Resolution of acute inflammation is an active process governed by specialized proresolving mediators, including resolvin (Rv)D2, that activates a cell surface G protein–coupled receptor, GPR18/DRV2. In this study, we investigated RvD2-DRV2–dependent resolution mechanisms using DRV2-deficient mice (DRV2-knockout [KO]). In polymicrobial sepsis initiated by cecal ligation and puncture, RvD2 (∼2.7 nmol/mouse) significantly increased survival (>50%) of wild-type mice and reduced hypothermia and bacterial titers compared with vehicle-treated cecal ligation and puncture mice that succumbed at 48 h. Protection by RvD2 was abolished in DRV2-KO mice. Mass spectrometry–based lipid mediator metabololipidomics demonstrated that DRV2-KO infectious exudates gave higher proinflammatory leukotriene B4 and procoagulating thromboxane B2, as well as lower specialized proresolving mediators, including RvD1 and RvD3, compared with wild-type. RvD2-DRV2–initiated intracellular signals were investigated using mass cytometry (cytometry by time-of-flight), which demonstrated that RvD2 enhanced phosphorylation of CREB, ERK1/2, and STAT3 in WT but not DRV2-KO macrophages. Monitored by real-time imaging, RvD2–DRV2 interaction significantly enhanced phagocytosis of live Escherichia coli, an action dependent on protein kinase A and STAT3 in macrophages. Taken together, we identified an RvD2/DRV2 axis that activates intracellular signaling pathways that increase phagocytosis-mediated bacterial clearance, survival, and organ protection. Moreover, these results provide evidence for RvD2-DRV2 and their downstream pathways in pathophysiology of infectious inflammation.
This article is featured in In This Issue, p.555
Inflammation is a protective response to defend the host against infection and injury (1). Ungoverned and excessive inflammation is an underlying pathology of many prevalent diseases, including cardiovascular diseases, diabetes, arthritis, and sepsis (2–4). Complete resolution of acute inflammatory responses was thought to be a passive process with dissipation or dilution of local chemoattractants and proinflammatory mediators, allowing tissues to return to homeostasis (1). In recent years, we obtained the first evidence, to our knowledge, that resolution of self-limited inflammation is not merely a passive termination, but rather an actively orchestrated programmed response that is rapidly turned on during acute inflammatory challenges, permitting inflamed and injured tissues to return via catabasis to function (5, 6). This event is driven in part by temporal lipid mediator (LM) class switching from generation of proinflammatory mediators (e.g., leukotriene [LT]B4) to the biosynthesis of lipoxins (LX) (7) and specialized proresolving mediators (SPM) (8–11). SPM are evolutionally conserved chemical structures derived from polyunsaturated fatty acids, including eicosapentaenoic acid–derived E-series resolvins and docosahexaenoic acid (DHA)–derived D-series resolvins, protectins, and maresins. Complete stereochemistries of these SPM are established and total organic synthesis achieved that also confirmed their potent proresolving actions. SPM are potent mediators governing not only innate immune responses in host defense, but also pain, organ protection, and tissue remodeling (recently reviewed in Ref. 8).
SPM derived from DHA including resolvin (Rv)D2 (7S,16R,17S-trihydroxy-docosa-4Z,8E,10Z,12E,14E,19Z-hexaenoic acid) were first identified and isolated from murine self-resolving exudates during the resolution phase of self-limited acute inflammation in vivo (12). The biosynthesis of RvD2 involves 17-lipoxygenation of DHA to 17S-hydroperoxy-DHA that is further transformed enzymatically to a 7(8)epoxide-containing intermediate in leukocytes via 5-lipoxygenase, followed by enzymatic hydrolysis to form RvD2. Endogenous RvD2 production is documented in human serum, plasma (13), adipose tissue (14), placenta (15), lung (16), breast milk (17), and sepsis patients (18). With isolated human polymorphonuclear neutrophils (PMN), RvD2 increases intracellular phagosomal reactive oxygen species generation for microbial killing (19). In whole blood at a single cell level using microfluidic chambers, RvD2 limits PMN chemotaxis and direct travel as well as increases random movement toward an IL-8 chemotactic gradient (20). RvD2 also decreases monocyte adhesion to adipocytes as well as their transadipose migration (14). RvD2 is a potent immunoresolvent that stereoselectively reduces excessive PMN trafficking in peritonitis and improves survival in sepsis (19). RvD2’s potent nanogram actions are also protective in disease models where RvD2 prevents inflammatory bowel disease such as colitis (21), alleviates inflammatory and fibromyalgia-induced pain (22, 23), increases survival following burn wound and reduces kidney and liver injuries in mice (24, 25), and reduces periodontitis (26) as well as nerve injuries as seen in Parkinson disease (27).
