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Immunology and Infection

Generation of Monocyte-Derived Dendritic Cells with Differing Sialylated Phenotypes

Published: October 20, 2023 doi: 10.3791/65525


A unique, comprehensive protocol to generate de-sialylated human monocyte-derived dendritic cells (mo-DCs) from isolated peripheral blood mononuclear cells (PBMCs) using a sialidase treatment is presented. Further, methods to assess the phenotypic and functional characterization of mo-DCs and evaluate how sialidase treatment improves the maturation level of mo-DCs are described.


Sialic acids are negatively charged monosaccharides typically found at the termini of cell surface glycans. Due to their hydrophilicity and biophysical characteristics, they are involved in numerous biological processes, such as modulation of the immune response, recognition of self and non-self antigens, carbohydrate-protein interactions, etc. The cellular content of sialic acid is regulated by sialidase, which catalyzes the removal of sialic acid residues. Several studies have shown that sialo-glycans are critical in monitoring immune surveillance by engaging with cis and trans inhibitory Siglec receptors on immune cells. Likewise, glyco-immune checkpoints in cancer are becoming crucial targets for developing immunotherapies. Additionally, dendritic cells (DCs) are envisioned as an important component in immunotherapies, especially in cancer research, due to their unique role as professional antigen-presenting cells (APC) and their capacity to trigger adaptive immune responses and generate immunologic memory. Nevertheless, the function of DCs is dependent on their full maturation. Immature DCs have an opposing function to mature DCs and a high sialic acid content, which further dampens their maturation level. This downregulates the ability of immature DCs to activate T-cells, leading to a compromised immune response. Consequently, removing sialic acid from the cell surface of human DCs induces their maturation, thus increasing the expression of MHC molecules and antigen presentation. In addition, it can restore the expression of co-stimulatory molecules and IL-12, resulting in DCs having a higher ability to polarize T-cells toward a Th1 phenotype and specifically activate cytotoxic T-cells to kill tumor cells. Therefore, sialic acid has emerged as a key modulator of DCs and is being used as a novel target to advance their therapeutic use. This study provides a unique approach to treat in vitro monocyte-derived DCs with sialidase, aimed at generating DC populations with different cell surface sialic acid phenotypes and tailored maturation and co-stimulatory profiles.


Sialic acid-carrying glycans (sialoglycans) have gained significant interest due to their immunomodulatory role. The monosaccharide sialic acid, which is most prevalent in humans in the form of N-acetylneuraminic acid, presents fundamental ligands for lectins with a recognized role in immunology, such as Selectins and Siglecs. These lectins recognize sialoglycans either on the same cell (cis) or on different cells (trans) and play a significant role in host-pathogen interactions and various physiological and pathological cellular activities1,2,3. Furthermore, since sialic acid occupies the terminal positions of cell surface glycoconjugates, it can conceal the underlying structures, thus inhibiting cell-to-cell contact via non-specific repulsive effects or by obstructing detection by other lectins4. The activity of a variety of sialyltransferases (that transfer sialic acids) and of sialidases (that cleave sialic acid bonds) within the cell determines the amount of sialic acid present at the surface. In addition, soluble sialyltransferases and sialidases expressed by the host or pathogens can extrinsically modify the amount of sialic acid on the cell surface5,6.

Aberrant sialylation is a feature of several pathological conditions. In autoimmune diseases, hyposialylation can contribute to unrestrained immune activation and organ damage, since sialic acid helps to discriminate self-antigens and regulate inflammatory responses7. Conversely, hypersialylation results in the over-expression of sialoglycans, such as sialyl-Tn, sialyl-Lewis antigens, polysialic acid, and gangliosides, which constitutes a hallmark of some cancers8,9. Hypersialylation also depends on increased expression of specific enzymes such as N-acetylglucosaminyltransferase (GNT-V), which generates hypersialylated tri- and/or tetra-antennary N-linked glycans, which have been associated with cancer growth and metastasis10. The sialic acid content also regulates protein stability and function, which are key for the role of relevant oncogenic players11. Therefore, increased sialylation can facilitate tumor development, metastasis, drug resistance, and immune evasion. Moreover, the upregulation of sialoglycans enables tumors to interact with inhibitory Siglec receptors on immune cells and avoid immune surveillance. For that reason, sialoglycans are now considered to be glyco-immune checkpoints and attractive therapeutic targets. For instance, inhibitors of the Siglec-immune axis are already in early clinical trials, since the immune cell receptor Siglec (sialic-acid-binding ImmunoGlobulin-like LECtin) plays an immune-inhibitory role12.

