1. Isolate CD14⁺ monocytes from peripheral blood mononuclear cells (PBMCs) using the CD14 MicroBeads, human or CD14 MicroBeads, human – lyophilized. Follow the protocol of the MicroBeads data sheet.
   
   Download the data sheet
   
   CD14 MicroBeads, human
2. Determine purity and recovery by labeling cells with CD14-FITC antibodies and analyzing them by flow cytometry on a MACSQuant® Analyzer 10.
3. Assess cell viability through propidium iodide (PI) staining.

Mo-DCs can be generated using various protocol formats, manually and automatically, and at various scales. Here we describe the procedure for 6-well plates as an example.

Day 0: monocyte seeding

1. Culture 3×10⁶ isolated monocytes in 3 mL of complete medium (RPMI 1640, 2 mM L-glutamine, 1% autologous plasma) supplemented^7,8 with 250 IU/mL IL-4 and 800 IU/mL GM-CSF.
2. Incubate the cultures at 37 °C and 5% CO₂ for 2 days.

Day 2: Stimulation 

Resuspend cells in 1.5 mL of complete medium supplemented with the 2-fold concentration of IL-4 and GM-CSF and put back into the original culture. 

Day 6: Stimulation

1. Again, remove a volume of 1.5 mL from the culture and centrifuge.
2. Resuspend cells in 1.5 mL medium supplemented^9,10 with 2000 IU/mL IL-6, 400 IU/mL IL-1β, 2000 IU/mL TNF-α, and 2 μg/mL PGE₂
3. Place the supplemented resuspension back into the original culture and culture for another 24 hours.

Day 7: Mo-DC assessment

Assess the viability, yield, and absolute cell count of mature Mo-DCs (mMo-DCs) by flow cytometry via light scatter signals and PI fluorescence. For the gating strategy, see the figure below.

Dendritic cells have a distinct morphology characterized by many cellular processes. To evaluate whether monocytes assume this morphology during differentiation and maturation to Mo-DCs, we recommend analyzing cells on days 0, 6, and 7 of cell culture, both by microscopy and flow cytometry.

1. Place isolated monocytes, as well as immature Mo-DCs (imMo-DCs) and mMo-DCs, in a 6-well plate and allow to sediment.
2. Capture images of the cells using a light microscope with phase-contrast at a 400× magnification.

Results from representative analysis

On day 0, cells had a spherical shape, which is normal for monocytes. After 6 days, cells showed cytoplasmic protrusions, which were even more pronounced after maturation to day 7. Moreover, size and granularity of the cells increased during differentiation and maturation, which is reflected in increased forward and side scatter signals in the flow cytometry analysis.

1. To analyze cell surface expression of various maturation markers, co-stimulatory molecules, and receptors for chemokines and antigens, label monocytes, imMo-DCs, and mMo-DCs with specific monoclonal antibodies (see table below). Use appropriate isotype controls to assess non-specific antibody binding. 
2. Determine the mean fluorescence intensity (MFI) of the respective markers by flow cytometry on a MACSQuant® Analyzer 10 using the MACSQuantify™ Software.
3. Exclude cell debris and dead cells from the analysis based on scatter signals and PI fluorescence.
   

Results from representative analysis

Flow cytometry analysis of cell surface markers showed that in vitro generated Mo-DCs assumed the typical DC phenotype. During differentiation, monocytes down-regulated the expression of CD14. In addition, mMo-DCs expressed various DC markers that are involved in the formation of immunological synapses between DC
and naive T cells, including the costimulatory proteins CD80 and CD86, the cell adhesion molecule CD54, and
the antigen-presenting molecules MHC I (HLA-ABC) and MHC II (HLA-DR). Mature Mo-DCs also expressed the DC activation markers CD83, CD25, and CD40, which were up-regulated accordingly. CCR7, which is required for migration of DCs to draining lymph nodes, was also up-regulated. The highest expression levels of the antigen uptake receptors CD209 and CD206 were detected on imMo-DCs, corresponding to their antigen uptake function.

