repetitively invert the 50 ml tubes during the incubation step at 37°C.
During the first filter step (see step 9) the tissue has to be ground over the filter using the sterile plug of a syringe.As the filters will clog faster without the dissociation using the Gentle MACS Dissociator, for each 50 ml tube filled with 2 g of tissue one 100 μm filter should be used instead of pooling two tubes over one filter.

Potential solution:
Potentially, the tissue was not completely digested.Therefore, it should be tested whether the concentrations of the enzymes have to be increased as the efficiency of the enzymes might vary from Lot to Lot.Additionally, an increase of the incubation time at 37°C might be tested.During filtering, incompletely digested tissue might be ground over the filter to release further cells from the tissue.In order to ease the digestion, the tissue should be mechanically reduced to the smallest pieces possible using scalpels.The Gentle MACS program 'm_liver_02.1'should be used in addition before the incubation at 37°C.Moreover, the thymus involutes with increasing age of the donor leading to less cells and more fat tissue.In this case, the fat from the thymic tissue is released during the processing and swimming in and on top of the cell suspension.This disturbs centrifugation/washing steps leading to cell loss.Therefore, the fat swimming on top of the cell suspension should be removed after the first washing step after digestion/filtering over 100 μm strainer (step 13) using a pipette.Further, instead of pouring off the supernatant, the liquid should be removed using a pipette to avoid losing cells.

Potential solution:
Either the tissue was not completely digested or too much cell death occurred during incubation at 37°C.Please refer to the potential solution of low cell yield (see above) for advice on how to improve the digestion of the tissue.

Potential solution:
Make sure that the lymphocyte separation medium was carefully overlaid with the cell suspension.In case the two liquids are mixing at the interphase even to a low amount, it will lead to a blurry interphase.In addition, verify that the deceleration of the centrifugation was set to zero.If the liquids were separated by a clear edge and the deceleration was set to zero, but still only few cells were located in the interphase, the digestion might be not efficient leading to fewer cells in the cell suspension.Refer to the suggestion on how to improve the cell yield above.

Potential solution:
In case one received appropriate amounts of total cells from the digestion of the thymus, but it was not possible to detect a clear DC compartment by flow cytometric analysis of the single cell suspension (see section 2 Flow cytometric analysis of the human DC compartment in lymphohematopoietic tissues), the digestion of the tissue might have to be improved.While thymocytes are easily be removed from the tissue without enzymatic digestion, for isolation of DC enzymatic digestion is necessary.If hardly any DC are detectable, please refer to the solutions for low cell yield.In addition, the age of the thymus donor can have an influence on the amount of DC as with involution of the thymus the number of DC decreases and is shifted more to plasmacytoid DC.
In order to increase thymic DC for flow cytometric analysis, DC might be enriched by depletion of thymocytes using biotincoupled anti-human CD3 and anti-human CD8α antibodies (see section 1.3.6Top tricks).
Further, it is very important to process the tissue as soon as possible after surgical removal.The tissue should be kept in PBS or RPMI-1640 at 4°C until preparation of single cell suspension to avoid cell death as well as activation of the cells.

Top tricks
Dendritic cells are a rare cell population in thymus tissue.In order to perform flow cytometric analysis or even functional assays such as internalization assays [10,11], it might be necessary to enrich the DC by negative depletion of thymocytes.Therefore, biotin-coupled anti-human CD3 and anti-human CD8α antibodies might be used followed by incubation with streptavidin-coated magnetic nanobeads (BioLegend).During an incubation inside a magnet, thymocytes labelled with streptavidin-coated magnetic nanobeads are depleted and DC negatively enriched.This enables functional assays as well as is necessary for efficient cell sorting of human DC (see section 3 Cell sorting of primary human DC).
Moreover, this protocol can be easily used for other lymphoid tissues, such as spleen, lymph nodes or tonsils, by adapting the concentration of the enzymes dependent on the tissue (see Table 11) [10][11][12].
While lymph nodes are rather small and rarely removed except for tumor-draining lymph nodes during excision of tumor, tonsils represent a more abundant source as tonsillectomy is a standard procedure for patients suffering from tonsillitis.Human tonsils were shown to contain conventional as well as plasmacytoid DC [12,43].As the structure and function of tonsils are comparable to lymph nodes, isolated tonsil DC can be used for the analysis of DC functions [44,45].

Reagents.
A complete list of reagents is provided in Table 12.Human tonsillar tissue was received under local ethical committee approval (Ethikkommission der Friedrich-Alexander-Universität Erlangen-Nürnberg, #3761).

Equipment.
Necessary equipment is listed in Table 13.

FCS
Quickly thaw FCS at 37°C in a water bath.Once completely thawed, incubate for 15 min at 42°C in the water bath to destroy

Trypan blue
Create a 0.9% (w/v) NaCl solution by dissolving NaCl in double-distilled H 2 O. Dissolve 0.36% (w/v) Trypan blue powder  Freezing media Add 10% of DMSO (v/v) to FCS.

Digestion of human tonsils.
1. Store the human tonsillar tissue in sterile RPMI-1640 on ice.Proceed as soon as possible with the preparation of the tissue to avoid any activation of the DC. 2. Transfer the tissue into a sterile cell culture plate using sterile forceps (Figure 4).2a.In case immunofluorescence analyses should be performed, remove small cubes (2×2×6 mm) of the tissue using sterile forceps and scalpel covering the whole architecture of the tissue (including the capsule) (for more detailed instructions, please refer to [32,33]).2b.In case immunofluorescence analyses should be performed, fill cryomolds with O.C.T. tissue tek and avoid any air bubbles.Transfer the tonsillar cubes into the cryomold and store them at -80°C until cutting with a cryotome (for more detailed instructions, please refer to [32,33]).
3. For preparation of single cell suspensions, proceed with mechanical disruption of the remaining tissue using forceps and scalpel.The tissue should be cut into pieces as small as possible as it will facilitate the enzymatic digestion.During tonsillectomy, tonsillar tissue is cauterized (marked with white arrows in Figure 4).Try to remove cauterized tissue as it will not be digested and might clog the filter.4. For digestion of the tissue, use C-tubes and a Gentle MACS Dissociator.Use one C-tube per ∼2 g tonsillar tissue.Thus, if preparing 4 g of total tissue, use two C-tubes.More than 2 g of tissue can lead to overloading of the C-tubes.In case a Gentle MACS Dissociator is not available in the lab, please refer to section 1.4.5 Pitfalls.5. Add 5 ml of RPMI-1640 to each C-tube as well as 500 μl Collagenase IV and 100 μl DNase I. Tightly close the C-tubes and proceed with mechanical disruption using the Gentle MACS Dissociator.6. Run the program "m_liver_01.1"and incubate the C-tubes at 37°C for 45 min.After 30 min, run the program "m_liver_02.1" to facilitate the digestion of the tissue.7.After the end of the incubation time, run the program "m_liver_02.1"again, add 5 ml of washing buffer, and put the C-tubes on ice to stop the digestion of the tissue.8. Prepare one 50 ml tube for each two C-tubes.Insert a 100 μm filter into each 50 ml tube and wet it with 5 ml washing buffer.9. Pipet or pour the cell suspension of one C-tube onto the filter.
Use the sterile part of the plug of a 2 ml syringe to facilitate the passage over the filter by stirring the cell suspension in the filter.10.Wash the filter with 5 ml of washing buffer.Add the second C-tube to the same filter.Again, use the plug of the syringe to facilitate the passage over the filter.11.Wash the filter with further 5 ml of washing buffer.12. Remove the filter from the 50 ml tube and fill up to 50 ml with washing buffer.13.Centrifuge for 5 min at 520 × g at 4°C.Carefully pour off the supernatant and resuspend the cell pellet in 10 ml of washing buffer.14.Prepare for each 50 ml tube a new 50 ml tube and insert a 70 μm filter.Wet the filter with 5 ml washing buffer and pipet or pour the resuspended cell suspension onto the filter.A new plug of a syringe can be used in case the filter is clogging.15.Wash the filter with 5 ml of washing buffer.Remove the filter and fill up to 50 ml with washing buffer.16.Centrifuge for 5 min at 520 × g at 4°C.Pour off the supernatant and resuspend the cell pellet in 10 ml of RPMI-1640.
Fill up to 40 ml with RPMI-1640.17. Prepare for each 50 ml tube filled with single cell suspension two new 50 ml tubes.Pipet 14 ml of lymphocyte separation media (ρ = 1.077 g/ml, RT) into each of the new 50 ml tubes.18. Carefully overlay 20 ml of the cell suspension onto 14 ml of lymphocyte separation media.Avoid any mixing of the liquids as it will reduce the final cell yield.19.To perform a density gradient centrifugation, centrifuge for 30 min at 520 × g at RT and set deceleration to 0. 20.Harvest the interphase between the lymphocyte separation media and the RPMI-1640 and transfer into a new 50 ml tube (marked with a black rectangle in Figure 3E).Pool up to four interphases into one 50 ml tube.Fill up to 50 ml with RPMI-1640 and centrifuge for 5 min at 520 × g at 4°C. 21.Pour off the supernatant, resuspend in 10 ml of RPMI-1640, and pool all 50 ml tubes into one.Fill up to 50 ml and centrifuge again at 520 × g at 4°C for 5 min.22.To determine the cell number, resuspend the cell pellet in 10 ml of RPMI-1640.Pipet 10 μl of the cell suspension into a 1.5 ml microtube and dilute it 1:200 to 1:1000 dependent on the size of the pellet.

