Modeling Tumor-Associated Macrophages In Vitro: The Role of Secreted Factors in Tumor Microenvironment Dynamics

Summary

This manuscript presents an in vitro protocol for generating tumor-associated macrophage-like cells by exposing human monocytes to conditioned media from tumor cell lines.

Abstract

Macrophages exhibit remarkable plasticity, enabling them to acquire distinct functional states in response to a variety of environmental cues. In solid tumors, they frequently constitute the most abundant immune cell population within the tumor microenvironment, where they are referred to as tumor-associated macrophages (TAMs). TAMs closely interact with malignant cells, actively promoting tumor progression, angiogenesis, and immune evasion. Tumor cells can reprogram macrophages through direct cell-cell contact and a wide array of secreted factors, shaping their polarization toward pro-tumoral phenotypes. To study these TAM polarization mechanisms in vitro, we propose a method that exposes human monocytes to soluble factors present in tumor cell line-conditioned media. This protocol enables the generation of TAM-like macrophages under controlled conditions, facilitating the analysis of phenotypic and functional changes induced by tumor-derived signals. Our approach offers a reproducible and physiologically relevant model to investigate tumor-macrophage interactions, deepens our understanding of macrophage reprogramming, and supports research into novel therapeutic strategies in cancer immunology and immunotherapy.

Introduction

Macrophages are multifunctional innate immune cells whose remarkable plasticity enables them to adopt diverse phenotypes in response to environmental cues¹. Within solid tumors, macrophages often become the most abundant immune cell population and differentiate into tumor-associated macrophages (TAMs)². These cells are actively reprogrammed by the tumor microenvironment to support tumor growth, promote angiogenesis, facilitate metastasis, and suppress effective antitumor immune responses. Tumor cells drive this polarization through both direct cell-cell interactions and a complex repertoire of secreted factors, including cytokines, chemokines, and growth factors, which reshape macrophage transcriptional and functional programs³. This dynamic interplay underpins the central role of TAMs in cancer biology and highlights macrophage plasticity as both a challenge and an opportunity for therapeutic intervention².

The clinical relevance of TAMs is increasingly recognized: their abundance in tumors often correlates with poor prognosis and resistance to therapies^4,5. Consequently, TAMs⁶ have emerged as attractive therapeutic targets. Strategies aimed at blocking their recruitment, inhibiting their survival, or reprogramming them toward inflammatory and tumoricidal phenotypes are currently under investigation⁶. Understanding the mechanisms by which tumor-derived factors polarize macrophages into TAMs is thus essential for developing effective immunotherapies. While immune checkpoint blockade has shown great success in many cancers, its efficacy remains limited in others⁷. Combining T cell-targeted therapies, such as anti-Programmed Cell Death Protein 1 (PD-1) or anti-Programmed Cell Death Ligand 1 (PDL-1) antibodies, with macrophage reprogramming strategies may enhance the overall antitumor immune response.

Traditional experimental models to study TAM biology have included in vivo murine cancer models, which offer a complex and physiologically relevant environment⁸. These models have provided fundamental insights into TAM recruitment and function. However, significant interspecies differences in immune signaling, tumor biology, and macrophage ontogeny can limit direct translation to human disease^9,10. Moreover, the complexity of in vivo systems makes it challenging to isolate and study specific signaling pathways or molecular events that drive macrophage polarization.

In contrast, in vitro models offer simpler and controlled conditions to dissect these mechanisms. The most widely used approach involves polarizing human monocyte-derived macrophages with defined stimuli: Interferon gamma (IFN) and bacterial lipopolysaccharide (LPS) to generate pro-inflammatory macrophages (M1), or Interleukin 4 (IL-4) and interleukin 10 (IL-10), or dexamethasone to obtain various anti-inflammatory macrophage (M2) subtypes^11,12,13,14. These models have helped identify molecular markers and transcriptional programs associated with canonical macrophage polarization states. Nonetheless, they fall short of replicating the complex, multifactorial environment that macrophages encounter within tumors, where polarization results from the simultaneous influence of numerous soluble mediators, metabolic signals, and cell-to-cell interactions^15,16. Although the classical M1/M2 classification remains useful to describe macrophage functional states, it is now clear that many more macrophage populations exist than previously thought. The work of Dr. Ginhoux's group and others, using single-cell transcriptomic approaches, has demonstrated the broad diversity of macrophage subsets found both under physiological conditions and in disease^17,18. This highlights that TAMs, although often described as resembling M2-like profiles, are not strictly equivalent to macrophages differentiated solely with canonical cytokines; rather, they share certain features but also display distinct and context-dependent differences.

