Augmentation of Human Monocyte Responses to Lipopolysaccharide by the Protein S and Mer/Tyro3 Receptor Tyrosine Kinase Axis

In response to injury or infection, dynamic temporal changes within the inflammatory microenvironment act to coordinate inflammatory cell recruitment (1), cell activation, tissue repair, (2) and ultimately, regeneration (3). Apoptosis has a key role in the termination and resolution of the inflammatory response. Recruited inflammatory cells that do not emigrate from the inflammatory site (4) undergo apoptosis (5) and are subsequently cleared by phagocytosis (also termed efferocytosis) (6, 7). In addition, resolution of inflammation and the restoration of normal tissue function is facilitated by functional alterations in phagocytes as a consequence of signaling events downstream of apoptotic cell recognition (8, 9).

The Tyro3/Axl/Mer (TAM) family of receptor tyrosine kinases (RTK) is widely expressed on phagocytic cells in many different tissue settings and forms one of the key molecular pathways by which phagocytes specifically recognize and subsequently internalize apoptotic cells (10). TAMs are able to bind to two specialized ligands, protein S (Pros1) and growth arrest specific protein 6 (Gas6) (11), that interact with the anionic phospholipid, phosphatidylserine (PtdSer), exposed on the plasma membrane of apoptotic cells following loss of plasma membrane asymmetry (12, 13). Gas6 functions as a ligand for all three TAM receptors, whereas Pros1 serves as a ligand for only Tyro3 and Mer, with little or no binding to Axl (14). The presence of a highly conserved Gla domain, containing multiple glutamic acid residues that are γ-carboxylated in a vitamin K–dependent reaction (15), confers the capacity of Pros1 and Gas6 to interact with PtdSer in a Ca²⁺-dependent manner (16).

TAM binding to Gas6 or Pros1 is critical for the physiological removal of apoptotic cells, and the absence of TAM-dependent apoptotic cell clearance mechanisms is associated with failed spermatogenesis in the testis (17), photoreceptor degeneration in the eye (18), and development of features of autoimmune and autoinflammatory disease (17, 19, 20). Signaling downstream of TAM ligation provides feedback inhibition of inflammatory cytokine production following TLR-driven responses (21, 22), allowing termination of inflammatory responses. Mer-dependent signaling in phagocytes promotes the internalization of apoptotic cells via autophosphorylation at Y867 and activation of PI3K, phospholipase C (PLC)-γ2, and protein kinase C (PKC), leading to focal adhesion kinase (FAK) phosphorylation that was not required for the suppression of NF-κB (23).

In this article, we have investigated the expression and function of TAMs by human monocytes, identifying a novel role for TAM signaling in the regulation of monocyte responses to LPS. We show that human monocytes express predominantly Mer, with low levels of Tyro3 and little or no expression of Axl. Our findings confirm that viable monocytes expose low levels of PtdSer (24). Pros1 was shown to bind to the surface of viable monocytes via both a Ca²⁺-dependent and -independent mechanism, likely to represent PtdSer-dependent and TAM-dependent binding interactions, respectively. Surprisingly, we found that Pros1 acts in a PtdSer-dependent manner to augment monocyte release of proinflammatory cytokines (TNF and IL-6) in response to LPS. Augmentation of monocyte proinflammatory cytokine release was attenuated when TAM RTK activity was blocked. We propose that low-level exposure of PtdSer on viable monocytes allows Pros1 binding to monocytes that leads to augmentation of proinflammatory cytokine production, identifying a potentially important novel role for TAM-mediated signaling during the initiation phase of inflammation.