Tissue Live cells per g of tonsillar tissue
Tonsil 8×10 7 -12×10 7   23.Dilute 10 μl of the diluted cell suspension 1:10 with Trypan blue and determine the cell number using a Neubauer chamber.Typically, 1.1×10 8 cells/g of tissue (±6.7 × 10 7 ) are retrieved (see Table 14).24.Cells can be either used directly for flow cytometric analysis (see Section 2 Flow cytometric analysis of the human DC compartment in lymphohematopoietic tissues) or cell sorting of DC subpopulations after further enrichment (see Section 3 Cell sorting of primary human DC).25.Unused cells can be stored in liquid nitrogen.In case tanks with liquid nitrogen for storage are not available, storage at -80°C is also possible.Therefore, resuspend the cells in freezing media to reach a final concentration of 5×10 7 cells/ml and transfer the cell suspension into 1.8 ml or 4 ml Cry-oTubes.Transfer the CryoTubes into isopropanol-containing Mr. Frosty TM Freezing Container and freeze/store at −80°C.

Data analysis
Examples of flow cytometry data analysis of DC subsets in human tonsils using the described single cell preparation are covered in detail in section 2.4 Flow cytometric analysis of human tonsillar DC.

Pitfalls
Problem: Gentle MACS Dissociator is not available

Potential solution:
In case a Gentle MACS Dissociator is not available, digestion can be performed in 50 ml tubes instead of C-tubes.Therefore, tissue should be cut into pieces as small as possible.Further, repetitively invert the 50 ml tubes during the incubation step at 37°C.During the first filter step (see step 9) the tissue has to be ground over the filter using the sterile plug of a syringe.As the filters will clog faster without the dissociation using the Gentle MACS Dissociator, for each 50 ml tube filled with 2 g of tissue one 100 μm filter should be used instead of pooling two tubes over one filter.

Potential solution:
Potentially, the tissue was not completely digested.Therefore, it should be tested whether the concentrations of the enzymes have to be increased as the efficiency of the enzymes might vary from Lot to Lot.Additionally, an increase of the incubation time at 37°C might be tested.During filtering, incompletely digested tis-sue might be ground over the filter to release further cells from the tissue.In order to ease the digestion, the tissue should be mechanically reduced to the smallest pieces possible using scalpels.The Gentle MACS program 'm_liver_02.1'should be used in addition before the incubation at 37°C.

Potential solution:
Either the tissue was not completely digested or too much cell death occurred during incubation at 37°C.Please refer to potential solution of low cell yield (see above) for advice how to improve the digestion of the tissue.Further, cauterized tissue should be removed before the digestion as it can lead to clogging of the cell strainer.

Potential solution:
Make sure that the lymphocyte separation medium was carefully overlaid with the cell suspension.In case the two liquids are mixing at the interphase even to a low amount, it will lead to a blurry interphase.In addition, verify that the deceleration of the centrifugation was set to zero.If the liquids were separated by a clear edge and the deceleration was set to zero, but still only a few cells were located in the interphase, the digestion might be not efficient leading to fewer cells in the cell suspension.Refer to the suggestion on how to improve the cell yield above.

Potential solution:
In case one received appropriate amounts of total cells from the digestion of the tonsil, but it was not possible to detect a clear DC compartment by flow cytometric analysis of the single cell suspension (see Section 2 Flow cytometric analysis of the human DC compartment in lymphohematopoietic tissues), the condition of the organ donor was a problem.As tonsils are often removed due to tonsillitis, highly inflamed tissue might have some changes in immune cell compartments.
Further, it is very important to process the tissue as soon as possible after surgical removal.The tissue should be kept in PBS or RPMI-1640 at 4°C until preparation of single cell suspension to avoid cell death as well as activation of the cells.

Top tricks
If single cell suspensions are generated to allow for cellular enrichment procedures, replace the classical FACS buffer with the buffer system suggested by the manufacturer of cellular enrichment kits during cellular enrichment procedures.

Introduction
Dendritic cells (DC) are important regulators of the immune system [2].After the first identification of human DC in the blood, the DC compartment was constantly characterized in more detail.While in the beginning DC were broadly identified as HLA-DR + and CD11c + cells, nowadays several subpopulations with specialized functions are distinguished [9].In general, DC lack the expression of so-called lineage markers, which are expressed on T cells (CD3), B cells (CD19/CD20), NK cells (CD56/CD335), and monocytes (CD14/CD16) but express HLA-DR, one of the human MHC class II molecules [9,12,46].Broadly, DC are further classified into conventional (cDC) and plasmacytoid DC (pDC).pDC can be easily identified by the expression of CD123, CD303, or CD304.However, recent reports show that precursors of cDC (precDC) also express CD123 and CD303 but can be distinguished using the markers Axl as well as Siglec-6 [47,48].While pDC lack CD11c expression, cDC express it at varying degrees.However, one has to be careful when using CD11c to identify human DC -as monocytes and macrophages highly express CD11c as well.
In the following, we provide an easy and ready-to-use protocol for analysis of cDC, pDC, and monocyte populations, including recently identified subpopulations of CD1c + DC, isolated from different tissues of the lymphohematopoietic system via a conserved core-panel by flow cytometry.Further, we show an exemplary gating strategy for the identification of DC subpopulations in the blood.

Reagents.
A complete list of reagents is provided in Table 15.

Equipment. Necessary equipment is listed in Table 16
and BD LSR Fortessa configuration in Table 17.

DAPI
Dissolve 4´,6-diamidino-2´-phenylindole dihydrochloride (DAPI) in ultrapure water to create a 1 mg/ml stock solution.Store the solution protected from light at 4°C.Dilute the stock solution 1:10,000 in FACS buffer to create a working solution for DAPI staining of cells directly before acquisition at the flow cytometer.

FACS buffer
Add 2% human serum type AB (v/v) to 500 ml of Phosphate buffered saline solution (PBS).

Antibody staining of single cell suspensions from human blood for flow cytometry.
In Section 1.1 Isolation of mononuclear cells from peripheral blood of this section, we described how to prepare PBMCs from human blood.
1. Following their preparation, 5 × 10 7 cells per donor are transferred per well to a 96-well V-bottom plate to perform antibody staining enabling flow cytometric analysis of the DC compartment.2. Before centrifugation of the samples, prepare the first staining mix. 3. Therefore, use 50 μl of FACS buffer per sample and add the antibodies listed in Table 18 in the indicated dilution.Please note that antibody dilutions are dependent on the flow cytometer and its setup.Suggested antibody dilutions were optimized for a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer see Table 17).If the cell count strongly differs from the expected cellular yields (>factor 1.5), scale the volume of antibody cocktails up and down accordingly to maintain a constant detector antibodyto-target ratio.Do not stain in less than 35 μl. 4. Commercially available Fc block solution is used to block binding to Fc gamma receptors, thereby avoiding unspecific recognition of cell-bound Fc gamma receptors; also other antibodies blocking Fc receptors or sera (mouse or rat, dependent on source of antibodies) might be suitable.However, optimal dilutions have to be determined.5.After preparation of the staining mix, centrifuge the samplecontaining 96-well V-bottom plates at 520 × g at 4°C for 5 min.6. Discard the supernatant, keep the plate up-side down and dip on a paper towel.17), the event rate should not exceed 15,000 events/s.25.Freshly prepared samples can be directly acquired, while thawed samples have to be filtered via 35 μm cell strainers into FACS tubes to avoid clogging of the flow cytometer.26.After performing the cytometer setup, acquire data at an appropriate flow cytometer (Table 17).

Data analysis
Data acquisition was performed at a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer, see Table 17).Subsequently, data were analyzed using FlowJo software.In the following, we provide exemplary gating strategies for PBMCs analyzed with our conserved panel.An overview of markers for DC and monocyte populations is provided in Table 20.Figure 5 shows an exemplary gating strategy for the identification of DC in human PBMCs.

Pitfalls
Problem: The signal resolution of a marker is not high enough to identify positive and negative cells

Potential solutions:
Check, if laser lines and employed filter sets are appropriate for excitation and detection of the employed dyes.Therefore, please  refer to Table 19, where we summarized the excitation and emission maxima of the employed dyes.