To bridge this gap, we present an in vitro protocol that models TAM-like polarization by exposing human monocyte-derived macrophages to conditioned media collected from tumor cell lines. This method captures the influence of the tumor secretome, a rich mixture of cytokines, chemokines, and growth factors secreted by cancer cells. It thus offers a relevant and experimentally accessible system to investigate the phenotypic and transcriptional reprogramming of macrophages in response to tumor-derived soluble factors. Unlike previous approaches in which monocytes were directly exposed to tumor-conditioned media, this protocol specifically focuses on the reprogramming of macrophages, thereby better reflecting resident macrophage behavior within the tumor microenvironment^19,20. In addition, it incorporates an optimized and straightforward panel of surface markers to assess macrophage polarization by flow cytometry, which is easy to implement and can be expanded according to the investigator's needs^14,21. Finally, since M-CSF has been shown to prime monocytes toward an M2-like phenotype^{10,15,22,23,24}, we deliberately omitted it to ensure that any observed macrophage polarization results specifically from exposure to tumor cell line-derived conditioned media.

This approach has several strengths: it retains the reproducibility and simplicity of in vitro systems, uses primary human cells for greater translational relevance, and reflects the heterogeneous, multifactorial nature of the tumor microenvironment better than single-cytokine models^25,26,27,28. Moreover, it facilitates comparative studies of macrophage responses to conditioned media from different tumor types or genetically modified cancer cells, and serves as a platform to test pharmacological agents that may alter TAM polarization. Beyond its general applicability, this method can be tailored to different tumor models by selecting appropriate cell lines or adjusting the proportion of conditioned media. Nonetheless, users should be aware that variability across healthy donors and tumor cell lines may affect reproducibility, underscoring the importance of careful experimental design and standardization.

Protocol

Peripheral blood samples were collected from healthy donors after informed consent in accordance with the Declaration of Helsinki. This protocol was approved by the Institutional Ethics Committee of the University of Buenos Aires. See the Table of Materials for details about all materials and reagents used in this protocol.

NOTE: Maintain sterile conditions throughout the whole protocol, working in a Biosafety level 2 (BSL-2) cabinet, and ensure cell viability >90%. For disposal, all biological waste must first be decontaminated by soaking in a 10% sodium hypochlorite solution for at least 30 min, and then placed in the appropriate biohazard disposal container.

