Reprograming Model of Human Monocyte-derived Macrophages for In-vitro Assays

Monocytes and macrophages are very plastic and reprogrammable immune cells crucial for health and disease. They sense and respond to a broad range of stimuli by adopting specific differentiation programs and phenotypes. We have standardized an in-vitro model to study macrophage polarization and reprogramming, providing a valuable tool for research.

Cells of the monocyte-macrophage lineage are multifunctional and found in almost all body tissues. They coordinate innate and adaptive immunity´s initiation and resolution phases, significantly affecting protective immunity and immune-mediated pathological injury. While tissue-resident macrophages are key players in maintaining homeostasis in a steady state, large amounts of monocytes are recruited from the peripheral blood into the tissue following damage or inflammatory insults. Monocyte-derived macrophages (M-DM) can differentiate into many dynamic subtypes, and their phenotypes and functions depend on the local tissue environment.

To compare different stimuli or environmental conditions during M-DM differentiation and polarization, we standardized an in-vitro model of human nonpolarized M-DM M0 and some cardinal cytokine-polarized macrophages, IFNγ/LPS-derived M1, IL-4-derived M2a, and IL-10 or dexamethasone-derived M2c, to analyze skewing reprogramming of M-DM by flow cytometry and real-time PCR. We found that CD64, CD206, CD163, CD14, and MERTK can clearly discriminate unpolarized M0 and polarized M1, M2a, and M2c by flow cytometry. Moreover, we defined IRF1 and CXCL10 as specific genes for classical IFNγ/LPS-derived M1-, IRF4-, CCL22-, and TGM2-specific transcripts for IL-4-derived M2a, and the MERTK gene for dexamethasone-derived M2c. To summarize, our standardized M-DM protocol could give the cardinal in-vitro map to analyze the differentiation and polarization of human M-DM under diverse stimuli.

Macrophages, first identified by Metchnikoff for their phagocytic ability, are ancient cells fundamental to Metazoan life. Found ubiquitously in adult mammals, they exhibit remarkable anatomical and functional diversity. As part of the mononuclear phagocytic system, alongside dendritic cells and monocytes, macrophages play crucial roles in various biological processes, from development and homeostasis to immune responses against pathogens^1,2. Resident macrophages, specialized for their tissue microenvironments, act as sentinels, monitoring tissue health and responding to physiological changes and external threats. Their limited plasticity is thought to be an evolutionary adaptation to maintain tissue homeostasis. In contrast, recruited monocytes are more flexible and can differentiate into diverse macrophage phenotypes during inflammation. This dual influence of inflammation and tissue niche shapes the functional diversity of macrophages³. Thus, depending on the local tissue milieu, monocyte-derived macrophages (M-DM) can differentiate into many subtypes. The local metabolites, growth factors, cytokines, and cell-cell interaction^4,5,6 outline their phenotypes and functions. Furthermore, macrophages are key producers of factors that dampen inflammation and drive re-vascularization and tissue repair^7,8.

Given that at least three main arms control polarization (extrinsic, intrinsic, and tissue environment conditions), macrophage polarization should be viewed as multidimensional². The main obstacles and pitfalls in describing macrophage differentiation and polarization are the heterogeneous experimental conditions across the literature and the lack of consensus on defining macrophage terms in in vitro and in vivo experiments⁹. Nonetheless, some unification of experimental standards has started for diverse experimental scenarios.

To better understand the human macrophage field, we need standardized and well-defined in-vitro models of M-DM to link, differentiation, polarization, phenotype, and re-programming, as well as specific functions. Our goal was to set a standardized in-vitro model of human nonpolarized M-DM M0 and cytokine-polarized macrophages to study skewing reprogramming of M-DM by flow cytometry and real-time PCR. Although macrophage polarization is a dynamic process because their highly plasticity and ability of integrating multiple signals from their environment, we characterized here the nonpolarized M-DM (M0) and some in-vitro polarization states, as the classical pro-inflammatory, commonly recognized also as M1, induced with IFNγ and LPS. Similarly, the tissue repair and regulatory M2 macrophages can be defined as M2a when induced with IL-4 and M2c when stimulated with dexamethasone or IL-10. These polarizing states capture a snapshot of the wide M-DM milieu spectrum in time and space but give us some cardinal points to set the analytic map where to locate the testing conditions.