Enzymes have been used to modulate the glycan profile as tools for study or for therapeutic strategies13,14. Sialidase has been employed to alter cancer cell malignancy since sialylated glycans such as sialyl Lewis X are critical for cell migration and cancer metastasis15. At the same time, sialidase inhibitors, which impede sialic acid cleavage, have reached clinics for treating sialic acid-dependent viral infections16. Recently, sialic acid modulation has gained further interest due to the critical role of sialic acids as ligands in the Siglec-immune axis, meaning they offer novel means to reduce cancer escape from immune responses. This interest was further strengthened by the 2022 Nobel laureate Bertozzi and her team's contribution of several strategies that selectively cleave diverse sialoglycans and improve anti-cancer immune responses17. Thus, sialidase-based strategies represent a promising modality for glyco-immune checkpoint therapy. The glycophenotype of cells of the immune system is dependent on the type of cell and their activation status. Regarding T-cells, glycans have a key role in the pathophysiological steps of T-cell development and thymocyte selection, T-cell activity, differentiation, and proliferation18. For instance, polylactosamine on glycoproteins influences basal levels of B lymphocytes and T lymphocytes and macrophage activation19. In macrophages, distinct glycan expression patterns have an important role in macrophage recruitment to the tumor microenvironment (TME)20. Hence, the expression of O-linked and N-linked glycans by immune cells could be used as potential glycobiomarkers for therapeutic approaches in the treatment of cancer and autoimmune diseases.

Dendritic cells (DCs) are specific antigen-presenting cells with a unique capacity to trigger immune responses, such as anti-cancer immunity21. DCs must undergo an upregulation of their antigen-presenting MHC molecules to present antigens to T-cells (signal 1), co-stimulatory molecules to activate T-cells (signal 2) and pro-inflammatory cytokines, such as IL-12, to trigger type 1 helper T-cell proliferation (signal 3)22. The resulting immune profile is tightly regulated, and checkpoints are essential for preventing healthy cells from being attacked. Since DCs can stimulate various immune responses against tumor cells, they are used as cell-based vaccines, and a sizable number of clinical studies have demonstrated their potential benefits23,24. After the FDA approved the first DC-based vaccine in 201025,26, many other DC-based vaccines have been developed. DC-based vaccines are mainly produced ex vivo and administered to patients to elicit immune responses against tumors. However, insufficient or brief maturation is currently one of the factors limiting the clinical efficacy of DCs and means expensive cytokine cocktails must be used. Without adequate maturation, DCs cannot activate T-cells in clinical circumstances. Instead, the DCs express immune checkpoints and trigger a tolerogenic immune response that prevents cytotoxic T-cells from acting against tumor cells.

Human DCs have heavily sialylated surfaces, and this sialylation decreases upon maturation and during an overall immune response27. The maturation of DCs can be induced by eliminating these sialic acids with sialidase. Desialylation greatly upregulates various cytokines, including IL-12, due to the translocation of the NF-kB transcription factor to the nucleus6,28. In addition, desialylation improves antigen cross-presentation through MHC-I and anti-tumour immune responses29,30. Accordingly, the knockout of the sialyltransferases ST3Gal.l and ST6Gal.l, which have a major role in DC sialylation, generates a more mature phenotype in murine DCs31.