Reagents were obtained from Sigma-Aldrich (Gillingham, U.K.) unless otherwise stated. Cell culture ware was purchased from either BD Biosciences (Oxford, U.K.) or Corning, and IMDM (with the addition of 1% l-glutamine, 25 mM HEPES) was purchased from Thermo Fisher Scientific (Paisley, U.K.). Human Pros1 was obtained from Enzyme Research Laboratories (Swansea, U.K.). Pros1 lacking the Gla domain (ΔGla Pros1) was generated using overlap-extension mutagenesis to delete the Gla domain from the Pros1 gene using plasmid pCIS2M-PS (25). The following primers were used: A = 5′-TCGATTGCTAGCGCACAG-3′; B = 5′-GAGCGAAGACAAACACGACGCTTCCTAACCAGG-3′; C = 5′-CCTGGTTAGGAAGCGTCGTGTTTGTCTTCGCTC-3′; and D = 5′-GACACAAAGCTGAGCACACAT-3′. These primers result in deletion of 135 nt coding residues Ala1 to Leu45, leaving the prepropeptide sequence (Met-41 to Arg-1) connected to Val46. The resulting PCR fragment was inserted after digestion with NheI and BlpI into the Pros1 gene in the pcDNA3.1⁺ expression plasmid to generate pcDNA3.1–∆Gla-PS. ∆Gla-PS was expressed in HK293 cells and purified by immunoaffinity chromatography (26).

Recombinant mouse Gas6 (full-length and Gla-less) was prepared as described (14) and provided by Prof. G. Lemke and Dr. E. Lew (Salk Institute for Biological Studies, La Jolla, CA). PE- or allophycocyanin-conjugated mouse mAbs (IgG1 unless otherwise indicated) against human CD14 (MΦP9, IgG2b), CD16 (3G8), CD62L (DREG56), CD162, (KPL1) CD11b (ICRF44), and isotype controls were obtained from BD Biosciences. PE-conjugated mouse mAbs against human Mer (122518, IgG2b), Tyro3 (96201, IgG1), and Axl (108724, IgG1) together with relevant PE-conjugated isotype controls (IgG1 and IgG2b) were from R&D Systems (Abingdon, U.K.). A rabbit monoclonal anti-Mer for immunoblot analysis (clone D21F11) was obtained from Cell Signaling Technology (Danvers, MA). A second PE-conjugated mouse mAb against human Mer (590H11G1E3, IgG1) was obtained from BioLegend (London, U.K.) to validate flow cytometry findings. ELISA reagents and Abs against IL-6 and TNF were obtained from R&D Systems. BMS777607 was obtained from Selleck Chemicals (Stratech, Newmarket, U.K.). Allophycocyanin-conjugated Annexin V was obtained from Thermo Fisher Scientific, and propidium iodide was obtained from BioLegend. LPS (Escherichia coli 0127:B8) was purchased from Sigma-Aldrich. Recombinant Human TNF, Human TNF DuoSet ELISA, and Human IL-6 Quantikine ELISA were from R&D Systems.

Peripheral blood leukocytes (human blood mononuclear cells and polymorphonuclear cells) were isolated from healthy volunteers as described (27). In brief, citrated blood (3.8%) was centrifuged at room temperature at 350 × g for 20 min; platelet-rich plasma was removed. Autologous serum was obtained by addition of CaCl₂ to a final concentration of 22 mM. Leukocytes were separated from erythrocytes by dextran sedimentation and further fractionated using isotonic Percoll (GE Healthcare) 50%/70%/81% discontinuous gradients. Polymorphonuclear leukocytes (>95% CD16+ve, hereafter termed “neutrophils”) were harvested from the 70%/81% interface and cultured for 18 h in IMDM supplemented with 5% autologous serum to induce apoptosis. Monocytes were enriched from the mononuclear cells (55%/70% interface) by negative selection using a Pan Monocyte Isolation Kit (Miltenyi Biotec) as described by the manufacturer. Monocytes were resuspended at 1.5 × 10⁶ cells/ml and cultured for 18 h in IMDM supplemented with 5% autologous serum in Falcon tissue culture plates (BD Biosciences). Monocyte-derived macrophages (MDM) were obtained by in vitro culture of monocytes for 6 d in IMDM supplemented with 5% autologous serum with the addition of 250 nM dexamethasone to induce a macrophage phenotype that predominantly uses Mer-mediated phagocytosis of apoptotic cells (28).