Despite the complex and dynamic combination of surface markers and transcriptional programs, when analyzing in vitro M-DM, we have set that CD64, CD206, CD163, CD14, and MERTK can separate unpolarized M0 condition, from IFNγ/LPS-induced M1, IL-4-induced M2a, and IL-10 or dexamethasone-induced by flow cytometry. Furthermore, IRF1 and CXCL10 gene expression was defined as specific for IFNγ/LPS-induced M1; IRF4-, CCL22-, and TGM2-specific transcripts for IL-4-derived M2a; and MERTK gene for dexamethasone-derived M2c, setting additional cardinal points to compare the skewing reprogramming of M-DM challenged with different stimuli or even co-cultured with another cell types.

All healthy volunteer blood donors provided written informed consent, and the study was approved by the Institutional Ethics Committee of the National Academy of Medicine (IMEX-CONICET-ANM) Argentina. See the Table of Materials for details about all materials and reagents used in this protocol.

1. Peripheral blood mononuclear cells (PBMCs) isolation

NOTE: PBMCs are isolated from 40 mL of anticoagulated peripheral blood with sodium citrate at 3.8% and using Ficoll-Hypaque density gradient (1077) centrifugation. This protocol is performed under sterile conditions in a BSL2 Biosafety cabinet.

1. Transfer the received anticoagulated peripheral blood sample into appropriate 15 mL or 50 mL tubes for centrifugation.
2. Centrifuge blood samples at 200 × g for 15 min (acceleration: 4 deceleration: 2, when the maximum is 5) at room temperature to obtain platelet-rich plasma (PRP) on top and cellular fraction, including red and white blood cells below. See Figure 1.
3. Transfer the upper fraction (PRP) to 15 mL tubes (if plasma will be stored), and centrifuge at 600 × g for 10 min (maximum acceleration and deceleration) to obtain the final plasma fraction without platelets. Store plasma at -20 °C.
4. Dilute the cellular fraction obtained from the first centrifugation, 2x (minimum) to 3x (maximum) using sterile 1x Phosphate-buffered Saline (1x PBS) prewarmed at room temperature. Gently homogenize it and use this diluted fraction for the Ficoll-Hypaque density gradient centrifugation.
5. Transfer 15 mL of Ficoll-Hypaque into a sterile 50 mL tube. Then, carefully and slowly add 30 mL of diluted blood on top of the Ficoll, avoiding mixing the phases. If working with 10 mL or less of diluted blood, transfer 4 mL of Ficoll-Hypaque into a 15 mL tube and then add the 10 mL of diluted blood sample.
   NOTE: This step is critical to obtain the mononuclear fraction.
6. Centrifuge the tube containing the Ficoll plus diluted blood at 600 × g for 25 min (acceleration: 1; Deceleration: 0) at room temperature.
7. Remove some of the yellow top layer with a sterile 3 mL Pasteur pipette. Then, carefully collect the interface (mononuclear "ring" fraction) formed between the yellow top layer and the Ficoll layer with a sterile Pasteur pipette. This "ring" contains the mononuclear cells (PBMC) of interest.
   NOTE: Some of the remaining plasma phase can be collected, but be sure to avoid the Ficoll phase, since it could be toxic for the PBMCs.
8. Transfer the collected PBMCs to a new 15 mL tube containing a minimum of 3 mL of sterile 1x PBS. Then, top up with the corresponding volume of sterile 1x PBS to 12-15 mL. Centrifuge the PBMCs at 600 × g (maximum acceleration and deceleration) for 10 min.
9. Discard the supernatant and wash the PBMCs again. Resuspend the cell pellet by adding 1 mL of sterile 1x PBS, then top up to 12-14 mL with additional sterile 1x PBS. Centrifuge the tubes at 600 × g (maximum acceleration and deceleration) for 10 min.
10. Discard the supernatant and resuspend the PBMC pellet in sterile 1x PBS containing 2% of inactivated fetal bovine serum (1x PBS-2% FBS) and bring the volume up to 10 mL. Use an automatic cell counter or count the cells, preparing a 1:5 cell suspension diluted in sterile cold 1x PBS and then (1:2) in Trypan Blue to obtain a final dilution of 1:10. Use a light microscope for counting.
11. Spin down the PBMC suspension at 300 × g for 5 min at 8-10 °C. Resuspend the cells in a desired volume of sterile, cold 1x PBS-2% FBS to obtain a final concentration of 10⁸ cells/mL. Leave these cells at 4 °C until the next step.
    NOTE: If PBMCs collected will not be used on the same day, it is recommended to freeze down in a cryogenic vial at 10⁸ cells/mL in appropriate freeze-down media (90% of FBS containing 10 % of dimethyl sulfoxide, DMSO). Store at -80 °C in a polyethylene glycol container overnight to allow the cells to freeze down slowly, thus minimizing cell damage. The next day, transfer the cells to a storage box and leave them at -80 °C for a couple of days or transfer to a liquid nitrogen tank for a long storage period.

