Generation of Monocyte-Derived Dendritic Cells with Differing Sialylated Phenotypes

Cells were isolated from the buffy coats of healthy anonymous blood donors, who were volunteers provided by the national blood bank, Instituto Português do Sangue e da Transplantação (IPST), after written and informed donor consent was obtained (IMP.74.52.4). The blood use was approved by the ethics committee (IPST 30072015), according to directive 2004/23/EC on standards of quality and safety for the donation, procurement, testing, processing, preservation, storage, and distribution of human tissues and cells (Portuguese Law 22/2007, June 29). The IPST biobank collects and stores blood in a specific plastic collection bag containing citrate phosphate dextrose (CPD), a preserving and anticoagulant solution, to maintain the blood's integrity until processing. To assess if the biological material is appropriate for manipulation, a serological control is performed for Treponema pallidum, hepatitis B virus (HBV), hepatitis C virus (HCV), and human immunodeficiency virus (HIV), all of which have to be negative. For the present study, the buffy coat was provided by IPST for investigation purposes, along with information regarding the collection date, serological results, blood type, and age of the donor³². The buffy coat can remain at room temperature for a maximum of 1 day.

1. Obtaining monocyte-derived dendritic cells

NOTE: It is important to mention that when human peripheral blood is being manipulated, one should consider specific universal safety precautions and appropriate material disposal. Before starting, confirm all the reagents and materials necessary are prepared and ready to use.