Sialidase treatment provides a method for stimulating all aspects of DC maturation, including increased antigen presentation, increased expression of co-stimulatory molecules, and increased cytokine production, to address the shortcomings mentioned above and enable DCs to elicit effective responses. This article presents a procedure to obtain viable desialylated human DCs through the use of a bacterial sialidase. De-sialylated DCs show an improved maturation profile and can be used as cell models to boost anti-tumour immune responses in vitro. The DCs are obtained from blood monocytes, which are then differentiated in vitro in the presence of the cytokine interleukin-4 (IL-4) and granulocyte macrophage colony-stimulating factor (GM-CSF). This work also describes lectin-based methods to analyze sialic acid at the cell surface and methods to immunophenotype the DC maturation level. The procedure described here can be used to desialylate other cell types, thus providing an approach to investigate the role of sialoglycans, which are vital glyco-immune checkpoints and relevant in immunomodulation.


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, 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 cytometry29,30.

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 preparation29,30.
  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.

Representative Results

Monocyte isolation and monocyte differentiation into mo-DCs
In accordance with the protocol, human PBMCs were isolated from the buffy coat using density gradient separation with density gradient medium and thoroughly washed. Trypan blue was used to perform viable cell counting on the day of isolation, as described previously in step 1.4.1. Subsequently, CD14+ monocyte isolation was carried out through positive selection. To achieve this, PBMCs were incubated with magnetic beads containing an antibody that recognizes the CD14 antigen. The selected CD14+ monocytes were cultured in a medium supplemented with GM-CSF and IL-4 for 5-6 days27 to differentiate into immature mo-DCs (Figure 1A). The maturation of mo-DCs can be obtained by applying a cocktail of cytokines, including IL-6, IL-1β, TNF-α, and PGE235 (Figure 1A).

During the differentiation process, as a result of IL-4 and GMCSF stimulation, the cell phenotype is expected to change. Data shows that mo-DCs lose expression of surface marker CD14, mainly expressed by monocytes (Figure 1B), and gain significant expression of CD1a, a marker expressed by human DCs36,37. mo-DCs also obtain higher MHC-II (HLA-DR) expression, an antigen-presenting molecule expressed by human DCs and other antigen-presenting cells38 (Figure 1B).

Cell surface sialic acid content
The treatment of mo-DCs with sialidase reduces the sialic acid content on the surface of mo-DCs, which can be confirmed by staining with lectins, which are proteins capable of binding to carbohydrates39. Since the enzyme used removes both α2,3 and α2,6-linked sialic acids from the cell surface, mo-DCs were stained with PNA, which recognizes T antigen-Galβ1-3GalNAcα1-Ser/Thr, as well as MAA and SNA lectins, which bind to α2,3- and α2,6-sialic, respectively. The effectiveness of the sialidase treatment was evaluated by flow cytometry and by confocal microscopy (Figure 2). As shown in Figure 2A, sialidase treatment significantly decreased MAA and SNA binding while increasing PNA staining. The decrease in SNA staining after sialidase treatment was further confirmed by confocal microscopy analysis showing a significantly reduced SNA staining at the cell surface (Figure 2B).

Functional characterization of sialidase-treated mo-DCs
To evaluate how sialidase treatment affects mo-DC functions, the maturation level of mo-DCs was assessed after sialidase treatment. As shown in Figure 3A, the sialidase treatment leads to a significant increase in the expression of the antigen-presenting molecules MHC I and MHC II and the expression of the CD80 and CD86 co-stimulatory molecules. To assess the effect of sialic acid removal on the ability of mo-DCs to induce T-cell responses, sialidase-treated mo-DCs loaded with tumor cell lysates were used to prime autologous T-cells (Figure 3). Next, the profile of the resulting T-cells was characterized based on their capacity to secrete the Th1 cytokine IFN-γ. As shown in Figure 3B, when compared with T-cells primed by fully sialylated mo-DCs, the T-cells primed by sialidase-treated mo-DCs secreted significantly higher levels of IFN-γ. These results suggest that sialidase-treated mo-DCs have improved capacity to prime autologous T-cells.