2. CD14⁺ monocyte sorting using magnetic beads

NOTE: The amount of CD14 monocytes found in PBMCs of healthy donors is variable depending on age and sex. When available, use a cell counter to obtain the % of monocytes in each sample and then calculate the number of PBMCs needed to sort the required CD14 monocytes. If no cell counter is available, 10% of monocytes per PBMC sample could be considered a wide approach^{4,6,7,10,11,12,13}.

CD14⁺ monocytes are isolated using a human CD14 positive selection kit⁵. We have standardized the isolation protocol using 1 to 2 × 10⁷ PBMCs in 100 µL of reaction; use this ratio for higher numbers. The minimum reaction volume suggested is 100 µL, even with a lower cell number than 1 × 10⁷ PBMCs.

1. Prepare PBMC cell suspension at 1-2 × 10⁸ cells/mL concentration in 1x PBS-2% FBS containing 1 mM of ethylenediaminetetraacetic acid (1x PBS-2% FBS-1 mM EDTA) buffer and transfer the desired amount of PBMCs into a 5 mL Polystyrene Round-bottom tube.
2. Add 10 µL of Positive Selection Cocktail for every 100 µL of cell suspension, mix well, and incubate at room temperature for 15 min.
3. Add 10 µL of magnetic nanoparticles per 100 µL of cell suspension, vigorously pipette up and down more than 5x to uniformly suspend the magnetic nanoparticles, and incubate at room temperature for 10 min.
   NOTE: Vortexing is not recommended.
4. Add 1x PBS-2% FBS-1 mM EDTA buffer to bring up the total volume of the cell suspension to 2.5 mL. Gently pipette the cells in the tube up and down 2-3x. Insert the tube into the magnet sorter and set it apart for 5 min.
5. Take the magnet and invert it and the tube in one continuous motion, discarding the supernatant fraction. The magnetically labeled cells will stay inside the tube. Leave the magnet and tube inverted for 2-3 s, then turn them vertically upright.
   NOTE: Leave any hanging drops from the tube's mouth; do not shake or blot them.
6. Remove the tube from the magnet and add 2.5 mL of 1x PBS-2% FBS-1 mM EDTA buffer into the tube again. Gently pipette the cell suspension up and down 2-3x to mix it. Insert the tube back into the magnet and set it apart for 5 min.
7. In one continuous motion, invert both the magnet and tube, discarding the supernatant fraction. Remove the tube from the magnet and resuspend the cells in a suitable amount of 1x PBS-2% FBS buffer (without EDTA). The positively selected cells are now ready to use.
   NOTE: A total of two 5 min separations are required in the magnet.
8. Wash sorted CD14 cells employing 1x PBS-2% FBS buffer, bringing the cell suspension to 12-14 mL to dilute any remaining EDTA, and then centrifuge at 600 × g (maximum acceleration and deceleration) for 10 min.
9. Discard the supernatant and repeat the washing by resuspending the cell pellet with 1 mL of 1x PBS-2% FBS buffer first and then topping up the volume to 12-14 mL. Centrifuge the tubes at 600 × g (maximum acceleration and deceleration) for 10 min.
10. Discard the supernatant and resuspend the positive selected CD14 monocytes in 2 mL of 1x PBS-2% FBS buffer. Use the automatic cell counter or count the cells by diluting the cell suspension (1:5) in sterile cold 1x PBS and then (1:2) in Trypan Blue using a light microscope. Leave the cells on ice until culture preparation.
    NOTE: Cell purity can be tested by flow cytometry, taking 50 µL of the cell suspension and staining it for CD14⁺ cells (clone HCD14). More than 90% purity is recommended.
11. For culturing, spin down the cell suspension at 300 × g for 5 min at 8-10 °C. Resuspend the pellet at 10⁶ cells/mL concentration using RPMI 1640 supplemented with 10% inactivated FBS and 1% penicillin-streptomycin (RPMI-10% FBS-1% P/S).
    NOTE: It is critical to use a high-quality, low-endotoxin, heat-inactivated FBS. Maintain always on ice to avoid monocyte adherence to the plastic wall before plating.