Potential solutions:
In case you have to fix the cells prior to analysis at a flow cytometer, it might lead to lower signal of some of the antibodies.As you can either fix the cells before or after staining with the fluorochrome-coupled antibodies, you should test which order gives you the best results.Fixing before staining will potentially mask some of the epitopes for the used antibodies, while fixing after staining can harm some of the fluorochromes.We usually prefer fixing after staining.In this case, you should use fluorochromes such as APC/Fire 750 instead of APC/Cy7 as it is more resistant to fixation.Further, excessive fixation should be avoided.Fixation with Cytofix/Cytoperm (BD Biosciences) for 15 min on ice does not impair gating of DC subpopulations as described in this protocol.However, you should change the dye for live/dead discrimination as described in the Top tricks section below.

Potential solutions:
When establishing a multicolour flow cytometry panel or incorporating new antibodies into an existing panel, isotype controls or fluorescence minus one (FMO) controls have to be performed.In case you observe binding of an antibody to a cell type, which should be negative for this marker, perform isotype or FMO controls by replacing the antibody with a corresponding isotype control or leaving the channel empty, respectively.If you perform isotype controls, adjust the dilution of the isotype control according to the concentration of the antibody.In case you observe positive signal for the antibody in the FMO control, the compensation of the flow cytometer was not performed properly or problems with another fluorochrome exist, which spills over into the channel.This might happen, when tandem dyes, such as PerCP/Cy5.5 or PE/Cy7 disaggregate.In case you observe positive signals in the isotype control, then the antibody is binding unspecifically to the cells.You should either titrate the antibody or test other Fcblocking reagents.

Top tricks
The current staining protocol includes several markers defining one cell population such as CD1c, CD301 (CLEC10A), and FcεR1A, which are all expressed on human cDC2.The same is true for cDC1 with CD141, XCR1, and CD370 (CLEC9A).Therefore, the panel can be reduced to a core panel by removing some of the redundant markers to use the free channels for analysis of receptor expression on DC subpopulations including costimulatory/inhibitory receptors for the activation status of the cells or cytokine expression.Additionally, the lineage cocktail, which is currently split up into two channels, can be combined in one to allow for the DC analysis with a reduced staining panel.If you incorporate new antibodies into the panel, please make sure to perform isotype or FMO control for the new antibodies, to ensure that the antibody binds specifically to the cells.Exemplary gating strategy of the dendritic cell compartment in human blood.After a morphology gate followed by elimination of doublets, dead cells (DAPI + ) as well as T cells (CD3 + and/or CD8 + ), B cells (CD19 + /CD20 + ) and NK cells (CD56 + /CD335 + ) were excluded.To distinguish DC from monocytes, CD88 and CD14 were employed, as CD88 is exclusively expressed on monocytes.CD14 low cells were also included in the gate as CD14 can be expressed on low level on human DC3.Remaining cells were separated into CD1c + CD123 − , CD1c − CD123 + , and CD1c − CD123 − cells.Human cDC1 were identified in the CD1c − CD123 − compartment by their high expression of CD141 as well as CLEC9A.cDC1 further express XCR1 (histogram plot).Human cDC2 are contained in the CD1c + CD123 − compartment and are positive for both, CLEC10A and FcεR1A.cDC2 were further subdivided into CD5 + CD163 − DC2 and CD5 neg-low CD163 + DC3.pDC (CD303 + Axl − ) and pre-cDC (CD303 + Axl + ) can be identified in the CD1c − CD123 + compartment using Axl and CD303.Data acquisition was performed at a BD LSR Fortessa and data was evaluated utilizing FlowJo software.The current use of the live-dead discriminator DAPI is not compatible with fixation and intracellular staining.To allow for parallel fixation and live-dead cell discrimination employ a fixable viability dye.In particular, the Zombie UV TM Fixable Viability Kit from BioLegend (#Cat 423107) might be attractive, since the dye is usually detected in the same channel as DAPI.But be aware that fixable viability dyes have to be stained separately in PBS without proteins such as BSA or sera.Further, fixable viability dyes such as Zombie UV bind to a lower extent also to living cells.There-fore, titrate Zombie UV appropriately to ensure proper exclusion of dead cells.
Dependent on the tissue, the dump channel containing the lineage markers can be shrinked or expanded.For instance, in thymus tissue CD3 is only expressed on low level on thymocytes.Here, the inclusion of CD8α is necessary to clearly distinguish cDC2 and thymocytes, while it can be neglected in most other tissues.Further, in tissues with high amounts of B cells (e.g., spleen and tonsils), either the dilution of the B cell-specific antibodies CD19 and CD20 can be lowered or additional B cell-specific markers such as CD21 can be added to improve the differentiation between cDC2 and B cells.
The suggested panel is suitable to analyze DC across various tissues including lympho-hematopoietic (spleen, lymph nodes, and thymus) and peripheral tissues (lung, liver) without the need for major modifications.

Summary of the phenotype
The overall phenotype of immune cells covered by the markers included in the panel is detailed in Table 20.The frequency of the analyzed DC subpopulations is covered in Table 21.

Introduction
Primary human DC were identified at the beginning of the 2000s in the blood using so-called blood DC antigens [13,14].In mice, the spleen is the major source for the isolation and analysis of murine primary DC.Also in humans, functional and transcrip- tomic analysis on human splenic DC was performed [12,19,[25][26][27].
In general, DC lack the expression of so-called lineage markers, which are expressed on T cells (CD3), B cells (CD19/CD20), NK cells (CD56/CD335), and monocytes (CD14/CD16) but express HLA-DR, one of the human MHC class II molecules [9,12,46].Broadly, DC are further classified into conventional (cDC) and plasmacytoid DC (pDC).pDC can be easily identified by the expression of CD123, CD303 or CD304.However, recent reports show that precursors of cDC (precDC) also express CD123 and CD303 but can be distinguished using the markers Axl as well as Siglec-6 [47,48].While pDC lack CD11c expression, cDC express it at varying degrees.However, one has to be careful when using CD11c to identify human DC -as human monocytes and macrophages highly express CD11c as well.The cDC compartment is composed of distinct subpopulations of DC: cDC1, also known as CD141 + DC, are identified by the high expression of CD141 as well as by the marker CLEC9A, XCR1, and CD272 (BTLA) and are the human equivalent to the cross-presenting murine cDC1 [12].cDC2, also known as CD1c + DC, express CD1c together with CD301 (CLEC10A) and FcεR1α and are the human equivalent to murine cDC2 [10,12,49].However, it was recently shown that CD1c + DC contain two subpopulations, called DC2 and DC3 [31,47,[49][50][51][52].While DC2 represent bona fide cDC, DC3 share transcriptomic as well as functional features with monocytes [53].Flow cytometric differentiation of blood CD1c + DC into DC2 and DC3 can be achieved using the markers CD5 and CD163, respectively [16,31,49,50].However, further classification of CD5 − CD163 − CD1c + DC, especially in lymphoid tissues, is necessary.
In the following, we provide an easy and ready-to-use protocol for analysis of cDC, pDC, and monocyte populations, including recently identified subpopulations of CD1c + DC, isolated from different tissues of the lymphohematopoietic system via a conserved core-panel by flow cytometry.Further, we show an exemplary gating strategy for the identification of dendritic cells in the spleen.

Reagents.
A complete list of reagents is provided in Table 22.23 and BD LSR Fortessa configuration in Table 24.

2.2.3
Step-by-step sample preparation

DAPI
Dissolve DAPI 4´,6-diamidino-2´-phenylindole dihydrochloride (DAPI) in ultrapure water to create a 1 mg/ml stock solution.Store the solution protected from light at 4°C.Dilute the stock solution 1:10,000 in FACS buffer to create a working solution for DAPI staining of cells directly before acquisition at the flow cytometer.

FACS buffer
Add 2% human serum type AB (v/v) to 500 ml of Phosphate buffered saline solution (PBS).

Antibody staining of single cell suspensions from human
spleen for flow cytometry.In 1.2 Preparation of human splenic single cell suspensions of this section, we described how to prepare single cell suspensions from human spleen.
1. Following their preparation, 5 × 10 7 cells per donor are transferred per well to a 96-well V-bottom plate to perform antibody staining enabling flow cytometric analysis of the DC compartment.2. Before centrifugation of the samples, prepare the first staining mix. 3. Therefore, use 50 μl of FACS buffer per sample and add the antibodies listed in Table 25 in the indicated dilution.Please note that antibody dilutions are dependent on the flow cytometer and its setup.Suggested antibody dilutions were optimized for a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer see Table 24).If the cell count strongly differs from the expected cellular yields (>factor 1.5), scale the volume of antibody cocktails up and down accordingly to maintain a constant detector antibodyto-target ratio.Do not stain in less than 35 μl.