1. Tumor cell line culture and conditioned media preparation

1. Thaw the MDA-MB-231 triple-negative human breast cancer (or other selected tumor) cell line from liquid nitrogen storage. Immediately immerse cryotubes containing cell lines in a 37 °C water bath and gently swirl the vial to ensure rapid and uniform thawing. Remove the vial promptly when mostly thawed.
2. Transfer the thawed cells into a 15 mL centrifugation tube containing 5 mL of sterile Phosphate-buffered Saline (PBS). Centrifuge the cell suspension at 400 × g for 5 min at room temperature (RT).
   NOTE: The thawing process must be performed quickly to preserve cell viability.
3. Carefully discard the supernatant and quickly resuspend the cell pellet in 1 mL of fresh complete Dulbecco's Modified Eagle Medium (DMEM) containing high glucose (4.5 g/L) with 10% fetal bovine serum (FBS), 4 mM Glutamine, and 1 mM sodium pyruvate. Count the cells in the resuspended pellet, performing the following two-step serial dilution:
   1. First prepare a 1:5 cell suspension dilution in sterile DMEM medium and then (1:2) with the viability stain Trypan Blue to obtain a final dilution of 1:10.
   2. Use a light microscope to count the viable cells within the Neubauer hemocytometer (or counting chamber). Count viable cells (only cells that exclude the blue dye) and determine the average number of viable cells per large corner square.
   3. Calculate the final concentration using the following formula: Cells/mL = (Average cells per square) x (Dilution Factor) x 10,000.
      NOTE: Resuspend the cells in a desired volume of complete DMEM medium to obtain a final concentration of 1x 10⁵cells/mL.
4. Seed 2.5 x 10⁵ to 5.0 x 10⁵ cells into a T25 culture flask containing 5 mL of complete DMEM medium. Maintain the culture at 37 °C in a humidified incubator with 5% CO₂ until the required cell density for harvesting is reached.
5. The next day after seeding, check cell attachment under a microscope. If more than 50 % of the cells are floating (indicating cell death), gently aspirate the old medium and replace it with 2.5 mL of fresh, pre-warmed medium.
6. Monitor cell growth daily. When the culture reaches approximately 90% confluency (an estimated 2.3 x 10⁶ cells in a T25 flask), typically within 5 days, the cells are ready to be subcultured (split). To split the cells at a 1:8 ratio, follow these steps:
   1. Remove the spent medium using a sterile Pasteur pipette.
   2. Rinse the cell monolayer once with 3 mL of sterile 1x PBS to remove residual serum, which inhibits trypsin. Aspirate and discard 1x PBS.
   3. Add to the cells 1 mL of 0.25% Trypsin- 1 mM Ethylenediaminetetraacetic acid (EDTA) solution and incubate at 37 °C, 5% CO₂ on a humidified incubator for 3 min to detach the cells.
      NOTE: The optimal incubation time with Trypsin/ EDTA is dependent on the specific cell line and may need to be adjusted. Check the cells under the microscope after 3 min to ensure they are fully detached.
   4. Neutralize the enzymatic activity by adding 3 mL of complete DMEM medium. Immediately after, gently pipette up and down to fully detach and suspend the cells.
7. Transfer 3.5 mL of the cell suspension into a 15 mL conical tube and centrifuge it for 5 min at 400 x g at RT.
8. Carefully remove the supernatant and resuspend the cell pellet in 1 mL of freezing medium (which must contain 90% FBS plus 10% dimethyl sulfoxide (DMSO). Transfer and aliquot the cell suspension into cryovials, ensuring a concentration of 5 x 10⁵ cells per tube for future use.
   1. Freezing needs to be gradually cooled by placing the cryovials into a Mr. Frosty (or similar controlled-rate freezing container). Transfer the container to a -80°C freezer and leave the cells to freeze overnight. Cells can be kept at -80°C for a few days, but should then be moved to liquid nitrogen for long-term storage.
      NOTE: Keep detailed records of cell passages and use early passages for experiments. Using cells with less than 20 passages is recommended.
9. Once new cultures again reach ~90% confluency, aspirate and collect supernatant into a 15 mL conical tube. If you wish to keep the culture ongoing, replace the collected volume with fresh complete DMEM medium.
   1. Centrifuge the collected supernatant at 400 × g for 5 min at 4 °C to pellet and remove cellular debris. Keep the supernatant (the clarified conditioned media), and discard the pellet.
10. Aliquot the clarified conditioned media into 1 mL portions, label the tubes, and store them at -20 °C until use.
    NOTE: For quality control, always keep a record of the conditioned media collection date and whether the aliquot has been previously thawed. It is strongly advised to avoid thawing the same aliquot more than twice. Furthermore, use the conditioned within one month of the freezing date to ensure its optimal activity.

2. Isolation of peripheral blood mononuclear cells (PBMCs)

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

1. Aliquot the anticoagulated peripheral blood sample into either a 15 mL or 50 mL conical tube before the upcoming centrifugation step.
2. Centrifuge blood samples at low speed, 200 × g for 15 min at RT to separate the blood components. This process yields platelet-rich plasma (PRP) as the top layer and the cellular fraction (red and white blood cells) below. Use a centrifuge setting of acceleration: 4 and deceleration: 2, when the maximum is 5.
3. After centrifugation aspirate the supernatant (the upper fraction), which is the PRP, and transfer it to a new sterile tube.
   NOTE: If needed for downstream experiments, the collected PRP can be centrifuged again at 600 × g at RT for 10 min (maximum acceleration and deceleration) to remove the platelets, and store the plasma without platelets at -20 °C for other assays.
4. The cellular lower fraction obtained from the initial centrifugation in step 2 should be gently diluted to a 2- to 3-fold final volume using sterile, room-temperature 1x PBS. Homogenize softly by inversion the resulting suspension and use it for the Ficoll-Hypaque density gradient centrifugation.
5. Dispense 15 mL of room-temperature Ficoll-Hypaque into a sterile 50 mL tube.
6. Carefully and slowly layer 30 mL of diluted blood over the Ficoll-Hypaque, ensuring the two phases do not mix.
7. Centrifuge the Ficoll-blood layered tube for 25 min at 600 × g at RT. Set the centrifuge without brake (acceleration: 1; Deceleration: 0).
8. Use a sterile Pasteur pipette to discard at least 4 mL of the obtained top fraction to avoid spilling. Then use the same pipette to recover the white ring containing the PBMCs that is formed between the upper fraction and the Ficoll-Hypaque transparent fraction. Avoid collecting Ficoll with the PBMC ring.
9. Transfer the PBMC ring from the tube to a sterile 15 mL centrifuge tube with at least 3 mL of 1x PBS.
10. Fill the 15 mL tube with more 1x PBS and centrifuge at 600 × g (maximum acceleration and deceleration) for 10 min at RT.
11. Remove the supernatant by inversion, fill again with 1x PBS, and centrifuge at 600 × g (maximum acceleration and deceleration) for 10 min at RT.
12. Remove the supernatant and resuspend the PBMC pellet in 2 to 3 mL of sterile 1x PBS plus 2% heat-inactivated FBS (1x PBS-2% FBS), then bring the volume up to 10 mL. Count the cells, preparing a 1:5 cell suspension diluted in sterile cold 1x PBS and then (1:2) in 0.4% Trypan Blue to obtain a final dilution of 1:10. Use a cell chamber on a light microscope as described in step 1.3.2 for counting.
13. 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: At this point, PBMCs collected can be frozen down in a cryogenic vial at 10⁸ cells/mL in freeze-down media (90% FBS-10 % DMSO). Store at -80 °C in a Mr. Frosty container overnight to allow the cells to freeze down slowly. 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 or continue the protocol.