Resolution at the cellular level consists of cessation of PMN entry into the tissue and elevated efferocytosis (i.e., macrophage phagocytosis of apoptotic PMN) (1). We introduced a quantitative definition of resolution of self-limited sterile acute inflammation denoted resolution indices that permits assessment of the resolution properties of SPM and pinpoints their unique mechanisms of action (6). SPM each lower the magnitude of leukocyte infiltration and/or shorten the resolution interval, which is the interval from the time point of maximum PMN infiltration to the time point of 50% PMN reduction in peritoneal exudates (17, 28). We also assessed the actions of SPM in resolution of Escherichia coli infection using these resolution indices together with cellular composition and functions (29). During E. coli infection, RvD2 given at the peak of PMN infiltration (12 h) significantly shortens the resolution interval by half and enhances PMN apoptosis and macrophage efferocytosis in self-limited E. coli infection (30). Based on these potent proresolving actions of SPM, we proposed the “cardinal signs of resolution” including clearance of debris (expurgatio reliquiorum), clearance of infective agents (expurgatio contagionem agentis), analgesia (doloris absentia), and gain of function (muneris lucrum) (31). With these cellular, molecular, and quantitative definitions, we now appreciate the mechanisms controlling the active resolution programs of inflammation and have pinpointed the proresolving actions of SPM.
Polymicrobial sepsis is a complex scenario containing phases of both excessive inflammatory responses temporally associated with immunosuppressive states (32). Sepsis remains an unmet clinical challenge with high mortality rates and increasing incidence (33, 34). Recently, we identified a G protein–coupled receptor for RvD2 termed DRV2/GPR18 and demonstrated specific binding of RvD2 to human recombinant DRV2 (30). In human macrophages, RvD2 stimulates phagocytosis and efferocytosis in a DRV2-dependent manner. RvD2–DRV2 interaction stimulates phagocyte functions to accelerate resolution of bacterial infections. In the present study, we tested the hypothesis that the DRV2 receptor is pivotal in systemic infection and report that RvD2–DRV2 interactions stimulated bacterial clearance and reduced mortality in microbial sepsis. This ligand–receptor activation initiates specific intracellular signal transduction pathways (e.g., protein kinase A [PKA], STAT3) that contribute to RvD2-DRV2–stimulated phagocytosis of live E. coli.
Materials and Methods
Cecal ligation and puncture
Targeted deletion of mouse gpr18 (NM_182806) was constructed by Lexicon Pharmaceuticals in a 129/SvEv-C57B/6 mixed background (30). Male mice (10–12 wk old) were used for all experiments and fed ad libitum laboratory rodent diet 20-5058 (Lab Diet; Purina Mills). All experimental procedures used were approved by the Standing Committee on Animals of Harvard Medical School (protocol no. 02570) and complied with institutional and US National Institutes of Health guidelines. Cecal ligation and puncture (CLP) was carried out essentially as in Spite et al. (19). Mice were anesthetized with a mixture of oxygen and isoflurane 5% for anesthesia induction and 1.5% isoflurane for maintenance during surgery. After mice were shaved, basal temperature was monitored. A cut (0.5 cm) midline incision to the mouse’s left side was performed; cecum was pulled out and ligated below the ileocecal valve with a 4-0 silk suture. Two punctures were performed with a 20-gauge needle, followed by a soft squeeze until stool was extruded. Cecum was then put back to the peritoneal cavity and animals were sutured with a 6-0 silk suture. Mice then received 1 μg of RvD2 i.p. or 500 μl of saline with 0.1% ethanol (vehicle), and survival was monitored during 120 h. In a second group of animals, after 12 h of CLP, peritoneal exudates were collected by lavaging with 5 ml of PBS−/−. Peritoneal bacterial titers were determined by plating lavages on Luria–Bertani agar plates at 100× dilution, and the CFU were counted 24 h later. Exudate cytokine levels were determined by proteome profiler mouse cytokine array following the manufacturer’s instruction.