3. Monocyte-derived macrophage (M-DM) differentiation and polarization

NOTE: M-DM culture is performed by plating 2.5 × 10⁵ CD14⁺ monocytes in 48-well plates containing 500 µL of RPMI-10% FBS-1% P/S and cultured in a humidified incubator at 37 °C with CO₂ (5%) for 7 days. Choose treated plates to enhance monocyte adherence to the plate.

1. Define all experimental conditions to run, including wells for basal M-DM (M0), the inflammatory M1, the tissue repair M2a, the regulatory M2c, and the desired testing conditions.
2. Pipette 250 µL of the cell suspension prepared in step 2.11 to seed 2.5 × 10⁵ CD14⁺ monocytes in each well.
3. Add 250 µL of RPMI-10% FBS-1% P/S plus 50 ng/mL of M-CSF to each well to complete the 500 µL of total volume.
4. Incubate the cell culture plate in a humidified incubator at 37 °C with 5% CO₂ for 4 days to allow the first step of macrophage differentiation.
   NOTE: Periodically check the cell culture differentiation to see if morphological changes occurred as expected or if undesired unattached cells exist.
5. Replace half of the culture medium (250 µL of RPMI-10% FBS-1% P/S) on day 4 of the M-DM culture. Add cytokines or reagents for the polarizing conditions and culture them for 3 days (the endpoint is day 7): LPS (1 ng/mL) plus IFNγ(50 ng/mL) for M1, IL-4 (40 ng/mL) for M2a, IL-10 (50 ng/mL) or dexamethasone (0.1 µM) for M2c.
6. Harvest M-DM on day 7 to perform phenotypic characterization, collect them for mRNA expression as indicated in the next two steps, or perform a functional assay.

4. Surface phenotype characterization in polarized M-DM by flow cytometry

1. Discard the total culture medium volume (500 µL of a 48-well plate) and add 200 µL of 1x PBS-2% FBS-1 mM EDTA buffer for cell harvesting. Incubate for 20 min on ice and then add 200 µL of 1x PBS-2% FBS to dilute the EDTA buffer. Collect the M-DM by pipetting up and down in 1.5 mL microtubes. Wash these harvested cells once with 1x PBS-2% FBS and centrifuge at 800 × g for 3 min.
2. Resuspend the M-DM pellet with 200 µL of 1x PBS-5% FBS and transfer them to a 96-well round-bottom microplate for blocking at room temperature for 15 min. Centrifuge the cells at 800 × g for 3 min.
3. Prepare the appropriate combination of directly conjugated antibodies in 1x PBS- 2% FBS. Stain each well or condition (1x) with 50 µL of the antibody panel and incubate at 4 °C for 30 min in the dark.
   NOTE: Here, the antibody panel consisted of CD11b-APC/Cy7, CD64-APC, CD163-PerCP/Cy5.5, CD206-AlexaFluor 488, CD14-PECy7, and MERTK-biotin plus Streptavidin-PE. Pretitered antibody concentration is listed in Table 1.
4. Wash the stained M-DM by adding 150 µL of 1x PBS and centrifugate at 800 × g for 3 min.
5. To measure cell viability, add 100 µL of the fixable viability dye Zombie Violet (1:1,000 in 1x PBS) and incubate at 4 °C for 30 min in the dark.
6. Wash the stained cells by adding 100 µL of 1x PBS and centrifuge at 800 × g for 3 min. Repeat the washing procedure using 200 µL of 1x PBS.
7. Fix by resuspending the M-DM pellet with 100 µL of the fixative reagent (from the fixative/permeabilization kit) and incubating them for 20 min on ice and in the dark.
   NOTE: To achieve appropriate fixation, it is essential to homogenize the cell pellet correctly.
8. Wash the M-DM by adding 100 µL of 1x PBS-2% FBS and centrifuge at 800 × g for 3 min. Repeat the washing procedure using 200 µL of 1x PBS-2% FBS.
9. Resuspend the M-DM pellet in 200 µL of 1x PBS and run the stained cells in the flow cytometer.

