The human monocytic cell lines U937 and THP-1 were purchased from American Type Culture Collection (ATCC, Manassas, VA, USA). Both cell lines were cultured in complete RPMI-1640 medium (Sigma-Aldrich, St. Louis, Missouri, USA) supplemented with 10% heat-inactivated fetal calf serum (Sigma-Aldrich, St. Louis, Missouri, USA) and 1% penicillin–streptomycin–glutamine (GIBCO, Carlsbad, CA, USA) in a 37°C, 5% CO₂/air atmosphere incubator.Human umbilical vein endothelial cells (HUVECs) (PromoCell, Heidelberg, Germany) were cultured in a flask coated with 0.2% gelatin type B solution (Sigma-Aldrich, St. Louis, Missouri, USA) and were maintained in complete Endothelial Cell Growth Medium 2 (PromoCell, Heidelberg, Germany). All cell washing steps were performed using 1X phosphate-buffered saline (PBS) (Amresco, Solon, OH, USA).

Human blood samples were collected from healthy donors with appropriate informed consent as approved by the Research Ethics Committee (Human) USM (USM/JEPeM/14120522). Briefly, 30–50 ml of blood was collected from each healthy donor. Adult peripheral blood mononuclear cells (PBMCs) were isolated by density gradient centrifugation using Lymphocyte Separation Medium (LSM) (Corning, Manassas, VA, USA). PBMCs were then collected from the plasma–LSM interface layer and washed with 1X PBS before being resuspended in the appropriate buffer or media.

Isolation of human whole-blood monocytes from PBMCs was performed using a Pan Monocyte Isolation Kit (Miltenyi Biotec, Auburn, CA, USA) according to the manufacturer’s protocol.

U937s, THP-1s, and blood monocytes were stimulated with 1 µg/ml lipopolysaccharide (LPS) from Escherichia coli (E. coli) 026:B6 (Sigma-Aldrich, St. Louis, Missouri, USA) for 18 hours at 37°C in a 5% humidified CO₂ incubator.

HUVECs were seeded onto a treated 6-well plate at 1x10⁵ cells/ml in complete Endothelial Cell Growth Medium 2. Cells were grown for 24 hours. To mimic inflammatory conditions, the confluent endothelial monolayer was either stimulated with 1 µg/ml LPS or incubated with 300 µg/ml9 unstimulated or LPS-stimulated monocytes derived mMPs for 18 hours at 37°C in a 5% CO₂/air atmosphere incubator. Trypsinization was performed using 0.25% trypsin-EDTA (Sigma-Aldrich, St. Louis, Missouri, USA) for 5 minutes at room temperature to detach the cells.

To isolate mMPs or eMPs, all culture supernatants were centrifuged at 500 xg for 5 minutes and 1,500 xg for 5 minutes, and finally ultracentrifuged at 20,000 xg for 60 minutes at 4°C to pellet down the MPs.

HUVECs and eMPs were stained with anti-CD31-PerCP/Cy5.5 (BioLegend, San Diego, CA, USA) and Annexin-V-FITC (BD Bioscience, San Jose, CA, USA) for 20 minutes in 1X binding buffer. Samples were analyzed by a FACS CANTO II flow cytometer (BD Bioscience, San Jose, CA, USA). HUVECs and eMPs were assessed on SSC/FSC profiles before being further gated based on the expression of CD31/Annexin-V.

Isolated mMPs were assessed for endotoxin interference using a ToxinSensor Chromogenic Limulus Amebocyte Lysate Endotoxin Kit (GenScript, Piscataway, NJ, USA).

THP-1 and U937 cells at a concentration of 5x10⁵ cells were cultured with their derived mMPs at a concentration of 3x10⁶ at a ratio of 1:6 (monocytic cell lines:mMPs). Whole monocytes at a concentration of 5x10⁵ cells were cultured in the presence of 1 µg/mL LPS with or without 1x10⁶ monocyte-derived mMPs in a 1:2 ratio (monocytes:mMPs)14. After 18 hours of culture, the supernatants were collected via centrifugation at 200 xg for 5 minutes and subsequently used for cytokine quantification using ELISA.