3. CD14+ monocyte sorting using magnetic beads

NOTE: Purified CD14+ monocytes are obtained from PBMC using a human CD14+ positive selection magnetic sorting kit¹¹. CD14+ cell percentage in PBMC is donor-dependent but can be estimated at 10% (ranging from 5-20%). Adapt the initial number of PBMC and the final reaction volumes based on the required number of monocytes for your experiment. The suggested minimum reaction volume is 100 µL, even when processing fewer than 1 × 10⁷ PBMC.

1. Add 10 µL of the positive selection cocktail for each 100 µL of PBMCs, pipette up and down, and keep on ice for 15 min.
2. Vortex the beads suspension for 30 s prior to use. Then add 10 µL of the beads per 100 µL of PBMC suspension. Mix well and incubate for 15 min on ice.
3. Add 1x PBS- 2% FBS -1 mM EDTA to a final volume of 2.5 mL, gently mix the suspension and insert the tube on the magnet and incubate for 5 min.
4. Invert the magnet (without removing the tube) over a discard recipient tube for 2-3 s, avoiding any shaking movement. Positive cells will stay in the tube.
5. Take the tube out of the magnet and add 1x PBS- 2% FBS -1 mM EDTA over the tube walls until a final volume of 2.5 mL.
6. Insert the tube in the magnet again and incubate for 5 min.
7. Invert the magnet (without removing the tube) over a discard recipient tube for 2-3 s, avoiding any shaking movement. Retrieve the tube from the magnet and add 3 mL of 1x PBS-2% FBS over the tube walls to recover all the cells.
8. Take 10 µL to count the number of CD14+ cells in a Neubauer chamber with a light microscope. Dilute with Trypan Blue 1:2.
   NOTE: Calculate the number of cells following this equation: Number of CD14+ cells = media number of live cells per quadrant x dilution factor x 10.000.
9. Place the CD14+ cell suspension in a new sterile 15 mL tube and wash cells by centrifuging at 600 × g (maximum acceleration and deceleration) for 10 min at 4 °C.
10. Resuspend CD14+ cells with RPMI 1640 medium supplemented with 10% FBS and 1% penicillin-streptomycin (P/S) complete medium at a convenient volume to obtain 2.5 x 10⁶ cells/mL for the next step, considering the number of cells obtained.
    NOTE: Maintain the CD14+ monocytes on ice until the next step to avoid attachment of these cells to the plastic tube wall.

4. Differentiation of macrophages

1. Plate purified CD14+ monocytes at 2.5 x 10⁵ cells suspended in 250 µL of complete RPMI medium (10% FBS-1% PS) per well in a 48-well treated plate.
   NOTE: It can also be done with half the number of cells in a 96-well treated plate.
2. Add sterile water to the surrounding empty wells to reduce evaporation during plate incubation.
3. Incubate cells for 5 days at 37 °C with 5% CO₂ in a humidified incubator without disturbing, monitoring daily for morphology, differentiation, and viability.
   NOTE: While M-CSF (25 ng/mL) can be added for differentiation, we confirm that our protocol uses 10% high-quality FBS alone to obtain M0 macrophages after five days of culture. We specifically recommend avoiding M-CSF due to its potential to prime monocytes toward an M2-like phenotype^10,22.