Liquid chromatography–tandem mass spectrometry-based metabololipidomics were performed with infectious exudates. Prior to sample extraction, ice-cold methanol containing deuterium-labeled d4-LTB4, d4-5S-HETE, d4-PGE2, and d5-RvD2 internal standards (500 pg each) were added to facilitate quantification. All samples were kept at −20°C for 45 min to allow protein precipitation and then subjected to solid-phase extraction as described (13). Extracted samples were analyzed by a liquid chromatography–tandem mass spectrometry system (QTRAP 5500; AB Sciex) equipped with an LC-20AD HPLC (Shimadzu, Tokyo, Japan). A Poroshell 120 EC-18 column (100 mm × 4.6 mm × 2.7 μm; Agilent Technologies, Santa Clara, CA) was kept in a column oven maintained at 50°C, and LM were eluted with a gradient of methanol/water/acetic acid from 55:45:0.01 (v/v/v) to 100:0:0.01 at 0.5 ml/min flow rate. To monitor and quantify the levels of targeted LM, a multiple reaction monitoring (MRM) method was devised with signature ion fragments for each molecule. Identification was conducted using published criteria including retention times and at least six diagnostic ions. Calibration curves were obtained using synthetic and authentic LM mixtures, including d4-LTB4, d5-LXA4, d4-PGE2, d5-RvD2, RvD1, RvD2, RvD3, RvD4, RvD5, PD1, MaR1, RvE1, RvE2, LXA4, LXB4, PGE2, PGD2, PGF2α, thromboxane (TX)B2, and LTB4 at 1.56, 3.12, 6.25, 12.5, 25, 50, and 100 pg. Linear calibration curves for each compound were obtained with r2 values of 0.98–0.99. Quantification was carried out based on peak areas of the MRM transitions.
Proteome profiler array
Peritoneal lavages were collected as described at 12 h after CLP. Cell-free supernatants were collected by centrifugation. A 1:10 dilution of the supernatant (150 μl) was incubated with the precoated Proteome Profiler array membranes (ARY028; R&D Systems) and processed according to the manufacturer’s instructions. Densitometric analysis of dot blots was performed using ImageJ software (National Institutes of Health, Bethesda, MD).
Real-time imaging of phagocytosis
Mouse bone marrow cells were collected and differentiated to macrophages with mouse GM-CSF (10 ng/ml) for 6 d. These macrophages were plated onto eight-well chamber slides (0.5 × 105 cells per well) overnight before the experiments. Chamber slides were kept in a Stage Top incubation system for microscopes equipped with a built-in digital gas mixer and temperature regulator (Tokai Hit model INUF-K14). Cells were treated with RvD2 (10 nM), PKA inhibitor H89 (3 μM; Sigma-Aldrich), RvD2 plus H89, or vehicle control for 15 min at 37°C, followed by addition of BacLight Green–labeled E. coli at a proportion of 50:1 (E. coli/macrophage) to initiate phagocytosis. Fluorescent images were then recorded every 10 min for 100 min (37°C) with Keyence BZ-9000 (Biorevo) inverted fluorescence phase-contrast microscope (×20 objective) equipped with a monochrome/color switching camera using BZ-II Viewer software (Keyence). Three separate experiments were performed. In each experiment, three fields (×20) per condition (per well) were recorded. Green fluorescence intensity was quantified using a BZ-II analyzer.
For cAMP measurements, resident peritoneal macrophages from naive WT or DRV2-KO mice were collected with 5 ml of PBS−/− and plated in a 12-well plate (0.5 × 106 cells per well) with RPMI 1640 supplemented with 10% FBS. The next day, media were replaced with PBS+/+ and cells incubated with RvD2 (0.1–100 nM), forskolin (10 μM; Sigma-Aldrich), or vehicle control for 15 min at 37°C. Lysis buffer (100 μl) was added to stop incubation and cells were homogenized. cAMP levels were measured by ELISA following the manufacturer’s instruction (Elite cAMP ELISA assay kit; eEnzyme, CA-C315).