5. Gene program profile to discriminate M-DM polarization by qPCR

NOTE: On day 7 of the culture, 2.5 × 10⁵ M-DM can be harvested using the RNA extraction reagent for the isolation of high-quality total RNA. RNA isolation can also be performed using alternative methods, such as column-based RNA extraction and an elution kit.

1. Discard the total culture medium volume and lysed M-DM by directly adding 350 µL of the extraction reagent. Homogenize and transfer this cell suspension into a 1.5 mL RNase-/DNase-free microtube and continue with the RNA isolation.
   NOTE: If RNA isolation will not be performed on the same day, the sample from step 5.1 can be stored at 4 °C overnight or -20 °C for up to a year.
2. To allow complete dissociation of the nucleoproteins complex, incubate the homogenized samples from step 5.1 for 5 min at 25 °C
3. Add 70 µL of chloroform to each sample. Securely cap each tube to prevent leakage and thoroughly mix the contents by shaking the tube. Incubate the samples for 2-3 min and then, centrifuge the samples at 12,000 × g for 15 min at 4 °C. This step will fully separate the phases into a lower phenol-chloroform phase, an interphase, and a colorless upper aqueous phase.
   NOTE: The volume of chloroform is based on a ratio of 0.2 mL of chloroform per 1 mL of extraction reagent used during the lysis step.
4. Transfer the aqueous phase containing the RNA to a new 1.5 mL RNase-/DNase-free microtube.
   NOTE: Leave behind a small volume when taking the aqueous phase to avoid transferring any of the interphase or organic layers.
5. Add 175 µL of isopropanol to each sample to precipitate the RNA. Incubate for 10 min at 4 °C or alternatively overnight at -20 °C.
   NOTE: The volume of isopropanol is based on a ratio of 0.5 mL of isopropanol per mL of initial extraction reagent.
6. Centrifuge the samples at 12,000 × g for 10 min at 4 °C. A white gel-like pellet of total RNA will precipitate at the bottom of the tube. Carefully remove and discard the supernatant using a micropipette.
7. Wash the RNA by resuspending the white gel-like pellet in 350 µL of 75% ethanol (1 mL per 1 mL of extraction reagent (v/v) used for lysis).
   NOTE: The RNA can be stored in 75% ethanol for up to 1 year at -20 °C or up to 1 week at 4 °C.
8. Briefly vortex the resuspended pellet and centrifuge at 7,500 × g for 5 min at 4 °C. Remove and discard the supernatant using a micropipette. Air-dry or vacuum the RNA pellet for 5-10 min.
   NOTE: To solubilize the RNA, avoid overdrying of the RNA pellet.
9. Resuspend the pellet in 20-30 µL of RNase-free water by pipetting up and down. Incubate in a water bath or heat block set at 55-60 °C for 10-15 min. Proceed to measure RNA concentration and store the RNA at -70 °C.
10. Perform the reverse transcription (cDNA copy) using 100-500 ng of RNA (minimal and maximal, respectively) in 20 µL of reaction volume by employing a cDNA synthesis kit.
11. Run the real-time PCR reactions using 1 µL of cDNA (5-25 ng).
    1. Prepare the reaction mix for each tube (1x) using 5 µL of SYBR green mixture (2x), 0.2 µL of each primer (forward and reverse), and 3.6 µL of nuclease-free water. Scale all components according to sample number. Mix the reaction mix thoroughly and dispense 9 µL into each qPCR tube and finally the 1 µL of cDNA to reach the final total volume of 10 µL.
    2. Program the thermal cycling protocol on a real-time PCR instrument as follows: 2-3 min at 98 °C (1x); 5-15 s at 98 °C plus 15-30 s at 60 °C (35-40x); finally include melt curve analysis (65-95 °C) with 0.5 °C increment at 2-5 s/step or use the instrument's default setting.
       NOTE: This reaction was standardized employing the universal SYBR Green mix. Primers recommended for M-DM polarization analysis are listed in Table 2.
12. Normalize the results using as the reference gene the eukaryotic translation elongation factor 1 alpha 1 (EEF1A1), and check the specificity of the amplified products through the analysis of the dissociation curves.
    NOTE: The dissociation curve analysis allows confirming the presence of only one qPCR amplification product in the reaction.