Commercially available Fc block solution is used to block
binding to Fc gamma receptors, thereby avoiding unspecific recognition of cell-bound Fc gamma receptors; also other antibodies blocking Fc receptors or sera (mouse or rat, dependent on the source of antibodies) might be suitable.However, optimal dilutions have to be determined.5.After preparation of the staining mix, centrifuge the samplecontaining 96-well V-bottom plates at 520 × g at 4°C for 5 min.6. Discard the supernatant, keep the plate up-side down and dip on a paper towel.
7. Resuspend each sample in 50 μl of primary antibody staining mix including the Fc-block (Trustain FcX, BioLegend) and incubate for 30 min at 4°C. 8. Fill up with 100 μl FACS buffer per well.9. Centrifuge at 520 × g and 4°C for 5 min.For a BD LSR Fortessa as described in this protocol (for detailed configuration, see Table 24), the event rate should not exceed 15,000 events/s.25.Freshly prepared samples can be directly acquired, while thawed samples have to be filtered via 35 μm cell strainers into FACS tubes to avoid clogging of the flow cytometer.26.After performing the cytometer setup, acquire data at an appropriate flow cytometer (table 24).

Data analysis
Data acquisition was performed at a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer, see Table 24).Subsequently, data were analyzed using FlowJo software.In the following, we provide exemplary gating strategies for mononuclear cells from human spleen analyzed with our conserved panel.An overview of markers for DC subpopulations and monocytes/macrophages is provided in Table 27.Figure 6 shows an exemplary gating strategy for the identification of DC in human spleen.

Pitfalls
Problem: The signal resolution of a marker is not high enough to identify positive and negative cells

Potential solutions:
Check, if laser lines and employed filter sets are appropriate for excitation and detection of the employed dyes.Therefore, please refer to Table 26, where we summarized the excitation and emission maxima of the employed dyes.

Top tricks
The current staining protocol includes several markers defining one cell population such as CD1c, CD301 (CLEC10A), and FcεR1A, which are all expressed on human cDC2.The same is true for cDC1 with CD141, XCR1, and CD370 (CLEC9A).Therefore, the panel can be reduced to a core panel by removing some of the redundant markers to use the free channels for analysis of receptor expression on DC subpopulations including costimulatory/inhibitory receptors for the activation status of the cells or cytokine expression.Additionally, the lineage cocktail, which is currently split up into two channels, can be combined in one to allow for the DC analysis with a reduced staining panel.
The current use of the live-dead discriminator DAPI is not compatible with fixation and intracellular staining.To allow for parallel fixation and live-dead cell discrimination employ a fixable viability dye.In particular, the Zombie UV TM Fixable Viability Kit from BioLegend (#Cat 423107) might be attractive, since the dye is usually detected in the same channel as DAPI.But be aware that fixable viability dyes have to be stained separately in PBS without proteins such as BSA or sera.Further, fixable viability dyes such as Zombie UV bind to a lower extent also to living cells.Therefore, titrate Zombie UV appropriately to ensure proper exclusion of dead cells.
Dependent on the tissue, the dump channel containing the lineage markers can be shrinked or expanded.For instance, in thymus tissue CD3 is only expressed on low level in thymocytes.
Here, the inclusion of CD8α is necessary to clearly distinguish cDC2 and thymocytes, while it can be neglected in most other tissues.Further, in tissues with high amounts of B cells (e.g., spleen and tonsils), either the dilution of the B cell-specific antibodies CD19 and CD20 can be lowered or additional B cell-specific markers such as CD21 can be added to improve the differentiation between cDC2 and B cells.
The suggested panel is suitable to analyze DC across various tissues including lymphohematopoietic (spleen, lymph nodes, and thymus) and peripheral tissues (lung, liver) without the need for major modifications.

Summary of the phenotype
The overall phenotype of immune cells covered by the markers included in the panel is detailed in Table 27.The frequency of the analyzed DC subpopulations is covered in Table 28.

Introduction
DC are not only involved in peripheral T cell tolerance [34,35], but also have a role in central tolerance [36][37][38].Therefore, anal-Figure 6. Exemplary gating strategy of the DC compartment in human spleen.After a morphology gate followed by elimination of doublets, dead cells (DAPI + ) as well as T cells (CD3 + and/or CD8 + ), B cells (CD19 + /CD20 + ) and NK cells (CD56 + /CD335 + ) were excluded.To distinguish DC from monocytes/macrophages, CD88 and CD14 were employed, as CD88 is exclusively expressed on monocytes/macrophages.CD14 low cells were also included in the gate as CD14 can be expressed on low level on human DC3.Remaining cells were separated into CD1c + CD123 − , CD1c − CD123 + , and CD1c − CD123 − cells.Human cDC1 were identified in the CD1c − CD123 − compartment by their high expression of CD141 as well as CLEC9A.cDC1 further express XCR1 (histogram plot).Human cDC2 are contained in the CD1c + CD123 − compartment and are positive for both, CLEC10A and FcεR1A.cDC2 were further subdivided into CD5 + CD163 − DC2 and CD5 neg-low CD163 + DC3.pDC (CD303 + Axl − ) and pre-cDC (CD303 + Axl + ) can be identified in the CD1c − CD123 + compartment using Axl and CD303.Data acquisition was performed at a BD LSR Fortessa and data was evaluated utilizing FlowJo software.ysis of thymic DC subpopulations is important in order to unravel how they contribute to negative and positive selection in the thymus.Several reports showed that conventional as well as plasmacytoid DC are present in human thymus [12,[39][40][41][42].
In general, human DC lack the expression of so-called lineage markers, which are expressed on T cells (CD3), B cells (CD19/CD20), NK cells (CD56/CD335), and monocytes (CD14/CD16) but express HLA-DR, one of the human MHC class II molecules [9,12,46].Broadly, DC are further classified into conventional (cDC) and plasmacytoid DC (pDC).pDC can be easily identified by the expression of CD123, CD303, or CD304.However, recent reports show that precursors of cDC (precDC) also express CD123 and CD303 but can be distinguished using the markers Axl as well as Siglec-6 [47,48].While human pDC lack CD11c expression, cDC express it at varying degrees.However, one has to be careful when using CD11c to identify human DC -as monocytes and macrophages highly express CD11c as well.The cDC compartment is composed of distinct subpopulations of DC: cDC1, also known as CD141 + DC, are identified by the high expression of CD141 as well as by the marker CLEC9A, XCR1, and CD272 (BTLA) and are the human equivalent to the crosspresenting murine cDC1 [12].cDC2, also known as CD1c + DC, express CD1c together with CD301 (CLEC10A) and FcεR1α and are the human equivalent to murine cDC2 [10,12,49].However, it was recently shown that CD1c + DC contain two subpopulations, called DC2 and DC3 [31,47,[49][50][51][52].While DC2 represent bona fide cDC, DC3 share transcriptomic as well as functional features with monocytes [53].Flow cytometric differentiation of blood CD1c + DC into DC2 and DC3 can be achieved using the markers CD5 and CD163, respectively [16,31,49,50].However, further classification of CD5 − CD163 − CD1c + DC, especially in lymphoid tissues, is necessary.
In the following, we provide an easy and ready-to-use protocol for analysis of cDC, pDC, and monocyte populations, including recently identified subpopulations of CD1c + DC, isolated from different tissues of the lymphohematopoietic system via a conserved core-panel by flow cytometry.Further, we show an exemplary gating strategy for the identification of DC in the thymus.

Reagents.
A complete list of reagents is provided in Table 29.

Equipment.
Necessary equipment is listed in Table 30 and BD LSR Fortessa configurarion in Table 31.

DAPI
Dissolve DAPI in ultrapure water to create a 1 mg/ml stock solution.Store the solution protected from light at 4°C.Dilute the stock solution 1:10,000 in FACS buffer to create a working solution for DAPI staining of cells directly before acquisition at the flow cytometer.

FACS buffer
Add 2% human serum type AB (v/v) to 500 ml of Phosphate buffered saline solution (PBS).