5. Exposure of macrophages to conditioned media and analysis

1. On day 5 of differentiation, gently remove 50% (125 µL) of the medium from each well and replace with either:
   1. 125 µL of fresh RPMI complete medium (control),
   2. 125 µL of tumor cell line-conditioned media (e.g., from MDA-MB-231), or
   3. 125 µL of DMEM complete medium (control for conditioned media).
      NOTE: For initial optimization with new tumor cell lines, test a range of conditioned media concentrations (e.g., 10%, 25%, and 50%). This range is suggested to ensure macrophage viability and differentiation changes. Choose the higher concentration of conditioned media that shows high macrophage viability.
2. Continue incubation until day 7.
   NOTE: On day 5, M0 macrophages are obtained, then 2 more days of culture in the presence of conditioned media is enough to induce the expression of surface polarization markers. Polarization surface markers are determined by flow cytometry, similarly, when using standardized cytokines to generate M1 or M2 macrophages^11,14.
3. Assess macrophage viability and morphology under the microscope daily.
   NOTE: Monitor macrophage differentiation daily over the 7-day culture period. Cell adherence is used as an indicator of a healthy, viable culture. A higher proportion of fusiform cells is indicative of increased differentiation and polarization in the conditioned media (CM) treatment group, as shown in Figure 2.

6. Harvest macrophages for evaluation of polarization markers by flow cytometry

NOTE: Optionally, to further extend the phenotype characterization of tumor-associated macrophages, isolate RNA to measure the expression of polarization-associated genes (e.g., TGM2, IRF-4, and CXCL10) by quantitative polymerase chain reaction (qPCR) or collect the culture supernatants for measurement of canonical cytokine production (e.g., TNFα, IL-6, and IL-10) by enzyme-linked immunosorbent assay (ELISA) as reported by our group^11,14.

1. Discard by gently aspirating with a sterile Pasteur pipet or tip the total culture medium volume of each well and add 200 µL of cold 1x PBS-2% FBS-1 mM EDTA buffer for cell harvesting.
2. Incubate for 10 to 15 min on ice and then add 200 µL of 1x PBS-2% FBS to dilute the EDTA buffer. Collect the macrophages by pipetting up and down (still on ice) for 5 additional min until all cells get detached.
   NOTE: This step is crucial to obtain a good number of viable macrophages. It is important not extend the time under EDTA exposition more than recommended to preserve cell viability. Check the cells under the microscope after finishing the procedure to ensure they are fully detached.
3. Collect macrophages on a 1.5 mL microtube and wash them with 1 mL of 1x PBS-2% FBS by centrifugation at 400 × g for 5 min at 8-10 °C.
4. Resuspend the macrophages pellet with 200 µL of 1x PBS-5% FBS and transfer them to a 96-well round-bottom microplate for nonspecific blocking at RT for 15 min. Centrifuge the cells at 400 x g for 5 min at 8-10 °C.
5. Discard the supernatant by quickly inverting the plate.
6. Prepare the appropriate cocktail of directly conjugated antibodies in 1x PBS- 2% FBS using a 1:50 dilution factor for each antibody. Stain each sample with 50 µL of the antibody cocktail and incubate for 30 min in the dark on ice.
   NOTE: Antibody cocktail: CD11b-PE/Cy7 (200 µg/mL), CD64-AlexaFluor 647 (150 µg/mL), CD163-PerCP/Cy5.5 (200 µg/mL), HLADR-PE (40 µg/mL) and CD206-AlexaFluor 488 (400 µg/mL). Pre-tittered antibody concentration is listed in Table 1; catalog numbers can be found in the Table of Materials.
7. Wash the stained macrophages by adding 150 µL of 1x PBS and centrifugate at 400 × g for 5 min at 8-10 °C.
8. Discard the supernatant by quickly inverting the plate.
9. To measure cell viability, add 100 µL of the fixable viability dye Zombie Violet (1:1,000 in 1x PBS) and incubate in the dark for 30 min on ice.
10. Wash the stained cells by adding 100 µL of 1x PBS and centrifuging at 400 x g for 5 min at 8-10 °C. Repeat the washing procedure using 200 µL of 1x PBS.
11. Fix by resuspending the cell pellet with 20 µL of fresh prepared 1% paraformaldehyde and incubating them for 20 min on ice and in the dark.
    NOTE: 1% paraformaldehyde was used for fixation, without permeabilization. Optional: The commercial fixative component (from the BD fixative/permeabilization kit) may also be used for this step¹¹.
12. Wash cells by adding 180 µL of 1x PBS-2% FBS and centrifuge at 400 x g for 5 min at 8-10 °C. Repeat the washing procedure using 200 µL of 1x PBS-2% FBS.
13. Discard supernatant by plate inversion and resuspend the pellet in 200 µL of 1x PBS to take to the flow cytometry equipment for measurement.
    NOTE: If stained and fixed samples can't be run by flow cytometry on the same day, store them at 4 °C and shield them from light. Analyze the samples as soon as possible on the following days.
14. Analyze flow cytometry results using FlowJo software. The gating strategy starts by excluding debris using forward and side scatter (FSC/SSC). Subsequently, select live macrophages as CD11b⁺ Zombie Violet^- cells.
    NOTE: Because autofluorescence in macrophages is donor-dependent, it's essential to use controls derived from the same donor for each experiment.