Based on our work in macrophage characterization for several years, we have set an accurate combination of markers that clearly distinguishes the different subsets of in-vitro M-DM. The phenotype markers were selected based on the literature and a wide previous screening that we had previously performed^{9,14,15,16,17}. Furthermore, we determined that CD64, CD206, CD163, CD14, and MERTK can help clearly discriminate between unpolarized M-DM (M0), polarized IFNγ/LPS-induced M1, IL-4-induced M2a, and IL-10- or dexamethasone-induced M2c by flow cytometry^4,5,7,11,12. The description and function of each selected marker can be found in Table 2⁴.

The panel of markers analyzed by flow cytometry for in vitro analysis of M-DM polarization is shown in Figure 2. Comparing the different polarized macrophages, we see that M1, induced by IFNγ plus LPS, is characterized by the highest level of CD64, the absence of CD206 expression, and low levels of CD163 and MERTK. On the other hand, M2a macrophages induced by IL-4 are specifically characterized by an increase in CD206 expression and a reduction in CD64, CD163, and MERTK. Finally, the M2c phenotype induced by IL-10 or dexamethasone is characterized by a specific increase in CD163 expression, intermediate levels of CD64, and an increase in MERTK and CD14. In addition, there is a marked reduction of CD206 only when polarized with IL-10. The dexamethasone-induced M2c interestingly also increases level of CD206. All these markers were analyzed after gating of CD11b⁺ viable cells. In addition to the surface phenotyping, we also set a specific gene expression panel for the classical IFNγ/LPS-induced M1 (IRF1 and CXCL10) and IL-4-induced M2a (IRF4, CCL22, and TGM2). The dexamethasone-M2c transcriptional-induced program is characterized by MERTK expression.

Figure 1: Peripheral blood mononuclear cells isolation by Ficoll gradient. After platelet-rich plasma is discarded, the diluted peripheral blood sample (2-3x) is loaded on top of Ficoll-Hypaque density gradient and centrifuged at 600 × g for 25 min (Acceleration: 1; Deceleration: 0), at room temperature to obtain the PBMCs as indicated. Carefully collect the interface formed between the yellow top layer and the Ficoll layer with a sterile Pasteur pipette. This "ring" contains the mononuclear cells of interest. Abbreviations: PBMCs = peripheral blood mononuclear cells; RBC = red blood cell. Please click here to view a larger version of this figure.

Figure 2: Phenotypic characterization of polarized M-DM by flow cytometry. Monocyte-derived macrophages (M-DM) were in-vitro differentiated for 7 days in RPMI 1640 medium containing 10% FBS and 1% P/S plus 50 ng/mL of M-CSF. On day 4, half of the culture medium was replaced, and polarizing cytokines were added. Unpolarizing M-DM (M0) received only medium; LPS (1 ng/mL) plus IFNγ (50 ng/mL) was added for M1; IL-4 (40 ng/mL) was added for M2a; IL-10 (50 ng/mL), or dexamethasone (0.1 µM) were added for M2c. M-DM was cultured for an additional 3 days. On day 7, M-DM was harvested to perform phenotypic characterization by flow cytometry using a panel of five M-DM markers, (A) CD64, CD206, CD163, CD14, and (B) MERTK. The gating strategy also included viable cells and CD11b⁺ cells. Representative dot plots show the expression levels of each polarizing marker comparatively for each type of M-DM (M0, M1, M2a, and M2c). Please click here to view a larger version of this figure.