Antibody staining of single cell suspensions from human thymus for flow cytometry.
In Section 1.3 Preparation of human thymic single cell suspensions of this section, we described how to prepare single cell suspensions from human thymus.
1. Following their preparation, 5×10 7 cells per donor are transferred per well to a 96-well V-bottom plate to perform antibody staining enabling flow cytometric analysis of the DC compartment.2. Before centrifugation of the samples, prepare the first staining mix. 3. Therefore, use 50 μl of FACS buffer per sample and add the antibodies listed in Table 32 in the indicated dilution.Please note that antibody dilutions are dependent on the flow cytometer and its setup.Suggested antibody dilutions were optimized for a BD LSR Fortessa equipped with 355, 405 nm, 488 nm, 561 nm and 640 nm laser lines (for detailed configuration of the used flow cytometer see table 31).If the cell count strongly differs from the expected cellular yields (>factor 1.5), scale the volume of antibody cocktails up and down accordingly to maintain a constant detector antibody to target ratio.Do not stain in less than 35 μl. 4. Commercially available Fc block solution is used to block binding to Fc gamma receptors, thereby avoiding unspecific recognition of cell-bound Fc gamma receptors; also  other antibodies blocking Fc receptors or sera (mouse or rat, dependent on source of antibodies) might be suitable.However, optimal dilutions have to be determined.5.After preparation of the staining mix, centrifuge the samplecontaining 96-well V-bottom plates at 520 × g at 4°C for 5 min.6. Discard the supernatant, keep the plate up-side down and dip on a paper towel.7. Resuspend each sample in 50 μl of primary antibody staining mix including the Fc-block (Trustain FcX, BioLegend) and incubate for 30 min at 4°C. 8. Fill up with 100 μl FACS buffer per well.9. Centrifuge at 520 × g and 4°C for 5 min.10.During the following washing steps, prepare the second staining mix with Streptavidin-BV570.Prepare 50 μl of staining mix per sample by diluting Streptavidin-BV570 1:500 in FACS buffer.11.After centrifugation (step 9), discard the supernatant, keep the plate up-side down and dip on a paper towel.12. Resuspend each sample in 100 μl FACS buffer.13.Centrifuge at 520 × g at 4°C for 5 min.14.Discard the supernatant, keep the plate up-side down and dip on a paper towel.15.Resuspend each sample in 50 μl of the second staining mix (see step 10).16.Incubate for 15 min at 4°C in the dark.17.Fill up with 100 μl FACS buffer per well.18. Centrifuge at 520 × g and 4°C for 5 min.19.Resuspend each sample in 100 μl FACS buffer.20.Centrifuge at 520 × g at 4°C for 5 min.21.Discard the supernatant, keep the plate up-side down and dip on a paper towel.22. Resuspend the samples in 100 μl FACS buffer.23.Samples are ready for acquisition at a flow cytometer.24.Add at least 200 μl of FACS buffer containing DAPI in a dilution of 1:10,000 directly before acquisition.Dependent on the event rate during acquisition, adjust the volume accordingly.For a BD LSR Fortessa as described in this protocol (for detailed configuration, see table 31), the event rate should not exceed 15,000 events/s.25.Freshly prepared samples can be directly acquired, while thawed samples have to be filtered via 35 μm cell strainers into FACS tubes to avoid clogging of the flow cytometer.26.After performing the cytometer setup, acquire data at an appropriate flow cytometer (table 31).

Data analysis
Data acquisition was performed at a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer, see Table 31).Subsequently, data were analyzed using FlowJo software.In the following, we provide exemplary gating strategies for mononuclear cells from human thymus analyzed with our conserved panel.An overview of markers for DC subpopulations and monocytes/macrophages is provided in Table 34.Figure 7 shows an exemplary gating strategy for the identification of DC in human thymus.

Pitfalls
Problem: The signal resolution of a marker is not high enough to identify positive and negative cells

Potential solutions:
Check, if laser lines and employed filter sets are appropriate for excitation and detection of the employed dyes.Therefore, please refer to Table 33, where we summarized the excitation and emission maxima of the employed dyes.

Top tricks
The current staining protocol includes several markers defining one cell population such as CD1c, CD301 (CLEC10A), and FcεR1A, which are all expressed on human cDC2.The same is true for cDC1 with CD141, XCR1, and CD370 (CLEC9A).Therefore, the panel can be reduced to a core panel by removing some of the redundant markers to use the free channels for analysis of receptor expression on DC subpopulations including costimulatory/inhibitory receptors for the activation status of the cells or cytokine expression.Additionally, the lineage cocktail, which is currently split up into two channels, can be combined in one to allow for the DC analysis with a reduced staining panel.
The current use of the live-dead discriminator DAPI is not compatible with fixation and intracellular staining.To allow for parallel fixation and live-dead cell discrimination employ a fixable viability dye.In particular, the Zombie UV TM Fixable Viability Kit from BioLegend (#Cat 423107) might be attractive, since the dye is usually detected in the same channel as DAPI.But be aware that fixable viability dyes have to be stained separately in PBS without proteins such as BSA or sera.Further, fixable viability dyes such as Zombie UV bind to a lower extent also to living cells.Therefore, titrate Zombie UV appropriately to ensure proper exclusion of dead cells.
Dependent on the tissue, the dump channel containing the lineage markers can be shrinked or expanded.For instance, in thymus tissue CD3 is only expressed on low level on thymocytes.Here, the inclusion of CD8α is necessary to clearly distinguish cDC2 and thymocytes, while it can be neglected in most other tissues.Further, in tissues with high amounts of B cells (e.g., spleen and tonsils), either the dilution of the B cell-specific antibodies CD19 and CD20 can be lowered or additional B cell-specific markers such as CD21 can be added to improve the differentiation between cDC2 and B cells.
The suggested panel is suitable to analyze DC across various tissues including lymphohematopoietic (spleen, lymph nodes, and thymus) and peripheral tissues (lung, liver) without the need for major modifications.

Summary of the phenotype
The overall phenotype of immune cells covered by the markers included in the panel is detailed in Table 34.The frequency of the analyzed DC subpopulations is covered in Table 35.

Introduction
Primary human DC were identified at the beginning of the 2000s in the blood using so-called blood DC antigens [13,14].Human tonsils were shown to contain conventional as well as plasmacytoid DC [12,43].As the structure and function of tonsils are comparable to lymph nodes, isolated tonsil DC can be used for analysis of functions of lymphoid tissue-resident DC [44,45].
In general, human DC lack the expression of so-called lineage markers, which are expressed on T cells (CD3), B cells (CD19/CD20), NK cells (CD56/CD335), and monocytes (CD14/CD16) but express HLA-DR, one of the human MHC class II molecules [9,12,46].Broadly, DC are further classified into conventional (cDC) and plasmacytoid DC (pDC).pDC can be easily identified by the expression of CD123, CD303, or CD304.However, recent reports show that precursors of cDC (precDC) also express CD123 and CD303 but can be distinguished using the markers Axl as well as Siglec-6 [47,48].While pDC lack CD11c expression, cDC express it at varying degrees.However, one has to be careful when using CD11c to identify human DC -as monocytes and macrophages highly express CD11c as well.The cDC compartment is composed of distinct subpopulations of DC: cDC1, also known as CD141 + DC, are identified by the high expression of CD141 as well as by the marker CLEC9A, XCR1, and CD272 (BTLA) and are the human equivalent to the crosspresenting murine cDC1 [12].cDC2, also known as CD1c + DC, express CD1c together with CD301 (CLEC10A) and FcεR1α and are the human equivalent to murine cDC2 [10,12,49].However, it was recently shown that CD1c + DC contain two subpopulations, called DC2 and DC3 [31,47,[49][50][51][52].While DC2 represent bona fide cDC, DC3 share transcriptomic as well as functional features with monocytes [53].Flow cytometric differentiation of blood CD1c + DC into DC2 and DC3 can be achieved using the markers CD5 and CD163, respectively [16,31,49,50].However, further classification of CD5 − CD163 − CD1c + DC, especially in lymphoid tissues, is necessary.
In the following, we provide an easy and ready-to-use protocol for analysis of cDC, pDC, and monocyte populations, including recently identified subpopulations of CD1c + DC, isolated from different tissues of the lymphohematopoietic system via a conserved core-panel by flow cytometry.Further, we show an exemplary gating strategy for the identification of DC in the tonsil.

Reagents.
A complete list of reagents is provided in Table 36.

Equipment.
Necessary equipment is listed in Table 37.

DAPI
Dissolve DAPI in ultrapure water to create a 1 mg/ml stock solution.Store the solution protected from light at 4°C.Dilute the stock solution 1:10,000 in FACS buffer to create a working solution for DAPI staining of cells directly before acquisition at the flow cytometer.

FACS buffer
Add 2% human serum type AB (v/v) to 500 ml of phosphate buffered saline solution (PBS).

Antibody staining of single cell suspensions from human tonsils for flow cytometry.
In Section 1.4 Preparation of human tonsillar single cell suspensions of this section, we described how to prepare single cell suspensions from human tonsils.
1. Following their preparation, 5 × 10 7 cells per donor are transferred per well to a 96-well V-bottom plate to perform   antibody staining enabling flow cytometric analysis of the DC compartment.2. Before centrifugation of the samples, prepare the first staining mix. 3. Therefore, use 50 μl of FACS buffer per sample and add the antibodies listed in Table 39 in the indicated dilution.Please note that antibody dilutions are dependent on the flow cytometer and its setup.Suggested antibody dilutions were optimized for a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer see Table 38).If the cell count strongly differs from the expected cellular yields (>factor 1.5), scale the volume of antibody cocktails up and down accordingly to maintain a constant detector antibodyto-target ratio.Do not stain in less than 35 μl. 4. Commercially available Fc block solution is used to block binding to Fc gamma receptors, thereby avoiding unspecific recognition of cell-bound Fc gamma receptors; also other antibodies blocking Fc receptors or sera (mouse or rat, dependent on source of antibodies) might be suitable.However, optimal dilutions have to be determined.5.After preparation of the staining mix, centrifuge the samplecontaining 96-well V-bottom plates at 520 × g at 4°C for 5 min.6. Discard the supernatant, keep the plate up-side down and dip on a paper towel.7. Resuspend each sample in 50 μl of primary antibody staining mix including the Fc-block (Trustain FcX, BioLegend) and incubate for 30 min at 4°C. 25.Freshly prepared samples can be directly acquired, while thawed samples have to be filtered via 35 μm cell strainers into FACS tubes to avoid clogging of the flow cytometer.26.After performing the cytometer setup, acquire data at an appropriate flow cytometer (Table 38).