1. Isolation of peripheral blood mononuclear cells
   1. Access the human buffy coat.
      NOTE: Buffy coat is a by-product derived from blood collected via leukapheresis³², which is enriched in white blood cells through centrifugation. All the steps were performed in a vertical flow chamber biosafety cabinet (BSC).
   2. Open the buffy coat packaging by cutting the sealed outlet tube with a scalpel, and transfer the contents into a 50 mL tube. Transfer 7 mL of buffy coat sample per sterile 15 mL conical tube, and add 6 mL of phosphate-buffered saline solution (PBS) to perform a preliminary wash. This initial wash step is necessary in order to clean the sample from the considerable amount of red blood cells (RBC) and plasma so that the sample is optimized for gradient separation with a density gradient medium (see the Table of Materials).
   3. Centrifuge the tube at room temperature for 10 min at 1,100 x g in a centrifuge with a swing rotor and with the brake off (see the Table of Materials).
   4. After centrifugation, collect the leukocyte suspension (the white ring between the plasma and RBCs) with a Pasteur pipette, and transfer it to a new sterile 15 mL conical tube.
   5. Fill the leukocyte suspension up to 10 mL with PBS to aid the next separation step, and mix by gently pipetting up and down.
   6. Prepare the density gradient medium (density: 1.077 g/mL) solution: Place 3 mL of density gradient medium into a new sterile 15 mL conical tube, and let it warm to room temperature.
   7. Add 5 mL of the diluted leukocyte suspension (from step 1.1.5) into the conical tube containing the density gradient medium (5:3) to perform density gradient separation. Add the sample slowly, drop by drop, using the tube walls to avoid disturbing the density gradient medium.
   8. Gradient separation: Centrifuge the density gradient medium suspension at room temperature for 30 min at 1,100 x g in a centrifuge with a swing rotor and with the brake off.
   9. After centrifugation, carefully remove the conical tubes from the centrifuge. After this step, a range of well-defined layers are visible, including the following, starting from the bottom: a red layer (RBCs and granulocytes), density gradient medium, a thin pale layer of peripheral blood mononuclear cells (PBMC), and plasma.
   10. Collect the thin layer of PBMCs using a Pasteur pipette, and avoid taking up the density gradient medium below or too much plasma above. Place the PBMC sample into a new 50 mL conical tube, fill it up to 25 mL with PBS, and mix the sample by gently pipetting up and down.
   11. Centrifuge the samples at room temperature for 10 min at 600 x g (normal brake) to wash off residual cells and debris, and discard the supernatant by carefully inverting the tube.
       NOTE: If there is too much red blood cell contamination, which is observable when the cell pellet or the buffy coat is not fully separated or appears reddish, it is recommended to lyse the remaining RBCs. In this case, add 5 mL of RBC lysis buffer (see the Table of Materials), mix thoroughly, and incubate for 5 min. Fill up to 40 mL with PBS, centrifuge the samples at room temperature for 10 min at 900 x g (normal brake), and discard the supernatant by carefully inverting the tube.
   12. Fill up the sample to 10 mL with PBS, and take an aliquot to count the cells. To remove platelets, centrifuge at room temperature for 5 min at 400 x g (normal brake), and discard the supernatant by carefully inverting the tube.
       NOTE: In case there is a substantial number of platelets, centrifuge at room temperature for 10 min at 200 x g (normal brake) twice. The platelets are identified by visualizing the sample while counting the cells.
2. Monocyte isolation by immunomagnetic separation
   1. Prepare the microbeads buffer by supplementing PBS with 0.5% bovine serum albumin (BSA) and 2 mM ethylenediaminetetraacetic acid (EDTA). Sterilize the solution by filtration (0.2 µm), and keep the buffer refrigerated (2-8 °C).
   2. Perform monocyte CD14⁺ isolation by magnetic-activated cell sorting (MACS).
      1. After cell counting using an automated cell counter (step 1.4.1), calculate the appropriate volume of microbeads buffer and CD14 immunomagnetic beads (see the Table of Materials) required. Ensure these solutions are kept on ice. Add 80 µL of microbeads buffer per 1 x 10⁷ cells and 20 µL of CD14 beads per 1 x 10⁷ cells.
      2. Resuspend the cell pellet in the previously determined volumes, and incubate for 15 min at 4 °C (2-8 °C).
         NOTE: In case verification of the monocyte levels in the PBMC samples is needed, perform a flow cytometric analysis using staining antibodies (e.g., CD14 [Monoclonal TÜK4]). Follow step 3.2 for details on the flow cytometric analysis.
      3. Add 1-2 mL of microbeads buffer per 1 x 10⁷ cells, centrifuge at room temperature for 10 min at 600 x g (normal brake) to remove unbound beads, and discard the supernatant by carefully inverting the tube.
      4. Prepare the LS column. The LS columns contain ferromagnetic spheres that, when placed onto a magnet, allow positive, gentle retention of magnetically labeled cells³³. Immediately before use, place an LS column (see the Table of Materials) on the magnet, rinse with 3 mL of microbeads buffer without completely drying, and immediately proceed to the next step.
         NOTE: Never let the column dry out during the procedure to avoid compromising the yield.
      5. Resuspend the cell pellet in 500 µL of microbeads buffer per 1 x 10⁸ cells. If the cell number is higher than 4 x 10⁸, use a 40 µm cell strainer to prevent cell aggregation.
      6. Add the cell suspension to the column inlet, place a 15 mL conical tube below the column outlet to collect the negative cell fraction, and wash the column three times with 3 mL of microbeads buffer. The negative fraction comprises the cells that were not collected with the CD14 beads (i.e., the CD14⁻ cells).
      7. After the final wash, remove the column from the magnet, place it on a sterile 15 mL conical tube, pipette 5 mL of microbeads buffer into the column inlet, and immediately insert the syringe plunger into the column inlet and push to dispense the target cells.
      8. Collect the magnetically labeled cells (CD14⁺ cells), and take an aliquot to count the cells, as described in step 1.4.1.
      9. Centrifuge both cell fractions, CD14⁻ and CD14⁺ cells, at room temperature for 10 min at 600 x g (normal brake). Discard the supernatant, retain the CD14⁺ fraction for the next steps, and store the CD14⁻ fraction for future assays, such as co-culture assays, if necessary. If required, the cells from the CD14⁻ fraction can be cryopreserved in RPMI-1640 20% FBS and 10% DMSO at −80 °C.
3. Monocyte differentiation into dendritic cells
   1. Prepare complete RPMI-1640 medium by supplementing the RPMI-1640 base medium (containing 11.1 mM glucose) with 10% fetal bovine serum (FBS), 1% of 2 mM of L-glutamine, 1% non-essential amino acids (NEAA), 1% sodium pyruvate, and 1% of 100 µg/mL penicillin/streptomycin (see the Table of Materials).
   2. Perform monocyte differentiation into mo-DCs, which occurs over ~5-6 days.
      1. Calculate the volume of medium necessary for the number of CD14⁺ cells obtained, and plate the cells according to the following experiment setup.
         NOTE: In this protocol, the cells were plated at a concentration of 1.3 x 10⁶ cells/mL to take into consideration cell death and measurement errors, and the medium was prepared by adding 1,000 U/mL of GM-CSF and 750 U/mL of IL-4 (see the Table of Materials) into a complete medium and mixing it thoroughly.
      2. Add the appropriate volume of medium to the CD14⁺ cells, and resuspend by pipetting up and down with a Pasteur pipette. Plate the cell suspension into 24-well plates (per well: 1.3 x 10⁶ cells/mL), and incubate in a culture incubator at 37 °C with 5% CO₂.
      3. Change the culture medium, and supplement it with fresh cytokines every 2-3 days (usually once per differentiation process). To perform this, carefully remove half of the culture medium without disturbing the cells. Add the same amount of fresh medium with the appropriate concentration of cytokines, as described previously in the note of step 1.3.2.1, and incubate for the remaining differentiation period.
         NOTE: DCs, when differentiating from monocytes, are loosely adherent cells. Fully differentiated immature mo-DCs are spindle-shaped, free-floating, and loosely adherent cells. The cells may also form rosettes, especially when mature³⁴.
      4. To collect the cells after differentiation, use a micropipette to transfer the entire cell suspension to a sterile conical tube, and wash the wells of the plate twice with PBS, tapping gently on the bottom (Figure 1A).
         NOTE: Avoid collecting the heavily adherent cells, as these are probably macrophages. To avoid improper cell maturation or activation, ensure the cells are handled with extreme care.
      5. Centrifuge the cells at room temperature for 10 min at 180 x g (normal brake) to remove any residues or dead cells, and resuspend in the appropriate medium/buffer for the experimental setup.
   3. Perform the maturation of mo-DCs.
      1. In case the maturation of mo-DCs is required, use a well plate or flask, taking into consideration the cell concentration example that was used previously (1.3 x 10⁶ cells/mL), and administer a cytokine cocktail by supplementing the medium with a cytokine cocktail comprising IL-1β (10 ng/mL), IL-6 (1,000 U/mL), prostaglandin E2 (PGE2; 1 µg/mL), and tumor necrosis factor-α (TNF-α; 10 ng/mL) (see the Table of Materials). Incubate the cells at 37 °C with 5% CO₂ for 24 h or 48 h.
4. Cell counting and viability
   1. Perform cell counting and trypan blue staining.
      1. To determine the number of cells and the viability of a cell suspension, take an aliquot of 10 µL from the cell suspension, and mix it with 10 µL of trypan blue (1:1 dilution).
      2. Take 10 µL of the previous mix, and use the automated cell counter to count the number of cells according to the manufacturer's instructions.
         NOTE: If the concentration of cells is too high, dilute the aliquot, and after the cell counting, consider the dilution factor in the calculations.
      3. Adjust the cell number and medium/buffer for the experimental setup.
   2. Determine the cell viability and apoptosis³⁰.
      NOTE: In this work, following the sialidase treatment (section 2), the viability assay was performed.
      1. Stain the mo-DCs with 5 µg/mL 7-aminoactinomycin D (7-AAD) and annexin V, and determine the apoptosis according to the manufacturer's instructions (see the Table of Materials).
      2. Analyze the results using flow cytometry^29,30.