Results

We generated triple-negative breast cancer tumor-associated macrophages (TAM)-like cells by treating monocyte-derived macrophages with 50% conditioned media (CM) from the MDA-MB-231 cell line. Monocytes were sorted from peripheral blood mononuclear cells (PBMC) of six independent healthy donors and differentiated following our protocol. The CM preparation utilized MDA-MB-231 cells at 90 % confluency (Figure 1). CM was added to the macrophage ...

Discussion

The protocol described here provides a reproducible and practical approach to study tumor-associated macrophage-like polarization by exposing human monocyte-derived macrophages to tumor cell line-conditioned media. This system captures the influence of soluble factors secreted by tumor cells, which are known to play a pivotal role in macrophage reprogramming within the tumor microenvironment^7,9,10,15...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Alexa Fluor 488 mouse anti-human  CD206 Antibody clone 15-2	BioLegend, San Diego, CA, USA.	Cat# 321113; RRID:AB_571874
Alexa Fluor 647 mouse anti-human CD64 Antibody clone 10.1	BioLegend, San Diego, CA, USA.	Cat# 305012; RRID:AB_528867
BD Cytofix/Cytoperm	BD Biosciences, San Jose, CA, USA	Cat# 554714	fixative/permeabilization kit
Dulbecco's Modified Eagle Medium (DMEM), high glucose	Gibco Thermo Fisher Scientific, Waltham, MA USA	Cat# 11965092
EasySep Human CD14 Positive Selection Kit; EasySep Positive Selection Cocktail; EasySep magnetic Nanoparticles; Easy Sep Magnet	STEMCELL Technologies, Vancouver, Canada	Cat# 18058
FACS Canto cytometer	Becton Dickinson, Franklin Lakes, New Jersey, USA.
Fetal Bovine Serum, qualified, New Zealand	Gibco™ Thermo Fisher Scientific, Waltham, MA USA	Cat# 10091148
Ficoll-Paque PLUS Cytiva	GE Healthcare - Life Sciences, USA	Cat# 17144003
FlowJo	FlowJo LLC.	RRID:SCR_008520
GraphPad Prism	GraphPad Software Inc.	RRID:SCR_002798
Partec CyFlow space cytometer	Sysmex Partec, Germany
PE anti-human HLA-DR Antibody clone L243	BioLegend, San Diego, CA, USA.	Cat# 307605; RRID:AB_314683
PE/Cy7 rat anti-mouse/human CD11b Antibody clone M1/70	BioLegend, San Diego, CA, USA.	Cat# 101216; RRID:AB_312799
PerCP/Cy5.5 mouse anti-human CD163 Antibody clone GHI/61	BioLegend, San Diego, CA, USA.	Cat# 333607; RRID:AB_1134006
RPMI 1640 Medium	Gibco Thermo Fisher Scientific, Waltham, MA USA	Cat# 11875119
Sodium pyruvate solution	Sigma-Aldrich Merck, San Luis, MO, USA.	Cat#  S8636
Trypsin-EDTA (0.25%), phenol red	Gibco Thermo Fisher Scientific, Waltham, MA USA	Cat# 25200056
Zombie Violet Fixable Viability Kit	BioLegend, San Diego, CA, USA.	Cat# 423113