Phagocytosis of pHrodo-labeled apoptotic neutrophils by monocytes or MDM was assessed by flow cytometry as described (29, 30). To ensure TAM ligand–free conditions in the phagocytosis assay, cells were washed with HBSS without Ca²⁺/Mg²⁺ containing 2.5 mM EDTA to remove exogenous TAM ligands. Monocytes were labeled with CellTrace Far Red DDAO-SE (Thermo Fisher Scientific), according to the manufacturer’s instructions, prior to phagocytosis assay. For experiments using the RTK inhibitor BMS777607, monocytes or MDM were preincubated for 20 min with 100 nM BMS77607 prior to coincubation with pHrodo-labeled apoptotic neutrophils. For some assays, Pros1 or ΔGla Pros1 was added to a final concentration of 25 nM, as detailed in the figure legends. MDM or monocytes were then overlaid with apoptotic neutrophils at a phagocyte/target ratio of 1:6. After coculture for 40–45min at 37°C, noningested apoptotic cells were removed by aspiration and monocytes, or MDM were detached by vigorous pipetting following incubation with 0.5% Trypsin/1 mM EDTA solution for 5 min at 37°C. The percentage of monocytes or MDM that were fluorescent (corresponding to those that had ingested apoptotic neutrophils) was determined by flow cytometric analysis (31).

Proteins were labeled with Dylight488 or Dylight650 dye as recommended by the manufacturer (Thermo Fisher Scientific). Following incubation with 7–10-fold molar excess of dye for 45 min in the dark at 20°C, the reaction was terminated by addition of glycine (final concentration: 20 mM). Labeled protein was layered onto a 7K m.w. cut-off protein desalting column (Pierce; Thermo Fisher Scientific) pre-equilibrated in PBS to remove unbound dye and centrifuged for 30 s at 3000 × g. The degree of protein labeling was estimated from measurement of absorbance at 280 and 650 nm (Dylight 650) or 280 and 493 nm (Dylight 488), which was routinely between 2.5 and 3.5 of dye per mole of protein. Functionality of labeled Pros1 and Gas6 was confirmed by their capacity to augment TAM-dependent phagocytosis of apoptotic neutrophils (data not shown).

All incubations were performed on ice to prevent internalization of Ab. Adherent MDM were detached by incubation in ice-cold HBSS without divalent cations (HBSS^−/−), containing 0.5% (w/v) BSA and 2 mM EDTA for 15 min followed by vigorous pipetting. Detached cells were washed with HBSS^−/− containing 2 mM EDTA (3 × 10⁵–1.5 × 10⁶ per assay) and resuspended in 20 mM HEPES (pH 7.4) containing 0.14 M NaCl and 0.1% (w/v) BSA (flow buffer). Nonspecific binding of Abs to Fc receptors was reduced by preincubation for 5 min with 2% autologous serum. Cells were then labeled with saturating concentrations of fluorophore-conjugated Abs for 30 min on ice. For assessment of binding of TAM ligands or Annexin V, cells were incubated with saturating concentrations of fluorescently labeled proteins diluted in 20 mM HEPES (pH 7.4) containing 0.14 M NaCl with or without the addition of 2 mM CaCl₂ or 2.5 mM EDTA for 10 min on ice. Data were acquired using either a five-laser LSR or two-laser FACSCalibur flow cytometer (BD Biosciences) and analyzed with FlowJo software (FlowJo, Ashland, OR).

Cytokine release from 18-h cultured monocytes following stimulation with 2 ng/ml LPS, 10 ng/ml IL-1β, or 50 ng/ml TNF was measured by ELISA. Monocyte culture media were removed, and cells were washed with HBSS without divalent cations containing 2 mM EDTA. Fresh IMDM was added containing 25 nM Pros1, Gla-less Pros1, or BMS777607. After a 20-min incubation in the presence or absence of Pros1, ∆Gla Pros1, or inhibitor, LPS (2 ng/ml) was added, and cells were cultured for 8 h at 37°C with 5% CO₂. Cell culture supernatant was harvested and centrifuged at 6000 × g for 5 min to remove any cell remnants and stored at −20°C. ELISA for TNF, IL-6, IL-1β, and IL-8 were performed as indicated by manufacturer.