Cell viability
After sialidase treatment, a viability assay was performed to ensure the treatment was not cytotoxic to the cells. Following treatment, mo-DCs were stained with 7-AAD and Annexin V, to detect nonviable and apoptotic cells, and analyzed by flow cytometry (Figure 4). Data shows no significant difference in cell viability between untreated (Figure 4, left panel) and sialidase-treated cells (Figure 4, right panel).

Figure 1
Figure 1: Differentiation of isolated monocytes into mo-DCs. (A) CD14+ monocytes were isolated from buffy coats and cultured at a concentration of 1.3 x 106 cells/mL at 37 °C. The monocytes were differentiated in medium supplemented with 750 U/mL of IL-4 and 1,000 U/mL of GM-CSF. Microscopic analysis of the morphology of monocytes isolated from human buffy coat at day 0 (top image). Immature mo-DCs; cells were differentiated during a period of 5 days using IL-4 and GM-CSF (middle image). Matured mo-DCs were obtained by using IL-6, IL-1β, TNF-α, and PGE2 cytokines for 24 h (bottom image). Scale bars: 100 µm. (B) The cells were analyzed at day 0, day 2, and day 5 throughout the differentiation period using flow cytometry. The following antibodies were used to characterize cell surface markers: (a-c) CD14; (d-f) CD1a, and (g-i) HLA-DR (MHC class II). The figure shows representative histograms of at least three independent assays. Panel (B) has been modified from Videira et al.40, patent WO2017002045A1. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Sialidase treatment of human mo-DCs to remove α2,6- and α2,3-linked sialic acids from the cell surface. (A) Analysis of mo-DCs by flow cytometry using lectin staining to test the efficacy of sialidase treatment. Human mo-DCs were treated with sialidase (grey bars) or left untreated (white bars) and stained with SNA lectin (recognizing [2,6]-sialic acids), MAA lectin (recognizing [2,3]-sialic acids), and PNA lectin (recognizing T antigen-Galβ1-3GalNAcα1-Ser/Thr). The values represent the mean fluorescence intensity (MFI) of at least three independent assays. Statistical significance was determined using a two-tailed paired t-test (*P < 0.05 or ***P < 0.0001), referring to the difference between the untreated and sialidase-treated DCs. Sialidase treatment decreased MAA binding and increased PNA staining in human mo-DCs, resulting from the removal of α(2,3)-linked sialic acids; the removal of α(2,6)-linked sialic acids after sialidase treatment was detected by a decrease in SNA staining. (B) Confocal microscopy of mo-DCs treated with sialidase and prepared on coverslips for observation. A range of z-stack images was collected from different cells and processed to include the mean staining intensity. Scale bars: 20 µm. Panel (A) has been modified from Silva et al.30; panel (B) has been modified from Silva et al.29. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Sialidase treatment of mo-DCs inducing a higher expression of maturation markers. (A) mo-DCs treated with sialidase showed a higher maturation phenotype than fully sialylated mo-DCs. Flow cytometry was used to evaluate the expression of several maturation markers. The mo-DCs were treated with sialidase for 1 h at 37 °C; the graph values represent the mean fluorescence intensity (MFI) (average ± SEM) of at least three independent assays. Statistically significant differences were calculated using a t-test (*P < 0.05, **P < 0.01), referring to the difference between the untreated and sialidase-treated conditions. (B) Desialylated human mo-DCs loaded with whole tumor antigens induced specific T-cell responses. The mo-DCs were treated with sialidase for 1 h at 37 °C or left untreated, followed by loading with MCF-7 lysates (TL) as a source of whole tumor cell antigens. Co-culture between mo-DCs and autologous T-cells was performed for 4-8 days in the presence of IL-2 (10 U/mL). T-cells primed with desialylated mo-DCs showed significantly higher secretion of the Th1 cytokine, IFN-γ. Following T-cell stimulation with mo-DCs, the cytokines secreted into the co-culture supernatants were measured by ELISA (n = 7). The graph values represent the concentration (pg/mL) (average ± SEM). Statistically significant differences were calculated using a t-test (*P < 0.05). The figure has been modified from Silva et al.30. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Lack of impact of sialidase treatment on the viability of human mo-DCs. Untreated mo-DCs (left panel) and sialidase-treated mo-DCs (right panel) were subjected to dual staining with annexin V and 7-AAD, and the staining was analyzed by flow cytometry. The data showed no significant difference in cell viability between the untreated and sialidase-treated cells, suggesting that mo-DCs can tolerate sialidase treatment and remain viable to exert their immunologic function. The figure has been modified from Silva et al.30. Please click here to view a larger version of this figure.