Antigen uptake by antigen-presenting cells can occur specifically via receptor-mediated endocytosis or non-specifically via pinocytosis or phagocytosis. To assess the pinocytosis capacity of in vitro generated Mo-DCs in comparison to monocytes, cells are cultured at 37 °C for up to 1 h in the presence of FITC-dextran. Non-specific binding of FITC-dextran to the cell surface is determined by incubating samples on ice throughout the procedure. The MFI of FITC is then analyzed by flow cytometry after various time points, subtracting the MFI of samples on ice from the MFI of samples incubated at 37 °C. 

1. Incubate 1×10⁵ monocytes, imMo-DCs, or mMo-DCs with FITC-labeled dextran (1 mg/mL) in complete medium (RPMI, 2 mM L-glutamine, 1% autologous plasma) for 5, 10, 20, 30, and 60 minutes at 37 °C.
2. Keep a control sample on ice for 60 min to check for non-specific binding of FITC-dextran to the cell surface.
3. After 60 min, wash all samples twice. To do so, centrifuge samples for 5 min at 300×g and 4 °C and resuspend cells with ice-cold PBS supplemented with 1% fetal calf serum (FCS).
4. Repeat the centrifugation step above and resuspended cells in PBS supplemented with 0.5% BSA.
5. Keep samples on ice until flow cytometry analysis.
6. Determine the uptake of FITC-dextran by measuring the mean fluorescence intensity (MFI) of FITC. Exclude dead cells from the analysis by PI fluorescence. Specific uptake of FITC-dextran is calculated by subtracting the MFI of the control sample that was incubated on ice from the MFI of the samples incubated at 37 °C.

Results of representative analysis

Immature Mo-DCs had the highest antigen uptake capacity, reflected in a continuous increase in MFI over the entire time course. Antigen uptake was decreased when Mo-DCs were matured for an additional 2 days (day 9). Monocytes showed negligible antigen uptake capacity compared to Mo-DCs. Additionally, flow cytometry analysis showed that imMo-DCs exhibited the highest expression of the antigen uptake receptors CD206 and CD209. These results correlate with the finding that imMo-DCs lose their antigen uptake function upon maturation.

Upon activation of DCs in the periphery, the cells migrate to the draining lymph nodes where they encounter naive T cells. Migration depends on the expression of CCR7 on the DC surface¹¹. The CCR7 ligand CCL19 acts as a chemoattractant for DCs and is expressed in the lymph node areas characterized by high T cell densities.

1. Resuspend mMo-DCs in RPMI 1640 with 10% FCS at a density of 5×10⁵ cells/mL.
2. Place 200 µL of the suspension in the upper compartment of a 24-well Transwell® Plate.
3. Fill the lower compartment of the Transwell Plate with 600 µL of complete medium supplemented with Human CCL19 (MIP-3β) at different concentrations.
4. After 3 h, harvest the cells contained in the lower compartment of the Transwell Plate and count them by flow cytometry.
   

Results of example analysis

CCR7 expression was up-regulated during Mo-DC maturation. mMo-DCs migrated towards the CCL19 stimulus in a dose-dependent manner.

After migration to the lymph nodes, DCs can induce the proliferation of naive T cells. The capacity of
monocytes and Mo-DCs to induce T cell proliferation can be tested in an mixed lymphocyte reaction (MLR) assay. To this end, allogeneic naive CD45RA⁺CD45RO^– T cells are isolated to high purities by MACS®
Technology, and their plasma membranes are labeled with CellTrace™ Violet. After coculturing with monocytes or Mo-DCs for 7 d, the number of T cells and the CellTrace Violet staining is analyzed by flow cytometry. As with each division of the T cells the dye gets more diluted in the plasma membrane, an increase in unlabeled cells indicates a high T cell proliferation rate.
 

1. Isolate naive allogeneic CD4⁺ T cells from PBMCs using the Naive CD4⁺ T Cell Isolation Kit II, human. Follow the protocol of the kit data sheet. 
   