Results are presented as mean ± SD or SEM as indicated, in which n is the number of independent experiments using cells obtained from different donors and for multiple comparison tests; significance was analyzed by ANOVA with Mann–Whitney posttest with Bonferroni correction for multiple comparisons using InStat software (http://www.graphpad.com).

The expression of the TAM family of RTK on human PBMCs was examined by flow cytometry. Monocytes were identified on the basis of forward/side scatter characteristics (Fig. 1A, upper panel) and expression of CD11b, CD11c, CD14, CD16, CD64, and HLA-DR (Supplemental Fig. 1). We then used differential expression of CD14/CD16 (Fig. 1A, lower panel) to subdivide the monocytes into CD14⁺⁺/CD16⁻ and CD14⁺/CD16⁺ populations (32) to determine the expression of Tyro3, Axl, and Mer on these subsets (Fig. 1B). We did not detect expression of Axl on either of the monocyte populations when compared with binding of an isotype control Ab (Fig. 1B). In contrast, we consistently observed low levels of expression of Tyro-3 on monocytes (Fig. 1B). Mer expression was ∼2-fold higher for the CD16⁺-expressing monocytes (Fig. 1B and quantified in Fig. 1C). To exclude the possibility that the expression of Mer we observed was mAb dependent, a second anti-Mer Ab was used with similar results. In these experiments, Mer expression on CD16⁺ monocytes was found to be 1.7-fold higher than that of CD16-negative monocytes (n = 3). Expression of Mer was higher for CD16⁺ monocytes further subdivided into high or intermediate expression when compared with CD16-negative monocytes (relative mean fluorescence ± SD) (Mer on CD16 high: 1579 ± 340; Mer on CD16 intermediate: 2047 ± 202; Mer on CD16 negative: 862 ± 85; IgG control: 280 ± 100; n = 3). Expression of TAMs was not a consequence of platelet binding to monocytes (33) as extensive washing of cells with EDTA to remove bound platelets did not affect Tyro3 or Mer expression. Freshly isolated human lymphocytes did not express Tyro3, Axl, or Mer (data not shown).

We next investigated monocyte capacity to bind to TAM ligands. Because viable cells have been reported to expose low levels of PtdSer on their plasma membrane (24), which would potentially allow binding of TAM ligands, we examined binding of fluorescently labeled Pros1 or Gas6 in the presence or absence of Ca²⁺ to dissect PtdSer-dependent and PtdSer-independent components of binding. In addition, we compared binding of Annexin V to monocytes under the same assay conditions. Binding of Annexin V to monocytes was found to be strictly Ca²⁺ dependent, with no binding in the presence of EDTA (Fig. 2A). Importantly, Annexin V binding occurred independently of any morphological (nuclear condensation) or biochemical (activation of caspase 3) signs of monocyte apoptosis (Supplemental Fig. 2 and data not shown). Furthermore, the levels of Annexin V binding to viable monocytes was 12.5 ± 3.3-fold lower than that for apoptotic monocytes, consistent with previous reports (24). These findings suggest that there is constitutive, low-level exposure of PtdSer on the plasma membrane of viable monocytes.

Binding of both Pros1 and Gas6 (data not shown) to monocytes was distinct from Annexin V binding in that there were Ca²⁺-dependent and -independent components (Fig. 2A, quantified in Fig. 2B). When we compared binding of plasma-derived Pros1 and ΔGla Pros1, we observed binding to monocytes in the absence of Ca²⁺, consistent with a TAM-dependent binding component. In contrast, we did not observe Ca²⁺-dependent binding of ΔGla Pros1 (Fig. 2C). Together, our data demonstrate that Pros1 binds to monocytes in both a PtdSer-dependent and -independent manner and that the PtdSer-dependent component of Pros1 binding requires the presence of the Gla domain.