Confluent monolayers of HUVECs were serum-starved for 4 hours prior to culture with 300 µg/ml unstimulated or LPS-stimulated mMPs derived from monocytes for 2 hours in serum-free medium9 followed by RT–qPCR analysis.

ELISA was performed to quantify the levels of cytokines released using the Human TNF-α ELISA Max Deluxe and Human IL-1β ELISA Max Deluxe kits (Biolegend, San Diego, CA, USA). The level of cytokines released in the supernatants was measured using a spectrophotometer at 450 nm and analyzed by SkanIt Software 2.4.3 RE (Thermo Scientific, Waltham, MA, USA).

Total RNA was extracted using the RNeasy Mini Kit (Qiagen, Germantown, MD, USA) according to the manufacturer’s instructions, and the RNA concentration was determined using a NanoDrop ND-2000 spectrophotometer (Thermo Scientific, Waltham, MA, USA) at 450 nm. The detection of ICAM-1 and VCAM-1 was performed using RT–qPCR. Samples were prepared using the LUNA Universal One-Step RT–qPCR Kit (New England Biolabs, Ipswich, MA, USA) as recommended by the manufacturer. The primer sequences for RT–qPCR were as follows: GAPDH (forward: 5’ CCT GCA CCA ACT GCT TA 3’, reverse: 5’ GGC CAT CCA CAG TCT TCT GAG 3’) (NCBI, NM_002046.5), ICAM-1 (forward: 5’ GGC CGG CCA GCT TAT ACA C 3’, reverse: 5’ TAG ACA CTT GAG CTC GGG CA 3’) (NCBI, NM_000201.2), and VCAM-1 (forward: 5’ TCA GAT TGG AGA CTC AGT CAT GT 3’, reverse: 5’ ACT CCT CAC CTT CCC GCT C 3’) (NCBI, NM_001078.3). Data acquisition was performed using an Applied Biosystems 7500 RT–PCR machine (Applied Biosystems, Foster City, CA, USA), and the cycle threshold was obtained by ABi 7500 Real Time-PCR software (Applied Biosystems, Foster City, CA, USA).

The data obtained were analyzed using GraphPad 7 Prism software (De Novo, San Diego, CA, USA). Data are shown as the mean ± SD. Mean differences between the two groups were tested using a paired t-test. *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001 were considered significant.

Figure 1

Figure 2

Figure 3

Monocyte subtypes exhibit different cytokine secretion in the presence of mMPs⁶ mMPs. We found that culturing mMPs derived from stimulated U937 cells with their origin cells resulted in low secretion of IL-1β compared to U937 cells alone (p < 0.05) (Figure 1 A). In contrast, the secretion of IL-1β in the supernatants of stimulated THP-1-derived mMPs cultured with their origin cells was significantly increased compared to that of THP-1 cells cultured alone (p < 0.01) (Figure 1 B). However, mMPs derived from both stimulated U937 and THP-1 cells cultured with U937 and THP-1 cells failed to induce TNF-α secretion despite TNF-α being detected in the culture supernatants of U937 and THP-1 cells alone (data not shown).

We then tested whether mMPs derived from monocytes enhanced TNF-α and IL-6 secretion. The release of TNF-α from unstimulated monocytes in the presence of 1x10⁶ mMPs was low compared to that from unstimulated monocytes without mMPs (p < 0.01) (Figure 1 C). Similarly, TNF-α production from stimulated monocytes cultured with mMPs was also low compared to that from stimulated monocytes in the absence of mMPs (p < 0.01). IL-6 production by unstimulated monocytes cultured with mMPs was significantly higher than that from unstimulated monocytes cultured without mMPs (p < 0.05) (Figure 1 D). Meanwhile, IL-6 production from the supernatants of stimulated monocytes cultured with mMPs was also higher than that from stimulated monocytes cultured without mMPs, although the difference was not significant.