Table 1: Antibodies panel used for M-DM immune phenotyping. Please click here to download this Table.

Table 2: A brief description and function of each selected marker to discriminate in-vitro polarized monocyte-derived macrophage used in flow cytometry and qPCR. Please click here to download this Table.

Macrophage differentiation, activation, and polarization have become a central focus in immunology, tissue homeostasis, disease pathogenesis, and inflammation resolution. Additionally, the emerging picture that tissue macrophages may be derived from circulating monocytes under conditions of disrupted homeostasis emphasizes the necessity for precise M-DM models characterized by defined phenotypes, transcriptional programs, and activation pathways.

Some critical steps in this protocol must be discussed here for its successful implementation. First, it should be noted that blood samples from which primary M-DM cultures are established may be heterogeneous regarding age, sex, genetic background, or environmental variables which may generate considerable variation in some biological responses. Hence, it is critical to have basal biological and experimental controls from the same donor. A paired analysis is recommended. Nonetheless, the high variation could also be considered an advantage since the response spectrum better reflects the biological significance. Single-cell suspensions should have more than 70% viable cell populations before sorting. Similarly, the postsorting purity of CD14 monocytes should yield 90%. Another critical aspect of M-DM culture is the initial cell density. Low density does not allow appropriate cellular cooperation and macrophage differentiation, and high density increases cell death. Although the influence of cellular interactions in macrophage reprogramming has not been explored extensively, some pioneer studies demonstrated that co-culturing human monocytes with regulatory T cells skews the balance toward differentiation of M2 macrophages¹⁸. Similarly, apoptotic cells are key players in acquiring an M2 phenotype when exposed to interleukin-13 (IL-13) and IL-4¹⁹. We have also demonstrated that platelets reprogrammed monocytes toward the M1 pro-inflammatory phenotype in the presence of LPS and a cell-contact-dependent manner (GPIb-CD11b partnership)⁵.

Another essential prerequisite is the inclusion of appropriate controls for flow cytometry experiments. Autofluorescence and compensation controls are critical for the equipment set, but Fluorescence minus one (FMO) should be mandatory to establish thresholds for negative and positive signals. Although the in-vitro models may not fully recapitulate the specific nature of the tissue, numerous studies have shown that macrophages can acquire tissue-specific identity through phagocytic and inflammatory pathways^3,20,21. We have reported not only the re-programming effect of a drug (ketamine⁴) and a lipid mediator (C1P⁷) on macrophage differentiation and polarization but also developed a heterotypic tridimensional model to study the interaction of macrophages and glioblastoma in vitro¹².

This protocol has some limitations. A primary concern is that using circulating blood monocytes as a source of macrophages may not fully replicate the characteristics of tissue-resident macrophages, which often originate from distinct embryonic lineages. Furthermore, the inherent heterogeneity of human blood samples, influenced by factors such as age and sex, can introduce significant variability in macrophage responses.

Despite these limitations, the protocol offers several key advantages. It is relatively easy to establish with readily available reagents and yields a high number of primary macrophages, enabling a broad range of experimental conditions for proof-of-concept studies. The resulting high purity of these cultures allows for precise evaluation of specific macrophage responses. Importantly, these are primary cells, not immortalized or established cell lines, which often exhibit altered characteristics.

The protocol has numerous applications, including investigating the effects of drugs on macrophage differentiation and polarization, studying interactions with other cell types in co-culture, and analyzing the impact of conditioned media from various cultures or plasma from patients with different pathologies. While the current protocol allows for some co-culture interactions, the development of organoid platforms promises to readily address limitations related to three-dimensional culture in the near future.

The authors have no conflicts of interest to disclose.

This work was supported by the National Agency for the Promotion of Science and Technology (ANPCyT-FONCYT) through grants PICT 2018-3070 and 2021-I-A-00807 to E.A.C.S. and PICT 2021-I-A-00716 to A.E.E, and by the National Scientific and Technical Research Council (CONICET) through the grant PIP 2022-0763 and by the University of Buenos Aires through Proyectos de Investigación y Desarrollo en Áreas Estratégicas con Impacto Social (PIDAE) 2022 to A.E.E. E.A.C.S. and A.E.E. are career investigators at CONICET.