   Download kit data sheet
   
   Naive CD4+ Cell Isolation Kit II, human
   
   
2. Determine purity of the isolated cells. Label cells with CD4-PE or CD45RO-APC and CD45 RA-PE antibodies and analyze by flow cytometry.
3. Suspend 5×10⁶ purified naive CD4⁺ T cells in 400 μL PBS.
4. Label cells with 100 μL of a 10 μM CellTrace Violet solution for 5 min at room temperature.
5. Wash cells once with 1 mL FCS and three times with 2 mL MLR medium (RPMI 1640, 2 mM L-glutamine, nonessential amino acids, 0.1 mM sodium pyruvate, 5% human AB serum).
6. Resuspend the CD4⁺ T cells in MLR medium at a density of 5×10⁵ cells/mL.
   
7. Suspend monocytes, imMo-DCs, and mMo-DCs in MLR medium at a density of 1×10⁶ cells/mL and
   serially diluted according to the table below.
8. Place 100 µL of each cell dilution in a well of a 96-well plate.
9. Add 100 µL of the labeled T cells into each well and culture for 7 days at 37 °C, 5% CO₂.
10. Determine proliferation of CD4⁺ T cells by measuring the fluorescence of CellTrace Violet by flow cytometry. Exclude cell debris and dead cells from the analysis by scatter signals and PI fluorescence.
    

Results of representative analysis

In contrast to monocytes, both imMo-DCs and mMo-DCs increased the number of CellTrace Violet–
negative T cells in the culture. This correlated with higher T cell numbers, as determined by cell counting.
T cell proliferation was highest when mMo-DCs were used as antigen-presenting cells, which is in line with
the result from the immunophenotyping showing that receptors involved in T cell priming are up-regulated on Mo-DCs upon maturation.

In general, DCs have the capacity to secrete both pro-inflammatory and immunoregulatory cytokines, depending on the stimulus received. T cell–mediated CD40L stimulation of Mo-DCs induces the production
of T_H1-polarizing IL-12, but also the secretion of the immunosuppressive IL-10^12-14. However, for studies towards the development of cancer therapies it is desirable to generate Mo-DC populations secreting high amounts of IL-12 and low amounts of IL-10 as these cells generate a more effective antitumor response via induction of T_H1 cells.

Mo-DCs can be stimulated under various conditions to determine their capacity to secrete IL-12 and IL-10. They can be incubated with soluble Human CD40-Ligand, which form multimers in vitro, or they can be cocultured with a J558L cell line expressing CD40L.

1. Suspend mMo-DCs in RPMI 1640 with 2 mM L-glutamine and 1% autologous plasma at a density of 1×10⁶
   cells/mL.
2. Seed wells of a 96-well plate with 5×10⁵ cells.
3. Add 16 µg/mL Human CD40-Ligand to each well to stimulate the mMo-DCs. For comparison, coculture mMo-DCs with CD40L-expressing J558L cells at different densities (7×10⁴, 14×10⁴, or 28×10⁴ cells/well).
   ▲ Note: The CD40L-expressing J558L cell line was kindly provided by Professor Marina Cella (Washington University, St. Louis, MO, USA)
   
4. Stimulate Mo-DCs for 24 h at 37 °C, 5% CO₂.
5. Collect cell supernatants and centrifuge for 10 min at 300×g to remove any cells.
6. Determine the concentrations of IL-12p70 and IL-10 by specific ELISAs (eBioscience).

Results of representative analysis

Stimulation with soluble CD40L led to production of high levels of IL-12 and low levels of IL-10. Coculture with CD40L-expressing J588L cells resulted in an overall increase in secretion of both IL-12 and IL-10. However, the ratio of IL12/IL-10 was higher after stimulation with soluble CD40L than after coculture with the cell line. Thus, recombinant human CD40L multimers represent an attractive alternative to CD40L-transfected cell lines, allowing for CD40 stimulation of Mo-DCs under defined conditions.

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