Neutrophils undergo spontaneous apoptosis during in vitro culture (6), and a significant proportion of 18-h cultured neutrophils expose PtdSer. Annexin V [(34), data not shown] and Pros1 bound to a subset of neutrophils in the presence of Ca²⁺ but not in the presence of EDTA (28). These results contrast the data shown in Fig. 2 for monocytes and suggest that Pros1 only binds to apoptotic neutrophils via exposed PtdSer and that there was no Ca²⁺-independent binding. This suggestion was confirmed by demonstration of cobinding of Annexin V and Pros1, revealing that Annexin V–positive cells also bound Pros1 (Fig. 3A), consistent with previous data for apoptotic thymocytes (35). Because binding of Pros1 to PtdSer is dependent on amino acid sequences present in the Gla domain, we predicted that ΔGla Pros1 would not bind to apoptotic neutrophils. As shown in Fig. 3B, we observed very low-level binding of ΔGla Pros1 that was independent of the presence of Ca²⁺ (data not shown). Quantification of binding demonstrates that ΔGla Pros1 binds to apoptotic neutrophils at very low levels, consistent with the absence of the Ca²⁺-dependent PtdSer binding domain (Fig. 3C). A small population of 18-h cultured neutrophils were found to bind to both Pros1 and ΔGla Pros1 at high levels in either the presence or absence of Ca²⁺. These cells corresponded to propidium iodide–positive necrotic cells (data not shown). Our data confirm high-level opsonization of apoptotic neutrophils with Pros1 but not ΔGla Pros1, consistent with binding to apoptotic neutrophils via interaction with PtdSer exposed on the plasma membrane.

We next compared 18-h cultured monocyte and MDM capacity for Pros1-dependent phagocytosis of apoptotic neutrophils. We used an experimental system in which monocytes and neutrophils were isolated from the same donor and cultured for 18 h prior to use in an autologous cell interaction assay. Monocytes were isolated by negative immunomagnetic selection to yield cells that were 89.3 ± 3.8% (n = 8) CD64⁺ and were essentially free of bound platelets. Following in vitro culture for 18 h, monocytes maintained expression of CD11b, CD162, Mer, and Tyro3 (Supplemental Fig. 3A). Mer expression on 18-h cultured monocytes was further confirmed by immunoblot analysis (Supplemental Fig. 3C). Axl was not expressed by monocytes that had been cultured for 18 h. In contrast to freshly isolated monocytes, expression of CD62L was downregulated on 18-h cultured monocytes, consistent with initiation of monocyte–macrophage differentiation (Supplemental Fig. 3A). Because there was considerable overlap in laser scatter properties of cultured neutrophils and monocytes, we labeled monocytes with CellTrace Far Red to allow discrimination from apoptotic neutrophil targets. Neutrophils were labeled with pHrodo to allow quantification of monocyte phagocytosis using flow cytometry. In the absence of Pros1, there was a small proportion of phagocytic monocytes (<5% phagocytosis; see example plots in Fig. 3D, quantified in Fig. 3E). Although phagocytosis was increased ∼2-fold in the presence of Pros1, the absolute percentage (∼6%) of monocytes capable of Pros1-dependent phagocytosis was low (Fig. 3E). Pros1-dependent phagocytosis of apoptotic neutrophils by monocytes was blocked by preincubation with the RTK inhibitor, BMS777607, demonstrating a requirement for RTK activity (36). Consistent with the lack of binding to apoptotic neutrophils shown in Fig. 3C, addition of Gla-less Pros1 failed to increase monocyte phagocytosis of apoptotic neutrophils (Fig. 3D and quantified in Fig. 3E), demonstrating a requirement for the PtdSer-binding domain of Pros1. Together, these data demonstrate that although 18-h cultured monocytes express both Mer and Tyro3, the capacity for Pros1-dependent phagocytosis of apoptotic cells is relatively low.