Macrophage phagocytosis of E. coli
Resident peritoneal macrophages were collected from naive WT and DRV2-KO mice and plated onto 96-well plates (0.5 × 105 cells per well). RvD2 (1 pM to 10 nM) or vehicle controls were incubated with macrophages for 15 min at 37°C, followed by incubation with FITC-labeled serum-treated zymosan particles (STZ) at 10:1 ratio (zymosan/macrophage) or BacLight Green–labeled E. coli at a 50:1 ratio (E. coli/macrophage) for 60 min at 37°C. Plates were gently washed, extracellular fluorescence was quenched by trypan blue, and phagocytosis was determined by measuring total fluorescence (excitation 493/emission 535 nm) using a fluorescent plate reader (Molecular Probes). The following inhibitors were added together with RvD2 or vehicle control: PKA inhibitor H89 (3 μM; Sigma-Aldrich), pERK1/2 inhibitor (50 μM; Tocris Bioscience), and STAT3 inhibitor (100 μM; Tocris Bioscience)
Peritoneal macrophages from DRV2-KO mice and WT mice were collected as in Chiang et al. (30). Peritoneal cells were incubated with RvD2 (10 nM) for 0, 1, 5, 15, and 30 min at 37°C, followed by 1.6% paraformaldehyde for 10 min at room temperature. Cells were barcoded following the manufacture’s protocol with palladium isotopes (Pd 102, 104, 105, 108, and 110) (Fluidigm Sciences). Briefly, cells were washed twice using barcoding permeabilization buffer. Diluted barcodes were transferred to cells and incubated for 30 min at room temperature. Barcoded cells were washed twice in cytometry by time-of-flight (CyTOF) staining buffer (PBS with 0.5% BSA and 0.1% sodium azide) and then pooled for staining. Pooled barcoded cells were incubated for 10 min with Fc Block (BioLegend, San Diego, CA) for Fc receptor–mediated nonspecific Ab binding. Cells were then stained for 30 min with metal-labeled surface Abs at room temperature, then washed twice in CyTOF staining buffer. Cells were then permeabilized in 80% ice-cold methanol for 10 min at −20°C. After washing twice to remove the methanol, cells were stained with metal-conjugated Abs for intracellular markers at room temperature for 30 min. The Abs used for CyTOF are listed in Supplemental Table III. Cells were then washed twice and stained in 500 μl of 1:1000 iridium intercalator (DVS Sciences, Toronto, ON, Canada) diluted in PBS overnight at 4°C. Cells were then washed twice in CyTOF staining buffer and twice in MilliQ-filtered deionized water (35). Cells were reconstituted at a concentration of 5 × 106 cells/ml containing EQ calibration beads (EQ four elements calibration beads; Fluidigm Sciences) according to the manufacturer’s protocol. Barcoded cells were analyzed on a Helios CyTOF (Fluidigm Sciences) at an event rate of 400–500 cells/s. The data were normalized using v6.3.119 Helios software (Fluidigm Sciences) at the LMA CyTOF facility at Dana Farber Cancer Institute (Boston, MA). Files were debarcoded using a Fluidigm debarcoder application. Gating was performed in a Cytobank platform (Cytobank, Mountain View, CA). Sequential gating strategy was used to analyze signaling events in resident peritoneal macrophages (CD11b+F4/80+). Phosphorylation levels of signaling molecules were determined at 0, 1, 5, 15, and 30 min after exposure of RvD2 (10 nM) in WT and DRV2-KO macrophages. Phosphorylation levels were calculated as the difference between the inverse hyperbolic sine of the median signal intensity at indicated time points and the inverse hyperbolic sine of the median signal intensity in the unstimulated (0 min) signal (36).
Statistical analysis was performed using a Student t test (two-group comparisons) and one-way ANOVA (multiple-group comparisons). Kaplan–Meier survival curves were analyzed using a one-tailed log-rank (Mantel–Cox) test. In all cases a p < 0.05 was considered significant.