Recently, the study of mMPs has become important due to their association with monocyte and endothelial cell activation, particularly in inflammation. Emerging evidence has demonstrated the proinflammatory effects of MPs derived from distinct origin cells on monocytes and macrophages5, 14, 15. However, the interaction between mMPs and monocytes at inflammatory sites has not been explored. Therefore, the effects of mMPs during culture with their origin cells were assessed. In this study, LPS was used to induce the activation of CD14 on monocytes and mimic cellular inflammation, which subsequently produces mMPs. To isolate mMPs, cultured supernatants containing mMPs were subjected to two-step differential centrifugation at 500 xg and 1,500 xg, followed by ultracentrifugation at 20,000 xg as previously recommended16. Two-step differential centrifugation at 500 xg and 1,500 xg is crucial for removing cells and apoptotic bodies17, respectively, from mMPs in supernatants. Meanwhile, ultracentrifugation at 20,000 xg during mMP isolation eliminates exosome contamination and ensures proper separation, since mMPs are pelleted while exosomes remain in the supernatants18.

Our data showed that in the presence of mMPs derived from THP-1 cells and blood monocytes, IL-1β and IL-6, respectively, were secreted. In contrast, mMPs derived from U937 cells were unable to modulate the secretion of IL-1β by U937 cells despite having an mMP concentration comparable to THP-1 cells. In addition, the presence of mMPs did not trigger TNF-α secretion by U937, THP-1, or monocytes. These data indicate that mMPs may modulate proinflammatory cytokine secretion in a cell-origin-dependent manner. It has been previously reported that upon culture with MP, distinct cell types may release varying levels of different cytokines19. The variations in cytokine expression detected in our study were not surprising since U937 and THP-1 cells show different proteomic and cytokine profiles than blood monocytes20 and thus play different roles in inflammation. Similar to our study, a previous finding showed that mMPs containing IL-1 trigger the release of IL-1β from monocytes in an autocrine manner9. Although the exact underlying mechanism of monocyte activation by mMPs remains unknown, it is possible that mMPs contribute to an inflammatory response via their lipid fraction, which triggers the activation of Toll-like receptor 4 on monocytes as previously reported21. Moreover, the influences of mMP concentration on their biological effect have been greatly emphasized previously22. In this study, 3x10⁶ THP-1-derived mMPs and 1x10⁶ blood monocyte-derived mMPs activated their origin cells at different levels. Nevertheless, the correlations between different concentrations of mMPs and their biological effects on the activation of U937, THP-1, and blood monocytes were not directly assessed in this study. Several variables, including cell types, cell concentrations, and types of cytokines measured, should be considered for proper assessment in future concentration-response studies.

As elevated numbers of mMPs have been observed in inflammation23, the proinflammatory properties of mMPs in this study may be explained by their potential to enhance IL-1β and IL-6 release from monocytes, thereby exacerbating inflammation. In addition, our data have shown that mMPs inhibit the release of TNF-α by monocytes upon culture with mMPs. This finding suggests that mMPs may initiate a different mechanism that promotes an anti-inflammatory response in effector cells, as has been previously shown3. However, the pathway by which mMPs inhibit TNF-α has not been explored. A previous report has suggested that mMPs may increase the production of prostaglandin E224, resulting in the inhibition of TNF-α release by monocytes to resolve inflammation25. These data suggest that mMPs may act as TNF inhibitors, thus reducing inflammation.

Apart from cytokine release, the interaction between mMPs and endothelial cells is important in the progression of inflammation26. In this study, we therefore assessed the expression of the endothelial cell activation markers CD31, ICAM-1, and VCAM-1. Our data demonstrated that mMPs derived from LPS-stimulated monocytes upregulated CD31, ICAM-1, and VCAM-1, which is similar to a previous report on THP-19. This suggests the potential role of mMPs in activating endothelial cells in addition to exosomes27. A previous study suggested that mMPs may mediate the activation of endothelial cells via an IL-1β-dependent pathway9, although the exact mechanism was not assessed in our study. It has been previously shown that THP-1-derived mMP regulation of endothelial cells involves the activation of intracellular signaling pathways by ERK1/2 phosphorylation and IĸB-α degradation, resulting in NF-ĸB translocation and leading to the increased expression of ICAM-1, VCAM-1, and E-selectin by endothelial cells9. Additionally, our results suggest that mMPs may contribute to the amplification of inflammation and vascular injury, as previously reported28, by elevating the expression of adhesion molecules and by facilitating monocyte migration through endothelial cells to sites of inflammation29.