Monocyte isolation
This manuscript describes a protocol to generate mo-DCs from human-isolated monocytes CD14+ (Figure 1A), followed by performing a sialidase treatment to reduce the sialic acid content on the surface of these cells.

There are different ways to obtain human DCs, such as directly from peripheral blood or tissues or through differentiation from precursors such as stem cells or monocytes. Obtaining DCs differentiated from monocytes isolated from peripheral blood is far more straightforward due to the ease of obtaining high quantities of monocytes compared to other DC sources41. Still, to obtain a high percentage of isolated monocytes, all the protocol steps must be carefully followed. For instance, the density gradient medium may be toxic to the cells, and to prevent cell death, one must avoid prolonged cell contact with the density gradient medium and wash the cells thoroughly. Cell manipulation must be done as quickly as possible to avoid the loss of cell viability. From PBMCs, monocytes can be isolated through positive selection using the magnetic-activated cell sorting (MACS) method, which is a suitable technology for yielding a high number of monocytes. In addition, compared with other monocyte selection methods, mo-DCs derived from MACS-isolated monocytes possess a greater ability to stimulate anti-tumour T-cell activity42. In this protocol, once isolated, the monocytes were incubated with IL-4 and GM-CSF for a period of 5-6 days to achieve the differentiation into immature mo-DCs (Figure 1). The results showed that morphologically (Figure 1A) and phenotypically (Figure 1B), the isolated monocytes differentiated into immature mo-DCs. Moreover, throughout the differentiation, the mo-DCs lost the expression of CD14 markers and gained the expression of CD1a and MHC-II (Figure 1B), which are required for antigen presentation to T-cells.

This isolation and differentiation of monocytes into mo-DCs are limitations to this protocol. The isolation process is a sensitive step that must be carefully and swiftly executed to avoid cell death, and this step must also be done every time mo-DCs are needed for a new experiment. The differentiation process takes 5-6 days which poses a difficulty in terms of employing this method for high-throughput analyses. Nonetheless, the isolation method and using cytokines to differentiate mo-DCs are useful for generating a high number of functional mo-DCs in vitro for experimentation purposes. The mo-DCs generated in this protocol are able to undergo sialidase treatment, flow cytometry, ELISA, confocal microscopy, and so on, thus emphasizing the importance and usefulness of this method30.

Immature mo-DCs and sialidase treatment
Sialidases are essential in sialylation regulation and are responsible for removing sialic acids from the cell surface glycans. In mo-DCs, sialic acid removal by sialidase leads to the maturation of these cells, which increases antigen-cross presentation and subsequent T-cell activation and anti-tumor activity30.