To confirm that Pros1 was able to confer phagocytosis by MDM, we treated monocytes with glucocorticoids for 5 d to induce high levels of expression of Mer [(36, 37), Supplemental Fig. 3D, 3E] and to promote Mer-dependent apoptotic cell phagocytosis. The proportion of glucocorticoid-treated (GC) MDM capable of phagocytosis of pHrodo-labeled apoptotic neutrophils was ∼10-fold higher than that of monocytes (baseline phagocytosis, 44% versus 4.2%, respectively; compare Figs. 3E, 4B). Phagocytosis of apoptotic neutrophils by GC-MDM was increased 1.9-fold in the presence of 25 nM Pros1 (Fig. 4A; quantified in Fig. 4B). However, the absolute percentage increase in phagocytosis for GC-MDM in the presence of Pros1 was ∼40%, with the majority of GC-MDM being phagocytic after 45 min. The Pros1-dependent increase in phagocytosiswas not observed when GC-MDM were preincubated with the RTK inhibitor BMS777607, consistent with a requirement for Mer. In contrast, preincubation with BMS777607 did not affect GC-MDM phagocytosis of apoptotic neutrophils in the absence of Pros1. Finally, we tested our prediction that the ΔGla Pros1 would fail to confer increased phagocytosis by GC-MDM. When compared with full-length Pros1, equimolar concentrations of ΔGla Pros1 did not augment phagocytosis (Fig. 4A; quantified in Fig. 4B). Together, our data demonstrate that ΔGla Pros1 does not bind to apoptotic neutrophils and fails to act as a ligand for Mer-dependent phagocytosis of apoptotic cells by either 18-h cultured monocytes or GC-MDM.

Although monocytes were inefficient phagocytes for apoptotic cells, they expressed both Mer and Tyro3 and were able to bind to Pros1. We therefore sought to examine the impact of Pros1 on monocyte proinflammatory cytokine production. Monocytes were cocultured with apoptotic neutrophils in the presence or absence of the TLR4 ligand, LPS, for 8 h. Consistent with published findings, we observed powerful suppression of TNF release in response to LPS following coculture of monocytes with apoptotic neutrophils (Fig. 5A). Suppression of TNF release was observed whether Pros1 was present or not, indicating that the suppression was independent of Pros1 opsonization (Fig. 5A). We observed similar findings in experiments using MDM that had been cultured in IMDM and serum, although the effect was not as pronounced as for monocytes (Supplemental Fig. 4). However, because monocytes are not efficient phagocytes of apoptotic cells (see Fig. 3E), suppression of monocyte inflammatory cytokine production following coculture with apoptotic cells was likely independent of phagocytosis (38). In the course of these experiments, we were surprised to find that when monocytes were incubated with LPS and Pros1 in the absence of apoptotic cells, there was a significant, 2-fold increase in TNF release. We confirmed that the Pros1 used in these experiments did not contain significant levels of contaminating endotoxin, with levels of LPS being <2 pg/ml at the concentrations used in this assay. Furthermore, incubation of monocytes with Pros1 in the absence of LPS failed to induce TNF release (Fig. 5A), suggesting our results were not a direct effect of Pros1 on monocyte TNF release. These data raised the possibility that engagement of monocyte TAMs in the absence of apoptotic cells acted to amplify TLR responses.

The augmentation of LPS-induced TNF release from monocytes in the presence of Pros1 was concentration dependent, with significant effects at 75, 25, and 7.5 nM Pros1 (Fig. 5B). Furthermore, pretreatment of monocytes with the RTK inhibitor BMS777607 to block TAM-dependent signaling prior to incubation with Pros1 and LPS inhibited the augmentation of TNF release, suggesting that TAM-dependent signaling was required (Fig. 5C). In contrast, ΔGla Pros1 failed to augment monocyte TNF production in response to LPS, indicating a dual requirement for the Gla domain and PtdSer binding for augmentation of LPS-induced TNF release (Fig. 5D). Importantly, the effect of Pros1 was not restricted to the release of TNF, as LPS-induced production of IL-6 was also increased in the presence of Pros1 (Fig. 5E). Together, these data suggest that Pros1 has a synergistic proinflammatory effect upon monocyte cytokine production/release in response to TLR4 ligands.

In this manuscript, we present important new findings relating to the TAM-mediated regulation of myeloid cell function.