RvD2 protects mice from sepsis in a GPR18/DRV2 receptor–dependent manner
Because RvD2 displays potent actions and RvD2 receptor, namely GPR18/DRV2, was identified (30), we set out to investigate whether the RvD2/DRV2 receptor axis is protective in polymicrobial sepsis. Using mice deficient in DRV2, we carried out CLP, a polymicrobial systemic sepsis model that closely resembles human pathology (37). In WT animals, RvD2 significantly increased survival (>50%) in CLP compared with vehicle-treated mice that all perished before 48 h (Fig. 1A, upper panel). In contrast, in DRV2-deficient mice (DRV2-KO), there were no significant differences in survival between RvD2- and vehicle-treated mice (Fig. 1A, bottom panel). We noted that with vehicle-treated CLP mice, DRV2-KO mice succumbed earlier (12 h) than did WT mice (24 h). Both WT and KO mice reached ∼50% mortality between 24 and 30 h. At 48 h the remaining WT mice perished whereas KO mice had 20% survival. A log-rank test comparing the survival distributions of these two groups (WT CLP plus vehicle versus KO CLP plus vehicle) did not reach statistical significance (p = 0.87). It is possible, however, that there was genetic and/or functional compensation in DRV2-KO mice, for example, expressions of other proresolving receptors and/or proinflammatory receptors might be altered in DRV2-KO animals. These could contribute to the apparent different outcome than that with WT mice survival following CLP.
We monitored body temperature at 24 h after CLP, which induced hypothermia in mice compared with naive mice. RvD2 reduced hypothermia (35.5 ± 0.8°C) compared with vehicle controls (31.7 ± 1.2°C) in WT mice but not in DRV2-KO mice (Fig. 1B). Additionally, in WT mice RvD2 enhanced bacterial killing, significantly reducing bacterial titers (105.5 ± 32.3 CFU/cm2) compared with the vehicle-treated group (234.8 ± 82.3 CFU/cm2), a response diminished in DRV2-KO mice (Fig. 1C). Thus, RvD2–DRV2 interactions in vivo increased survival in CLP mice, protected hypothermia, and enhanced bacterial clearance from infectious exudates.
LM metabololipidomics and proteome profiling
We next questioned whether DRV2-KO gives heightened inflammatory status in sepsis, and carried out mass spectrometry–based metabololipidomics focusing on local acting LM. Each LM was profiled using multiple reaction monitoring and identified by direct comparison with synthetic and authentic standards using matching criteria, including retention time, characteristic fragmentation patterns, and at least six diagnostic ions (13). In infectious exudates obtained from CLP mice, we identified resolvins and other SPM, specifically eicosapentaenoic acid–derived E-series resolvins, DHA-derived D-series resolvins, protectins, and maresin, as well as arachidonic acid–derived LT, prostanoids, and LX (Fig. 2A, Supplemental Table I). A representative tandem mass spectrometry spectrum of RvD1 used for identification is shown in Fig. 2B. Principal component analysis was used for exploring cross-covariance between WT and DRV2-KO. The three-dimensional loading plot showed two distinct clusters, one with mostly proinflammatory and procoagulating LM (i.e., PG, LT, and TX) that was associated with DRV2-KO. The other cluster contained proresolving mediators, for example, RvE1–3, RvD1–5, PD1, MaR1, and LXA4, that were associated with WT animals (Fig. 2C). We quantified each LM and found significant increases ∼85% in proinflammatory and procoagulating LM, for example, PG plus TX (Fig. 2D) and LTB4 plus 5-HETE (∼95% increase; Fig. 2E) in DRV2-KO compared with WT mice. Additionally, DRV2-KO mice showed significant decreases of ∼60% in D-series resolvins, including RvD1, AT-RvD1, RvD3, and AT-RvD3, compared with WT littermates (Fig. 2F).
Next, we carried out proteome profiling for CLP. Cell-free supernatants from infectious exudates showed significant upregulation of a panel of cytokines compared with naive mice at 12 h with both WT and DRV2-KO mice (Fig. 3A). RvD2 treatment significantly upregulated a panel of proteins, including matrix metalloproteinase (MMP)-2, MMP-3, and myeloperoxidase in WT (Fig. 3B, Supplemental Table II). However, these cytokines were not significantly altered in DRV2-KO by RvD2 treatment. It was reported that loss of MMP-2 leads to increases in MCP-3 levels and exacerbates myocarditis in mice, pointing to a potential protective role of MMP-2 via its function in chemokine cleavage (38).