Immature human mo-DCs display a high content of cell surface α(2,6)- and α(2,3)-linked sialic acids27 compared to mature mo-DCs31,43. Furthermore, removing sialic acids by treating mo-DCs with sialidase improves the maturation of the DCs28,30,31. The sialidase selected for this experiment was from the bacterium Clostridium perfringens. Still, other organisms also produce sialidases, such as the bacteria Streptococcus pneumoniae, Vibrio cholerae, or Salmonella typhimurium44, the leech Macrobdella decora45, and even Homo sapiens46, and sialidases from these organisms are also used experimentally. However, each sialidase has different substrate specificities. Additionally, using the sialidase enzyme can have its limitations; for example, the manipulation of mo-DCs during the treatment can further stimulate these cells. Furthermore, the amount of sialidase and the incubation times must be optimized based on the type of cells being used and their sialic acid composition. The sialic acid removal is not a permanent effect but rather a transient phenomenon, because the cell will restore its cell surface sialic acid content. Besides sialidase, there are other methods to reduce the sialic acid molecules at the surface of cells, such as using sialyltransferase inhibitors, gene knockouts of sialyltransferase genes, or metabolic blockade of sialic acid using sialic acid mimetics47,48,49. Nonetheless, these methods may present distinct effects on cells, and besides desialylation, the cell viability must be considered. The sialidase enzyme treatment is a practical method for effectively and transiently removing cell surface sialic acids while maintaining the cell viability.

In this work, sialidase was added to the immature mo-DCs at the concentration of 500 mU/5 x 106 cells/mL, and the cells were incubated at 37 °C for 60 min. The treatment was performed using RPMI-1640 without serum to preserve the cell viability and avoid any interaction between the sialylated molecules present in the serum30. Sialidase treatment can be performed with other buffers besides RPMI, such as 50 mM sodium acetate, pH 5.1 (in the case of C. perfingens sialidase), or PBS50,51,52. Nonetheless, RPMI-1640 is the most common culture medium for DCs as it maintains constant experimental conditions during the procedure, avoids inducing maturation, and reduces any stress that may be caused by sialidase buffers or PBS53,54,55,56. After incubation with sialidase, it is critical to wash the cells thoroughly with a serum-supplemented medium to guarantee that the enzyme reaction has stopped. The presence of sialylated molecules in the serum will compete as substrates for sialidase, thus assuring a rapid reaction stop.

Surface marker characterization by flow cytometry and confocal microscopy
For the determination of the sialic acid profile, in protocol section 3, we utilized lectin staining followed by flow cytometry and confocal laser scanning microscopy. For the cell staining procedure, in both cases, the lectin concentrations and incubation conditions were optimized to avoid cell agglutination and death. It is critical to perform the incubation at 4 °C in buffers containing at least 2% of either FBS or BSA to avoid non-specific binding of the lectins. In this protocol, RPMI-1640 containing 10% FBS was used to maintain constant experimental conditions and avoid cell stress. Regarding confocal microscopy, fixation of the cells prior to staining is essential to preserve the morphology, prevent autolysis, and maintain antigenicity.

The analysis of the mo-DC phenotype by flow cytometry showed that sialidase-treated mo-DCs had a significantly higher amount of PNA lectin bound to the cell surface compared to MMA and SNA lectins, which decreased after the sialidase treatment (Figure 2A). As expected, PNA staining increased, since PNA recognizes non-sialylated antigens, in contrast to MAA and SNA, which bind directly to α2,3- and α2,6-sialic acids, respectively30. This staining confirms the effective removal of sialic acids from the cell surface using this protocol. Another method that can be used to validate the treatment and analyze the cell surface sialic acid content is lectin staining followed by confocal microscopy, as exemplified in Figure 2B.

Besides the former examples, alternative approaches exist to evaluate and characterize sialic acid content, such as lectin probing by western blotting. Alternative sialic acid-specific lectins are also available, such as Siglecs, a group of lectins that have a distinct preference for sialic acid types and linkages57. Besides using lectins in either technique (flow cytometry, microscopy, or western blot), it is also possible to characterize the sialic acid content using antibodies; for instance, α2,8-sialic acids can be assessed by antibodies such as clone 735, which is specific for polysialic acid58. In addition, after sialidase treatment, cells can be functionally tested for their biological or therapeutic efficiency by evaluating their phenotype and ability to activate T-cells40. In fact, as shown in the examples provided, sialidase-treated mo-DCs showed higher maturation phenotype, as well as an elevated expression of antigen-presenting and co-stimulatory molecules.