2. Treatment of the cells with sialidase

NOTE: After the differentiation into mo-DCs, on the sixth day, the cells are ready for the sialidase treatment assay.

1. Considering the desired experimental setup, collect ~10 x 10⁶ mo-DCs from 10 wells of the 24-well plates with 1.3 x 10⁶ cells/well, and transfer them into a new sterile 15 mL conical tube.
   NOTE: Assume some cell loss; typically, at this stage, the concentration found is 1.3 x 10⁶ cells/mL because the mo-DCs and their precursors do not proliferate and experience 20% viability loss during differentiation into mo-DCs.
2. Centrifuge at room temperature for 5-7 min at 300 x g (normal brake), and discard the supernatant to remove dead cells and debris.
3. Add 10 mL of RPMI-1640 medium (containing 11.1 mM glucose), centrifuge at room temperature for 4 min at 300 x g (normal brake), discard the supernatant, add 2 mL of RPMI-1640, and mix thoroughly.
4. Place 1 mL of cells in RPMI-1640 into new sterile microtubes, #1 and #2; each microtube will contain approximately 5 x 10⁶ cells.
5. To microtube #1, add 500 mU of sialidase from Clostridium perfringens (see the Table of Materials). To microtube #2, add mock-treated sialidase, which is a negative control, to confirm if the effects observed are directly related to sialic acid removal and are not due to artifacts. The mock-treated sialidase is heat-inactivated sialidase, which is obtained by boiling the enzyme for 20 min at 100 °C.
6. Incubate for 60 min at 37 °C.
7. After incubation, place the cells into new sterile 15 mL conical tubes with the same numeration, #1 and #2. Add around 4 mL of complete RPMI-1640 medium (containing 10% FBS) to both tubes to stop the enzymatic reaction.
8. Centrifuge at room temperature for 4 min at 300 x g (normal brake), and discard the supernatant.
9. Add 5 mL of complete RPMI-1640 medium to each tube, and plate 1 mL of cells per well.