Data analysis
Data acquisition was performed at a BD LSR Fortessa equipped with 355, 405, 488, 561, and 640 nm laser lines (for detailed configuration of the used flow cytometer, see Table 38).Subsequently, data were analyzed using FlowJo software.In the following, we provide exemplary gating strategies for mononuclear cells from human tonsils analyzed with our conserved panel.An overview of markers for DC subpopulations and monocytes/macrophages is provided in Table 41. Figure 8 shows an exemplary gating strategy for identification of DC in human tonsils.

Pitfalls
Problem: The signal resolution of a marker is not high enough to identify positive and negative cells

Potential solutions:
Check, if laser lines and employed filter sets are appropriate for excitation and detection of the employed dyes.Therefore, please refer to Table 40, where we summarized the excitation and emission maxima of the employed dyes.

Top tricks
The current staining protocol includes several markers defining one cell population such as CD1c, CD301 (CLEC10A), and FcεR1A, which are all expressed on human cDC2.The same is true for cDC1 with CD141, XCR1, and CD370 (CLEC9A).Therefore, the panel can be reduced to a core panel by removing some of the redundant markers to use the free channels for analysis of receptor expression on DC subpopulations including costimulatory/inhibitory receptors for the activation status of the cells or cytokine expression.Additionally, the lineage cocktail, which is currently split up into two channels, can be combined in one to allow for the DC analysis with a reduced staining panel.
The current use of the live-dead discriminator DAPI is not compatible with fixation and intracellular staining.To allow for parallel fixation and live-dead cell discrimination employ a fixable viability dye.In particular, the Zombie UV TM Fixable Viability Kit from BioLegend (#Cat 423107) might be attractive, since the dye is usually detected in the same channel as DAPI.But be aware that fixable viability dyes have to be stained separately in PBS without proteins such as BSA or sera.Further, fixable viability dyes such as Zombie UV bind to a lower extent also to living cells.Therefore, titrate Zombie UV appropriately to ensure proper exclusion of dead cells.
Dependent on the tissue, the dump channel containing the lineage markers can be shrinked or expanded.For instance, in thymus tissue CD3 is only expressed on low level of thymocytes.Here, the inclusion of CD8α is necessary to clearly distinguish Figure 8. Exemplary gating strategy of the DC compartment in human tonsils.After a morphology gate followed by elimination of doublets, dead cells (DAPI + ) as well as T cells (CD3 + and/or CD8 + ), B cells (CD19 + /CD20 + ) and NK cells (CD56 + /CD335 + ) were excluded.To distinguish DC from monocytes/macrophages, CD88 and CD14 were employed, as CD88 is exclusively expressed on monocytes/macrophages.CD14 low cells were also included in the gate as CD14 can be expressed on low level on human DC3.Remaining cells were separated into CD1c + CD123 − , CD1c − CD123 + , and CD1c − CD123 − cells.Human cDC1 were identified in the CD1c − CD123 − compartment by their high expression of CD141 as well as CLEC9A.cDC1 further express XCR1 (histogram plot).Human cDC2 are contained in the CD1c + CD123 − compartment and are positive for both, CLEC10A and FcεR1A.cDC2 were further subdivided into CD5 + CD163 − DC2 and CD5 neg-low CD163 + DC3.pDC (CD303 + Axl − ) and pre-cDC (CD303 + Axl + ) can be identified in the CD1c − CD123 + compartment using Axl and CD303.Data acquisition was performed at a BD LSR Fortessa and data were evaluated utilizing FlowJo software.3 Cell sorting of primary human DC 3.1 Cell sorting of primary human blood DC

Introduction
DC are major regulators of immune responses as they are able to induce T cell responses against invading pathogens or tumors but also maintain peripheral tolerance [46,54].The DC compartment comprises different DC subpopulations: Murine cDC1 are characterized by the dependence on the transcription factor Batf3 and the expression of CD8α (lymphoid tissues) or CD103 (peripheral tissues) as well as the chemokine receptor XCR1, while cDC2 are shown to partially depend on Irf4 and express the surface molecules CD11b as well as Sirpα [46].Furthermore, the DC compartment contains plasmacytoid DC, which depend on ID2 and are identified by the expression of PDCA-1, B220 as well as Siglec-H.
In vivo studies in mice showed that the DC subpopulations differed in functional properties as only cDC1 were able to crosspresent antigens to CD8 + T cells [55][56][57].While cDC1 predominantly induce Th1 responses, cDC2 are prone to Th2 as well as Th17 responses [58].
In humans different DC subpopulations exist as well, which show transcriptional homologies to murine DC subpopulations [9].Human cDC1, also known as CD141 + DC, show high mRNA expression of BATF3 as well as XCR1, while cDC2, also known as CD1c + DC, share the expression of SIRPα and IRF4 with murine cDC2.In order to perform functional analysis with human primary DC, isolation of the different DC subpopulations to high purity and high cell numbers is necessary.After the identification of specific antigens on the different human DC subpopulations, namely CD1c on cDC2, CD303 on pDC, and CD141 on cDC1, positive enrichment using magnetic beads became possible [13,14].However, isolation of the different DC subpopulations from the same donor using magnetic beads will reduce the final cell yield as the cell suspension has to be split up or the enrichment has to be performed in consecutive steps.Therefore, we here present a protocol combining negative enrichment by depletion of unwanted cell lineages followed by cell sorting of DC to high purity [10,11].Further, the panel for cell sorting can be easily adapted to isolate newly defined DC subpopulations such as DC3 (see section 3.1.6Top tricks).

Reagents.
A complete list of reagents is provided in Table 43.Further, blood or LRS cones are needed for the isolation of peripheral blood mononuclear cells (PBMCs) (see Section 1.1 Isolation of mononuclear cells from peripheral blood).

Equipment.
Necessary equipment is listed in Table 44.

Trypan blue
Create a 0.9 % (w/v) NaCl solution by dissolving NaCl in double-distilled H 2 O. Dissolve 0.36 % (w/v) Trypan blue powder in 0.9 % NaCl solution.Sterile filter the solution via a 0.22 μM membrane and store at room temperature.

FACS buffer
Add 2% human serum type AB (v/v) to 500 ml of Phosphate buffered saline solution (PBS).

Staining of the enriched cells.
In order to purify primary human DC, the enriched DC have to be stained with a panel of fluorochrome-conjugated antibodies and subsequently cell sorted.The depletion of lineage-positive cells, such as T cells, B cells, and NK cells, efficiently enriches all DC subpopulations ∼50-fold (see Section 3.1.3.2).Thus, after enrichment of DC from 9 × 10 8 PBMCs (as in this chapter), typically 5×10 7 cells have to be stained for the cell sorting.The enriched cells can either directly be stained in 500 μl of cell sort staining mix (Table 46) or cells can be counted using a Neubauer chamber.In case you have markedly more or less cells (factor >1.5), please adjust the volume of the cell sort staining mix accordingly.
1. Prepare 500 μl of cell sort staining mix (Table 46) to stain the enriched DC for the cell sort.2. Discard the supernatant from the cell pellet (see step 5 in section 3.1.3.2) and resuspend the cells in 500 μl of the prepared cell sort staining mix.Incubate the cells on ice for 30 min in the dark.3.During incubation, you can prepare DAPI-containing FACS buffer (FACS-DAPI) for the cell sort by diluting DAPI 1:10,000 in FACS buffer.Further, add 500 μl of DC medium to four 5 ml polypropylene round bottom tubes, which serve as collecting tubes for the cell sort.4.After incubation, fill the 15 ml Falcon up to 15 ml and centrifuge with 520 × g for 5 min at 4°C.Discard the supernatant and wash again with 10 ml FACS buffer.Centrifuge with 520 × g for 5 min at 4°C.Discard the supernatant and resuspend the cells in 1,000 μl FACS-DAPI buffer.The cells are now ready for cell sorting.

Cell sorting of enriched blood DC.
For this protocol, we describe cell sorting of human blood DC using a BD FACSAria II cell sorter with five laser lines (355, 405, 488, 561, and 635 nm; for detailed configuration see Table 45).To minimize spillover and maximize available fluorochromes for subsequent assays (e.g., costimulatory molecules expression after TLR stimulation), a core panel of markers for identification of DC subpopulations is used.If necessary, other DC subpopulation-defining markers can be easily added (e.g., XCR1-FITC, CD5-PE, CD163-A647).In case cell sorts are performed by yourself, prepare the cell sorter accordingly; i.e., perform a performance check, determine the proper drop delay using Accu Drop Breads, insert the device for parallel sorting of four cell populations, turn on the cooling device and set it to 4°C, and so on.Perform a compensation for all stained colors using the very same antibodies (same vendor, same LOT) as you used for staining (see Section 3.1.3.3.1).