Additionally, as mMPs are also released constitutively under physiological conditions, especially during cell growth30, 31, their roles in physiology during steady state should not be ignored. It has been previously reported that under normal conditions, MPs derived from platelets mediate the transfer of adhesion molecules from platelets to hematopoietic cells and endothelial cells, enhancing their adhesion capacity and engraftment32. Similarly, we found that CD31, ICAM-1, and VCAM-1 expression by endothelial cells can be enhanced by mMPs derived from unstimulated monocytes. This finding suggests that this mMP may aid in cell adhesion to endothelial cells.

The ability of mMPs to induce vesiculation in eMPs, in addition to CD31 and adhesion molecule expression in endothelial cells, was also assessed. This study demonstrates that mMPs enhance the release of eMPs in culture conditions with endothelial cells, suggesting that they may be involved in the exacerbation of pulmonary and capillary leakage as previously demonstrated in vivo33. A previous report showed that eMP elevation is a hallmark of endothelial cell activation34, while others have demonstrated that mMPs are able to induce the release of eMPs in an in vitro brain inflammation model9. Taken together, mMPs are not only cell residues but are also capable of altering the responses of their effector cells by acting as biological mediators in cell-to-cell interactions22.

This is the first study conducted using human blood monocyte-derived mMPs on monocyte and endothelial cell activation. Monocytic MPs cause deleterious effects, particularly under pathological conditions. Monocytic MPs derived from different origin cells may display distinct cytokine profiles, thus regulating effector cells via different cytokine mechanisms. Additionally, the ability of mMPs derived under pathogenic conditions to elicit high expression of adhesion molecules by endothelial cells and to induce vesiculation of eMPs may be the key to worsening inflammation, thus promoting the progression of endothelial dysfunction. However, further studies are necessary to better understand the underlying mechanism mediated by mMPs in inflammation.

eMP: Endothelial microparticles, ERK1/2: Extracellular regulated kinase 1/2, HUVEC: Human umbilical vein endothelial cells, ICAM-1: Intracellular adhesion molecule 1, IL-1: Interleukin 1, LPS: Lipopolysaccharides, MP: Microparticles, mMP: Monocytic microparticles, NF-ĸB: Nuclear Factor ĸB, TNF- α: Tumor necrosis factor α, TLR: Toll-like receptor VCAM-1: Vascular cell adhesion molecule 1

NAMN was supported by the USM Fellowship scheme and USM Graduate Assistance scheme. NAFMA was supported by MyBrain15 and USM Graduate Assistance scheme.

Conceptualization and design of the study: Rapeah Suppian and Maryam Azlan

Acquisition of data: Nur Azira Mohd Noor, Siti Zulaiha Marhalim, and Nur Azrah Fazera Mohd Ariffin.

Analysis and/or interpretation of data: Nur Azira Mohd Noor, Siti Zulaiha Marhalim and Nur Azrah Fazera Mohd Ariffin.

Drafting the manuscript: Nur Azira Mohd Noor, Siti Zulaiha Marhalim, and Nur Azrah Fazera Mohd Ariffin.

Revising the manuscript critically for important intellectual content: Nur Azira Mohd Noor and Maryam Azlan

Approval of the version of the manuscript to be published: Nur Azira Mohd Noor, Siti Zulaiha Marhalim, Nur Azrah Fazera Mohd Ariffin, Rapeah Suppian, and Maryam Azlan

All authors read and approved the final manuscript.

This work was supported by the Short Term Grant, Universiti Sains Malaysia (Grant number: 304/PPSK/61313135).

Data and materials used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Not applicable.

Not applicable.

The authors declare that they have no competing interests.