Furthermore, sialidase-treated mo-DCs can be loaded with antigens and co-cultured with T-cells or other cells and then can be studied regarding the phenotype, cytokine secretion profile, or other features. In the example provided, the data show that sialidase-treated mo-DCs can be loaded with tumor antigens and then used to activate T-cells. In fact, the resulting T-cells showed increased IFN-γ secretion, which is in agreement with previous reports on the effect of sialic acid shortage on boosting the capacity of mo-DCs to activate T-cells27,28,29,30,31.

In conclusion, this protocol shows a feasible, viable, and practical method to generate mo-DCs for sialic acid content manipulation by treatment with sialidase. This protocol presents a methodology that can serve different purposes and applications. This method can not only have a crucial role in understanding the role of sialic acids in the maturation and response of immune cells but can also be used as an immunomodulatory tool.


The authors declare no competing financial interests or other conflicts of interest.


The authors acknowledge funding from the European Commission GLYCOTwinning GA 101079417 and EJPRD/0001/2020 EU 825575; the Fundação para a Ciência e Tecnologia (FCT) Portugal, under grants FCT 2022.04607.PTDC, UIDP/04378/2020, UIDB/04378/2020 (UCIBIO), and LA/P/0140/2020 (i4HB). FCT-NOVA. and Stemmatters were also funded by the Fundo Europeu de Desenvolvimento Regional (FEDER), through the Programa Operacional Regional do Norte (Norte 2020) for the SI I&amp;DT DCMatters project (NORTE-01-0247-FEDER-047212). We acknowledge Biolabs facility at FCT-NOVA and GLYCOVID NOVA Saude.