Data analysis
For examples, describing functional assays that can be performed with sorted primary human DC, please refer to [10,11,31,44,47,48,50].The volume of the master mix and the dilution of the antibodies suggested in this chapter are optimized for the specific instrument used in our laboratory.In case the segregation between the different DC is not good enough for cell sorting, you might have to increase the volume of the master mix or lower the dilution of the antibodies.
Further, if the configuration of your cell sorter strongly differs from the configuration of the BD FACSAria II cell sorter described here (see Table 45), you might have to check the compensation or change some of the fluorochromes used.For example, the tandem dye BV711 (CD141) can spill over into the channels BV421 (CD3, CD19, CD20, CD56) as well as A700 (CD14).As BV421 + as well as A700 + cells are excluded before DC are gated, this could lead to a loss of cDC1 as these cells highly express CD141.Thus, you can alternatively use an A647-or APC-labelled αCD141 antibody (#344124, Biolegend; 564123, BD Biosciences; 130-113-314, Miltenyi Biotech).

Potential solutions:
Typically, you should isolate 12 × 10 6 cDC2 and pDC as well as 24 × 10 5 cDC1, if you start with 9 × 10 8 PBMCs.As working with human material, the variability between different donors is high and can lead to markedly higher or lower yield in rare cases.In case you repetitively have lower cell numbers than expected, either the isolation of PBMCs or the enrichment is not working properly.
During preparation of the PBMCs, make sure that you carefully overlay the lymphocyte separation medium with the diluted blood product and collect the whole interphase after the density gradient centrifugation.In addition, verify that the deceleration of the centrifugation was set to zero.
During enrichment of the PBMCs, you might incubate the cells on ice or at 4°C to avoid unspecific binding or ingestion of the magnetic beads to the DC as this might lead to depletion of DC.Make sure that you do not remove the 14 ml round bottom tube from the magnet before you pour off the cell suspension into the 15 ml tube.The unwanted cells are attached to the wall of the 14 ml round bottom tube only as long as the tube is inside the magnet.Before starting with the staining of the cells for the cell Figure 9. Gating strategy for cell sorting of pre-enriched human blood DC.PBMCs of a healthy donor were enriched and stained as described above.Cells were acquired using a BD ARIA II cell sorter.After a morphology gate (FSC-A/SSC-A), doublets (FSC-A/FSC-H) as well as dead cells (DAPI + ) were excluded.Then, lineage (CD3/CD19/CD20/CD56) negative cells were selected and only HLA-DR + cells gated.Monocytes were identified by high expression of CD14.The CD14 − cells were subdivided into CD1c + CD123 − (Q4) cells, CD1c − CD123 + (Q1) cells as well as CD1c − CD123 − cells (Q3).Cells in Q4 were further gated as CD1c + CD11c + cDC2, cells in Q1 as CD123 + CD303 + pDC, and cells in Q3 as CD141 + CD11c int cDC1.According to the frequency, cDC1, cDC2, pDC, and monocytes were assigned to the FL, FR, L, and R channels and subsequently sorted using a 70 μm nozzle with four-way purity using a BD Aria II cell sorter.

Figure 2 .
Figure 2. Mechanical disruption of human spleen tissue.(A) Human spleen tissue before mechanical disruption.(B) Embedding of small pieces (cubes of 2×2×6 mm) of the spleen with capsule in cryomolds filled with O.C.T. tissue tek.(C) Remaining tissue was cut into small pieces (1×1×1 mm) for further mechanical disruption using C-tubes and a Gentle MACS Dissociator followed by enzymatic digestion using Collagenase IV and DNase I.
Figure 2C).4. For digestion of the tissue, use C-tubes and a Gentle MACS Dissociator.Use one C-tube per ∼2 g splenic tissue.Thus, if preparing 10 g of total tissue, use five C-tubes.More than 2 g of tissue can lead to overloading of the C-tubes.In case a Gentle MACS Dissociator is not available in the lab, please refer to section 1.2.5 Pitfalls.5. Add 5 ml of RPMI-1640 to each C-tube as well as 500 μl Collagenase IV and 100 μl DNase I. Tightly close the C-tubes and proceed with mechanical disruption using the Gentle MACS Dissociator.6. Run the program 'm_liver_01.1'and incubate the C-tubes at 37°C for 30 min.Run the program 'm_liver_02.1'and incubate an additional 15 min at 37°C for 30 min.7.After the end of the incubation time, run the program 'm_liver_02.1'again, add 5 ml of washing buffer, and put the C-tubes on ice to stop the digestion of the tissue.8. Prepare one 50 ml tube for each two C-tubes.Insert a 100 μm filter into each 50 ml tube and wet it with 5 ml washing buffer.9. Pipet or pour the cell suspension of one C-tube onto the filter.

Figure 3 .
Figure 3. Mechanical disruption of human thymic tissue.(A) Human thymus tissue before mechanical disruption.(B) Thymus tissue was cut into small pieces (1×1×1 mm) for (C) further mechanical disruption using C-tubes and a Gentle MACS Dissociator followed by enzymatic digestion using Collagenase IV and DNase I. (D) Filtered single cell suspension was overlaid on Lymphocyte Separation Media (1,077 g/ml).(E) After centrifugation without brake, interphase (marked with a black rectangle) containing the mononuclear cells can be collected.

1. 4 . 3
Step-by-step sample preparation 1.4.3.1 Preparation of stocks and solutions.DNase I Dissolve Deoxyribonuclease I (DNase I) in Hank´s balanced salt solution (HBSS) to reach a final concentration of 4200 Units/ml.Sterile filter the solution through a sterile 0.22 μm membrane.Make aliquots.Store at -20°C and avoid freeze-thaw cycles.Use sterile solutions and aseptic techniques.Collagenase IV/ Collagenase D Dissolve Collagenase IV (Col IV) in Hank´s balanced salt solution containing 3 mM CaCl 2 to reach a final concentration of 1500 Units/ml.Sterile filter the solution through a sterile 0.22 μm membrane.Prepare aliquots.Store at -20°C and avoid freezethaw cycles.Use sterile solutions and aseptic techniques.

Figure 4 .
Figure 4. Mechanical disruption of human tonsillar tissue.Human tonsillar tissue retrieved from tonsillectomy was transferred into a sterile cell culture plate.White arrows indicate cauterized tissue that should be removed prior to mechanical disruption using forceps and scalpel.

Figure 5 .
Figure5.Exemplary gating strategy of the dendritic cell compartment in human blood.After a morphology gate followed by elimination of doublets, dead cells (DAPI + ) as well as T cells (CD3 + and/or CD8 + ), B cells (CD19 + /CD20 + ) and NK cells (CD56 + /CD335 + ) were excluded.To distinguish DC from monocytes, CD88 and CD14 were employed, as CD88 is exclusively expressed on monocytes.CD14 low cells were also included in the gate as CD14 can be expressed on low level on human DC3.Remaining cells were separated into CD1c + CD123 − , CD1c − CD123 + , and CD1c − CD123 − cells.Human cDC1 were identified in the CD1c − CD123 − compartment by their high expression of CD141 as well as CLEC9A.cDC1 further express XCR1 (histogram plot).Human cDC2 are contained in the CD1c + CD123 − compartment and are positive for both, CLEC10A and FcεR1A.cDC2 were further subdivided into CD5 + CD163 − DC2 and CD5 neg-low CD163 + DC3.pDC (CD303 + Axl − ) and pre-cDC (CD303 + Axl + ) can be identified in the CD1c − CD123 + compartment using Axl and CD303.Data acquisition was performed at a BD LSR Fortessa and data was evaluated utilizing FlowJo software.

10 .
During the following washing steps, prepare the second staining mix with Streptavidin-BV570.Prepare 50 μl of staining mix per sample by diluting Streptavidin-BV570 1:500 in FACS buffer.11.After centrifugation (step 9), discard the supernatant, keep the plate up-side down and dip on a paper towel.12. Resuspend each sample in 100 μl FACS buffer.13.Centrifuge at 520 × g at 4°C for 5 min.14.Discard the supernatant, keep the plate up-side down and dip on a paper towel.15.Resuspend each sample in 50 μl of the second staining mix (see step 10).16.Incubate for 15 min at 4°C in the dark.17.Fill up with 100 μl FACS buffer per well.18. Centrifuge at 520 × g and 4°C for 5 min.19.Resuspend each sample in 100 μl FACS buffer.20.Centrifuge at 520 × g at 4°C for 5 min.21.Discard the supernatant, keep the plate up-side down and dip on a paper towel.22. Resuspend the samples in 100 μl FACS buffer.23.Samples are ready for acquisition at a flow cytometer.24.Add at least 200 μl of FACS buffer containing DAPI in a dilution of 1:10,000 directly before acquisition.Dependent on the event rate during acquisition, adjust the volume accordingly.