First, human peripheral blood monocytes express Mer and low levels of Tyro3 but not Axl. Levels of expression of Mer were higher on CD14⁺/CD16⁺-expressing monocytes than CD14⁺⁺ monocytes (Fig. 1), consistent with increased expression associated with maturation status. However, we found that Mer-expressing monocytes were relatively poor phagocytes of Pros1-opsonized apoptotic cells (Fig. 3). These data suggest that, for monocytes, additional receptors are required for the capture and internalization of apoptotic cells (39). Alternatively, there may be a threshold level of Mer or Tyro3 expression required to confer function as a phagocytic receptor (40).

Second, we observed low-level binding of Annexin V to viable monocytes [as reported by others (24)], suggesting that PtdSer is exposed on the plasma membrane of viable monocytes. PtdSer is exposed on the surface of activated platelets where it has an important role in the regulation of coagulation (41). PtdSer exposure on viable cells is also critical for “pruning” of neuronal synapses by astrocytes and microglia (42) and removal of the outer segments of rod cells in the retina by retinal pigment epithelial cells (43). Exposure of PtdSer in viable myoblasts is critical for myotube formation (44). In addition, viable or activated leukocytes have also been shown to expose PtdSer (24, 45, 46), suggesting that PtdSer exposure is not restricted to apoptotic cells.

Using fluorescently labeled Pros1 and Gas6, we demonstrated that viable monocytes bind to TAM ligands (Fig. 2) and that low-level exposure of PtdSer on monocytes allows Ca²⁺-dependent binding of Pros1 that requires the presence of the Gla domain. However, we also identified a Ca²⁺-independent component of Pros1 binding that does not require the Gla domain, which we speculate is mediated via the sex hormone–binding globulin domain of Pros1. Ca²⁺-independent binding of TAM ligands was not observed for viable or apoptotic neutrophils (Fig. 3A) or lymphocytes (data not shown).

Third, we observed Pros1-dependent augmentation of monocyte TNF release in response to LPS. This effect of Pros1 markedly contrasts the suppression of LPS-induced inflammatory cytokine release observed in the presence of apoptotic cells (9). One possibility is that Pros1 acted to alter monocyte viability or metabolism. However, we did not observe differential recovery of monocytes following incubation with Pros1 (82% versus 85%). Furthermore, metabolic profiling using Seahorse analysis did not reveal changes in aerobic or anaerobic energy use in Pros1-treated monocytes (data not shown). Data from preliminary experiments suggest that Pros1 also amplifies release of MCP1 and CXCL8 from LPS-treated monocytes (data not shown). In view of the lack of effect of Pros1 alone upon monocyte TNF release, we suggest that Pros1 has a synergistic effect upon LPS-mediated regulation of monocyte cytokine production/release. Augmentation of monocyte proinflammatory cytokine production by Pros1 was not seen with ΔGla Pros1, demonstrating a requirement for PtdSer binding for the observed augmentation of monocyte cytokine release.

Fourth, augmentation of LPS-induced TNF release by Pros1 was blocked by RTK inhibition, implying a requirement for TAM kinase activity. Previous studies have suggested that there are divergent signaling mechanisms downstream of Mer autophosphorylation, with activation of FAK/PLCγ being required for Rac activation necessary for phagocytosis (23). In contrast, Mer-dependent suppression of NF-κB activation has been suggested to be independent of this pathway (23). Results from preliminary experiments showing that Pros1 does not affect proinflammatory cytokine release (TNF and CXCL8) in response to IL-1 (IL-1β) stimulation raise the possibility that there is a specific association between Mer and/or Tyro3 and TLR4-mediated signaling that leads to amplification of proinflammatory cytokine production. LPS has been reported to induce downregulation of Mer, with loss of receptor expression [(47), Supplemental Fig. 3B] and downstream suppressive signaling potentially contributing to generation of proinflammatory cytokines (48). We therefore examined whether Pros1 affected the LPS-induced Mer downregulationat early time points. After 30 min of LPS treatment, expression of Mer was 61 ± 12% at time = 0 in the absence of Pros1 and 58 ± 15% in the presence of 25 nM Pros1. Similarly, after 60 min of LPS treatment, expression of Mer was 55 ± 8% at time = 0 in the absence of Pros1 and 56 ± 11% in the presence of 25 nM Pros1 (n = 3). The reduction of expression of Mer induced by LPS was similar in the presence or absence of Pros1, suggesting that differential reduction in Mer expression did not account for increased proinflammatory cytokine production in the presence of Pros1.