Name Company Catalog Number Comments
15 mL conical tube AstiK’s CTGP-E15-050 Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase
24-well plate Greiner Bio-one 662 160 Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase
50 mL conical tube AstiK’s CTGP-E50-050 Obtaining Monocyte-derived Dendritic Cells
7-Aminoactinomycin D (7-AAD) BioLegend 420404 Obtaining Monocyte-derived Dendritic Cells
Annexin V Immunotools 31490013 Obtaining Monocyte-derived Dendritic Cells
Attune Acoustic Focusing Flow Cytometer Thermo Fisher Scientific  Determination of Sialic Acid Profile; Maturation Profiling of mo-DCs
BSA Sigma - Aldrich A3294-100G Obtaining Monocyte-derived Dendritic Cells; Determination of Sialic Acid Profile
CD14 (Monoclonal TÜK4) Miltenyi Biotec 130-080-701 Obtaining Monocyte-derived Dendritic Cells
CD80 Immunotools 21270803 Maturation Profiling of mo-DCs
CD86 Immunotools 21480863 Maturation Profiling of mo-DCs
Cell counting slides and trypan blue EVE EVS-050 Obtaining Monocyte-derived Dendritic Cells
Centrifuge Eppendorf 5430 R Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase; Determination of Sialic Acid Profile; Maturation Profiling of mo-DCs
Density gradient medium (Histopaque) Sigma - Aldrich 10771-100ML Obtaining Monocyte-derived Dendritic Cells
EDTA Gibco, ThermoFisher 15400054 Obtaining Monocyte-derived Dendritic Cells
Elisa kit (IFN-γ) Immunotools 31673539 Maturation Profiling of mo-DCs
EVE automated cell count NanoEntek 10027-452 Obtaining Monocyte-derived Dendritic Cells
Fetal bovine serum (FBS) Gibco 10500064 Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase; Determination of Sialic Acid Profile
Granulocyte-macrophage colony-stimulating factor (GM-CSF) Miltenyi Biotec   130-093-864 Obtaining Monocyte-derived Dendritic Cells
Human CD14 microbeads (Immunomagnetic beads) Miltenyi Biotec 130-050-201 Obtaining Monocyte-derived Dendritic Cells
Interleukin (IL)-1β Sigma - Aldrich I9401 Obtaining Monocyte-derived Dendritic Cells
Interleukin (IL)-4 Miltenyi Biotec 130-093-919 Obtaining Monocyte-derived Dendritic Cells
Interleukin (IL)-6 Sigma - Aldrich SRP3096 Obtaining Monocyte-derived Dendritic Cells
L-glutamine Gibco A2916801 Obtaining Monocyte-derived Dendritic Cells
LS column and plunger Miltenyi Biotec 130-042-401 Obtaining Monocyte-derived Dendritic Cells
Maackia amurensis (MAA) lectin (MAA lectin - Biotinylated) Vector labs B-1265-1 Determination of Sialic Acid Profile
MHC-I (HLA-ABC) Immunotools 21159033 Maturation Profiling of mo-DCs
MHC-II (HLA-DR) Immunostep HLADRA-100T Maturation Profiling of mo-DCs
Microtubes AstiK’s PCRP-E015-500 Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase; Determination of Sialic Acid Profile; Maturation Profiling of mo-DCs
Neuraminidase (Sialidase) Roche 11585886001 Treatment of Cells with Sialidase
Non-essential amino acids (NEAA) Gibco 11140-050 Obtaining Monocyte-derived Dendritic Cells
Paraformaldehyde (PFA 2%) Polysciences Europe 25085-1 Determination of Sialic Acid Profile; Maturation Profiling of mo-DCs
Paraformaldehyde (PFA 4%) Biotium 22023 Determination of Sialic Acid Profile
Pasteur pipettes Labbox PIPP-003-500 Obtaining Monocyte-derived Dendritic Cells
Peanut (Arachis hypogaea) Agglutinin (PNA) lectin (PNA lectin - FITC) Vector labs FL-1071 Determination of Sialic Acid Profile
Penicillin/streptomycin Gibco 15140163 Obtaining Monocyte-derived Dendritic Cells
Phosphate Buffered Saline (PBS) NZYTech   MB18201 Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase; Determination of Sialic Acid Profile; Maturation Profiling of mo-DCs
Prostaglandin E2 (PGE2) Sigma - Aldrich P0409 Obtaining Monocyte-derived Dendritic Cells
RBC lysis buffer BioLegend 420302 Obtaining Monocyte-derived Dendritic Cells
RPMI-1640 medium (containing 11.1 mM glucose) Gibco 31870074 Obtaining Monocyte-derived Dendritic Cells; Treatment of Cells with Sialidase; Determination of Sialic Acid Profile
Sambucus nigra lectin (SNA lectin - Biotinylated) Vector labs   B-1305-2 Determination of Sialic Acid Profile
Sambucus nigra lectin (SNA lectin - FITC) Vector labs FL-1301-2 Determination of Sialic Acid Profile
Sodium pyruvate Thermofisher 11360-070 Obtaining Monocyte-derived Dendritic Cells
SpectroMax190 Molecular Devices Maturation Profiling of mo-DCs
Streptavidin-PE BioLegend   405203 Determination of Sialic Acid Profile; Maturation Profiling of mo-DCs
Tetramethylbenzidine (TMB) Sigma - Aldrich T0440 Maturation Profiling of mo-DCs
Tumour necrosis factor-α (TNF-α) Sigma - Aldrich H8916 Obtaining Monocyte-derived Dendritic Cells
Zeiss LSM710 confocal microscope Zeiss Determination of Sialic Acid Profile



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Monocyte-derived Dendritic Cells Sialylated Phenotypes Sialic Acids Immune Response Modulation Immune Surveillance Glyco-immune Checkpoints Immunotherapies Dendritic Cells (DCs) Antigen-presenting Cells (APC) Maturation Of DCs Sialic Acid Content T-cell Activation
Generation of Monocyte-Derived Dendritic Cells with Differing Sialylated Phenotypes
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Luz, V. C. C., Silva, Z., Sobral,More

Luz, V. C. C., Silva, Z., Sobral, P., Tanwar, A., Paterson, R. L., Videira, P. A. Generation of Monocyte-Derived Dendritic Cells with Differing Sialylated Phenotypes. J. Vis. Exp. (200), e65525, doi:10.3791/65525 (2023).

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