Figure 7 .
Figure 7. Exemplary gating strategy of the DC compartment in human thymus.After a morphology gate followed by elimination of doublets, dead cells (DAPI + ) as well as T cells/thymocytes (CD3 + and/or CD8 + ), B cells (CD19 + /CD20 + ) and NK cells (CD56 + /CD335 + ) were excluded.To distinguish DC from monocytes/macrophages, CD88 and CD14 were employed, as CD88 is exclusively expressed on monocytes/macrophages.CD14 low cells were also included in the gate as CD14 can be expressed on low level on human DC3.Remaining cells were separated into CD1c + CD123 − , CD1c − CD123 + , and CD1c − CD123 − cells.Human cDC1 were identified in the CD1c − CD123 − compartment by their high expression of CD141 as well as CLEC9A.cDC1 further express XCR1 (histogram plot).Human cDC2 are contained in the CD1c + CD123 − compartment and are positive for both, CLEC10A and FcεR1A.cDC2 were further subdivided into CD5 + CD163 − DC2 and CD5 neg-low CD163 + DC3.pDC (CD303 + Axl − ) and pre-cDC (CD303 + Axl + ) can be identified in the CD1c − CD123 + compartment using Axl and CD303.Data acquisition was performed at a BD LSR Fortessa and data was evaluated utilizing FlowJo software.

1 .
Isolate human PBMCs as described in Section 1.1 Isolation of mononuclear cells from peripheral blood.2. Transfer 9 × 10 8 cells into a 14 ml round bottom tube.Centrifuge the cells for 5 min with 520 × g at RT. Discard the supernatant and resuspend the cells in 9 ml EasySep buffer.3. Add 150 μl Fc blocking reagent, 200 μl of Depletion Cocktail A, and 200 μl Depletion Cocktail B (all included in EasySep Pan-DC Pre-Enrichment Kit).Mix the cells by pipetting up and down and incubate at RT for at least 30 min.If the cells settle at the bottom of the tube, resuspend the cells again to facilitate binding of unwanted cells with the depletion cocktail.4. Vortex Dextran Rapidspheres for at least 30 s until the suspension is homogenously brownish.Without washing, add 400 μl of Dextran Rapidspheres to the cell suspension.Incubate for 15 min at RT. 5. Remove the lid from the tube and transfer it inside an Easy-Sep magnet.Incubate for 5 min until the cell suspension is cleared from the magnetic particles, which bind to the wall of the 14 ml round bottom tube during the incubation inside the magnet.Without removing of the tube from the magnet, pour off the cell suspension into a fresh 15 ml Falcon.Centrifuge the tube for 5 min with 520 × g at 4°C.Discard the supernatant and resuspend the cell pellet in FACS buffer.Centrifuge the cells again for 5 min with 520 × g at 4°C.The cells are now enriched and can be stained with fluorochrome-coupled antibodies for cell sorting or FACS-analysis (see Section 3.1.3.3.1).

3. 1 . 5
Pitfalls (consider any relevance for single cell sequencing/OMICS techniques if applicable)Problem: The enrichment of DC is not strong enough to efficiently sort high numbers Potential solutions: Typically, most lineage-positive cells, such as T cells, B cells, NK cells, and also monocytes, are efficiently depleted.The remaining cells should consist of at least 20% cDC2, 20% pDC, and 2% cDC1.In case you detect high amounts of T cells, B cells, NK cells, or monocytes, you should adjust the depletion cocktail (see step 3, section 3.1.3.2) by increasing it to 300 μl as well as the Dextran Rapidspheres (see step 4, section 3.1.3.2) to 600 μl.Alternatively, you can repeat the enrichment by performing steps 3-5 with the enriched cell suspension.As the cells are already preenriched, scale down the depletion cocktail to 25-50 μl (dependent on the size of the pellet) as well as the Dextran Rapidspheres to 50-100 μl.Assure that you incubate the cells at RT.If you perform the enrichment on ice to avoid potential activation of the cells, increase the incubation time to at least 45 min.Problem: The discrimination of the different DC subsets is not possible Potential solutions:

Table 2 .
Necessary equipment

Table 3 .
Expected cellular yields resulting from isolation of PBMCs from LRS cones

Table 4 .
Reagents, enzymes, chemicals, and solutions Sterile filter the solution through a sterile 0.22 μm membrane.Make aliquots.Store at -20°C and avoid freeze-thaw cycles.Use sterile solutions and aseptic techniques.Dissolve Collagenase IV (Col IV) in Hank´s balanced salt solution containing 3 mM CaCl 2 to reach a final concentration of 1500 Units/ml.Sterile filter the solution through a sterile 0.22 μm membrane.Prepare aliquots.Store at -20°C and avoid freezethaw cycles.Use sterile solutions and aseptic techniques.

Table 5 .
Necessary equipment

Table 6 .
Expected cellular yields resulting from single cell suspension generation

Table 9 .
Necessary equipment

Table 10 .
Expected cellular yields resulting from single cell suspension generation

Table 11 .
Adjustments for other lymphoid tissues TissueCollagenase IV/D DNase I Specifics Spleen 500 μl 100 μl Frequently clogging of the cell strainer due to the capsule of the spleen Tonsil 500 μl 100 μl Might be beneficial to remove cauterised tissue before digestion Lymph nodes 500 μl 100 μl For small lymph nodes, digestion in 50 ml Tubes instead of C-tubes; no density gradient centrifugation

Table 12 .
Reagents, enzymes, chemicals and solutions complement activity (inactivation for 30 min at 56°C is also possible but will result in lower activity of the growth factors contained in FCS).Directly filter the warm FCS through a sterile 0.22 μm membrane (Corning #431118) into a sterile storage bottle (Corning #430518) and aliquot into 50 ml portions.Use aseptic techniques during the whole procedure.Aliquoted FCS should be stored at -20°C.Avoid freeze-thaw cycles.

Table 13 .
Necessary equipment

Table 14 .
Expected cellular yields resulting from single cell suspension generation

Table 15 .
Reagents, antibodies, chemicals and solutions Fill up with 100 μl FACS buffer per well.9. Centrifuge at 520 × g and 4°C for 5 min.10.During the following washing steps, prepare the second staining mix with Streptavidin-BV570.Prepare 50 μl of staining mix per sample by diluting Streptavidin-BV570 1:500 in FACS buffer.11.After centrifugation (step 9), discard the supernatant, keep the plate up-side down and dip on a paper towel.12. Resuspend each sample in 100 μl FACS buffer.13.Centrifuge at 520 × g at 4°C for 5 min.14.Discard the supernatant, keep the plate up-side down and dip on a paper towel.15.Resuspend each sample in 50 μl of the second staining mix (see step 10).16.Incubate for 15 min at 4°C in the dark.17.Fill up with 100 μl FACS buffer per well.18. Centrifuge at 520 × g and 4°C for 5 min.19.Resuspend each sample in 100 μl FACS buffer.20.Centrifuge at 520 × g at 4°C for 5 min.21.Discard the supernatant, keep the plate up-side down and dip on a paper towel.22. Resuspend the samples in 100 μl FACS buffer.23.Samples are ready for acquisition at a flow cytometer.24.Add at least 200 μl of FACS buffer containing DAPI in a dilution of 1:10,000 directly before acquisition.Dependent on the event rate during acquisition, adjust the volume accordingly.For a BD LSR Fortessa as described in this protocol (for detailed configuration, see Table

Table 16 .
Necessary equipment

Table 17 .
Detailed configuration of the used BD LSR Fortessa

Table 18 .
First antibody staining mix for flow cytometry

Table 19 .
Optimal excitation and emission values for employed dyes Check, if the defined PMT voltages are adequate.Check, if the correct antibody clone was employed.Perform an antibody titration with your own flow cytometer set up.

Table 21 .
Frequency of analysed DC subpopulations in the blood (N = 7)

Table 23 .
Necessary equipment

Table 24 .
Detailed configuration of the used BD LSR Fortessa

Table 25 .
First antibody staining mix for flow cytometry

Table 26 .
Optimal excitation and emission values for employed dyes Check, if the defined PMT voltages are adequate.Check, if the correct antibody clone was employed.Perform an antibody titration with your own flow cytometer set up.

Table 27 .
Summary of marker expression on analysed cell populations in the spleen

Table 30 .
Necessary equipment

Table 31 .
Detailed configuration of the used BD LSR Fortessa

Table 32 .
First antibody staining mix for flow cytometry

Table 33 .
Optimal excitation and emission values for employed dyes

Table 35 .
Frequency of analysed DC subpopulations in the thymus (N = 7)

Table 37 .
Necessary equipment

Table 38 .
Detailed configuration of the used BD LSR Fortessa

Table 39 .
First antibody staining mix for flow cytometry

Table 43 .
List of needed reagents for the cell sorting of primary human blood DC

Table 45 .
Detailed configuration of the used BD FACSAria II cell sorter

Table 46 .
Cell sort staining mix