Although it is well established that loss of Mer signaling affects proinflammatory cytokine release by macrophages (45), there may be species- or cell type–specific effects in addition to temporal differences in responses that may be important. We found similar effects of Pros1 upon LPS-induced TNF release by human MDM, although we could not use the GC-MDM to further investigate the role of Mer as glucocorticoid treatment of macrophages strongly downregulates proinflammatory cytokine production. Thus, although treatment with glucocorticoid drives Mer-dependent phagocytosis, examination of the potential role of Mer signaling in regulation of TNF production is not possible. It is likely that changes in Mer expression during human monocyte–macrophage differentiation (49, 50) will also impact the engagement of downstream signaling pathways that drive pro- or anti-inflammatory signaling. Unlike human monocytes, mouse monocytes do not express Mer (50), and Pros1-dependent augmentation of TNF release may not be seen in mouse monocytes. Rothlin et al. (21) demonstrated that for bone marrow–derived dendritic cells, TAM-dependent suppression of LPS-induced IL-6 release was lost or reversed in the absence of signal transducer and activator of transcription-1 (STAT1) or IFNAR1 expression. In contrast, recent work from Earp and colleagues (51) has identified a new molecular mechanism by which exogenous Pros1 acts via Mer and Tyro3 to suppress macrophage polarization. In their studies, Mer formed a complex with protein tyrosine phosphatase 1B, acting to suppress intracellular signaling and proinflammatory gene expression. Interestingly, the suppressive effects of Pros1 upon some proinflammatory genes, including TNF and inducible NO synthase, were found to be less sensitive to Mer-dependent inhibition at early time points. The augmentation of TNF release by Pros1 we describe in this article may account for the early augmentation of TNF release observed following short periods of coculture of macrophages with apoptotic cells (52).

We suggest that TAMs could exert divergent regulatory effects upon monocyte/macrophage function during an inflammatory response in a manner that is dependent on the context of PtdSer exposure on the cell membrane (Fig. 6). During the initiation of inflammatory responses, there will be few apoptotic cells present at inflammatory sites. We speculate that when ligation of Mer (and Tyro3) by TAM ligands is uncoupled from apoptotic cell phagocytosis, downstream signaling may act to amplify, rather than suppress, LPS-induced monocyte proinflammatory mediator production. One possibility is that Pros1 binding to monocytes via PtdSer acts either in cis to engage TAMs on the same cell or in trans to TAMs expressed on adjacent monocytes and confers these effects. In contrast, as inflammation progresses, the presence of apoptotic cells with high-level opsonization by Pros1 would be predicted to promote TAM-dependent macrophage efferocytosis and engagement of anti-inflammatory signals that dampen proinflammatory cytokine release.

In summary, our data suggest that in the absence of apoptotic cells, Pros1 acts to amplify LPS-dependent proinflammatory cytokine production by monocytes, an effect that requires RTK activity and the presence of the PtdSer-binding Gla domain of Pros1. Our findings unveil potential new roles for TAMs in regulation of monocyte/macrophage function during progression of inflammation.

We thank our colleagues Chris Gregory, Chris Lucas (University of Edinburgh, Edinburgh), John Griffin (The Scripps Research Institute, La Jolla, CA), and Greg Lemke (Salk Institute for Biological Studies) for support and helpful discussions. We are grateful to Greg Lemke and Erin Lew (Salk Institute for Biological Studies) for the provision of Gas6 and ΔGla-Gas6. We acknowledge the facilities and support of staff in the Queen’s Medical Research Institute Flow Cytometry and Cell Sorting Facility (Shonna Johnston, Will Ramsay, and Mari Pattison).

The authors have no financial conflicts of interest.