3. Determination of the sialic acid profile

1. Lectin staining
   1. Collect and wash the cells at room temperature for 5 min at 300 x g (normal brake).
   2. Resuspend the cells in RPMI-1640 + 10% FBS, and distribute the cells (100,000/100 µL) into the microtubes.
   3. Perform staining for flow cytometry in RPMI-1640 with 10% FBS using a concentration of 0.01 mg/mL for each lectin: Sambucus nigra (SNA) lectin, peanut agglutinin (PNA) lectin, and Maackia amurensis (MAA) lectin (see the Table of Materials). Incubate for 30 min at 4 °C.
   4. Wash the cells with 1 mL of PBS containing 10% FBS or 10% BSA, and centrifuge at room temperature for 4 min at 300 x g (normal brake).
   5. To the cells stained with the biotinylated lectins, add 0.0005 mg/mL streptavidin-PE (see the Table of Materials), and incubate for 15 min at room temperature in the dark. Wash the cells with 1 mL of PBS, and centrifuge at room temperature for 4 min at 300 x g (normal brake).
   6. Discard the supernatant, and, to each tube, add 300 µL of 2% paraformaldehyde (PFA 2%). Protect the tubes from light, and, if required, store at 4 °C until data acquisition.
   7. Acquire the data using a flow cytometer within 1 week of sample preparation^29,30.
2. Flow cytometry
   1. Resuspend the cells in 1 mL of PBS, and acquire the sample with a flow cytometer for immediate data acquisition.
   2. For delayed data acquisition, resuspend in 300 µL of 2% PFA, and acquire the data within 1 week.
3. Confocal laser scanning microscopy
   1. Plate the cells on 12 mm diameter polylysine-coated glass coverslips, and incubate for 5 min at room temperature.
   2. Centrifuge the coverslips at room temperature for 1 min at 100 x g (normal brake) to promote cell adhesion.
   3. Fix at room temperature for 30 min with 4% PFA before washing with 1% BSA in PBS.
   4. Use FITC-conjugated SNA lectin (0.01 mg/mL) to stain α2,6-linked sialic acids on the cell surfaces (see the Table of Materials).
   5. Acquire images on a confocal microscope (see the Table of Materials).
   6. After Z-stack processing, select representative confocal cross-section images.
   7. Analytically quantify the staining intensity using the corrected total cell fluorescence (CTCF).
      ​NOTE: CTCF = Integrated density − (Area of selected cell × Mean fluorescence of background readings)²⁹.

4. Maturation profiling of mo-DCs

1. Antibody staining and flow cytometry
   1. Collect a new sample of the cells of interest to perform antibody staining. Wash the cells at room temperature for 5 min at 300 x g (normal brake), and distribute cells into the microtubes (100,000 cells per tube).
   2. Perform staining for flow cytometry using the desired antibodies (ab), MHI-I, MHC-II, CD80, and CD86 (see the Table of Materials).
   3. Incubate the fluorescence-conjugated ab for 15 min at room temperature in the dark.
   4. Wash the cells with 1 mL PBS, and centrifuge at room temperature for 5 min at 300 x g(normal brake).
      NOTE: If using unlabeled ab, add fluorescently conjugated secondary ab, and incubate in the dark for 15 min according to the manufacturer's instructions. Wash the cells with 1 mL of PBS, and centrifuge at room temperature for 5 min at 300 x g (normal brake).
   5. To all the microtubes, add up to 100 µL of PBS, resuspend the cells in 300 µL of 2% paraformaldehyde (PFA 2%), and keep the tubes in the dark at 4 °C until the data acquisition.
   6. Acquire the data using a flow cytometer.
      NOTE: After staining and fixation, the samples can be acquired by flow cytometry immediately or within a 1 week period. In this case, store the tubes at 4 °C in the dark.

5. Enzyme-linked immunosorbent assay (ELISA)

NOTE: In this work, the IFN-γ production was measured using the ELISA assay following the manufacturer's instructions (see the Table of Materials).

1. For coating the plate in a coating buffer, dilute the capture antibody (1:100, capture antibody in PBS), transfer 100 µL of this working solution to each well, and incubate overnight at room temperature.
2. Discard the capture antibody completely.
3. Add the blocking buffer (e.g., PBS + 2% BSA + 0.05% Tween20), and incubate for 1 h at room temperature before removing the blocking buffer.
4. Add the standard and sample, with the respective mix and dilutions, and incubate for 2 h at room temperature. Wash five times with washing buffer.
5. Add the biotinylated detector antibody, and incubate for 2 h at room temperature, followed by five washes.
6. Add poly-HRP-streptavidin-HS, and incubate for 30 min at room temperature, followed by five washes with washing buffer.
7. Add TMB substrate (see the Table of Materials), and incubate for up to 60 min at room temperature, taking into account the test system being used. Wash five times with washing buffer.
8. Read the samples on a microplate reader at 450 nm.