Additionally, we carried out statistical analysis between WT-RvD2 and KO-RvD2 groups and found statistically significant reduction in several proteins in KO mice compared with WT mice. These include IL-22, LIX (CXCL5), MMP-3, MMP-9, Pentraxin 2/SAP, Pentraxin 3/TSG-14, Reg3G, and Serpin F1 (Supplemental Table II). These proteins have been reported to have protective roles in regulating immune responses during infections. For example, IL-22 controls pathobionts via regulation of the complement system, promoting resistance after pathogen-induced intestinal damage (39). Reg3G restricts bacterial colonization of mucosal surfaces and reduces bacterial translocation (40). Pentraxin 3/TSG-14 improves survival in endotoxic shock and CLP (41). Therefore, reduction in these proteins in KO-RvD2 mice might contribute to their higher mortality rate and impaired bacterial clearance compared with the WT-RvD2 group (Fig. 1A, 1C).
Taken together, our results demonstrated that DRV2-KO mice showed dysregulated LM profiles, giving heightened proinflammatory LM and reduced SPM compared with WT in sepsis. Also, RvD2 regulates a selected panel of inflammation-related proteins in a DRV2-dependent manner.
RvD2-DRV2–dependent intracellular signaling
Next, we investigated RvD2-DRV2–initiated intracellular signals using mass cytometry (CyTOF) with naive mouse peritoneal macrophages. Macrophages collected from WT and DRV2-KO mice were incubated with RvD2 (10 nM) for 0–30 min, and cells were collected for staining with specific Abs for cell surface markers, followed by permeabilization and staining of intracellular targets, including a panel of phospho-proteins (see 2Materials and Methods). Macrophages were identified as CD11b+F4/80+ populations (see the gating strategy in Supplemental Fig. 1). RvD2 time-dependently increased phosphorylation of pAKT, p-p38 MAPK, pCREB, pS6, pERK1/2, pSTAT1, pSTAT3, and pSTAT5, each with different kinetics in macrophages. pAKT, p-p38 MAPK, pCREB, and pS6 levels reached maximum at 1 min and then gradually declined. pERK1/2 was detected at 1 min and peaked at 30 min after RvD2 stimulation (Fig. 4A). In comparison, pSTAT1, pSTAT3, and pSTAT5 were also markedly increased as early as 1 min, and their levels remained elevated until 15–30 min. In macrophages from DRV2-KO mice, upregulation of these phospho-proteins by RvD2 was abolished. In separate sets of experiments, pCREB levels were validated using flow cytometry (Fig. 4B). Quantification of pERK1/2, pSTAT3, and pCREB using CyTOF and flow cytometry are shown in Fig. 4C. These results indicate that RvD2 regulates phosphorylation of selected kinases and transcription factors with different kinetics in macrophages in a DRV2-dependent manner.
STAT3 and PKA pathways mediates RvD2-DRV2–stimulated phagocytosis
Next, we investigated the role of selected signaling components in RvD2-DRV2–stimulated phagocytosis identified using CyTOF, including PKA (that phosphorylates CREB), STAT3 (that is involved in phagosome maturation), and ERK. First, we monitored phagocytosis of live fluorescence-labeled E. coli by bone marrow–derived macrophages (BMDM) using real-time imaging. Fluorescence intensity that increased with time (0–120 min) represents increased macrophage ingestion of E. coli (Fig. 5A). RvD2 addition prior to E. coli (10 nM, 15 min) significantly enhanced phagocytosis (∼95% at 100 min; Fig. 5A). This RvD2 action was not observed in DRV2-KO BMDM, indicating the role of the RvD2/DRV2 axis in stimulating bacterial clearance via phagocytosis. Because cAMP activates PKA, which can phosphorylate pCREB (42), we determined cAMP levels with naive peritoneal resident macrophages and found that RvD2 (10–100 nM) significantly increased cAMP levels in WT but not DRV2-KO (Fig. 5B).
A suboptimal concentration of forskolin (10 μM) was used as a control and taken as 100% (Fig. 5B). This is consistent with earlier findings that RvD2 (10–100 nM) increases cAMP levels that are dependent on DRV2 in human macrophages (30).
We next questioned whether PKA, STAT3, and ERK1/2 play a role in RvD2-stimulated phagocytosis. RvD2 incubation prior to E. coli addition (10 nM, 15 min) significantly enhanced phagocytosis. Incubation of macrophages with a STAT3 inhibitor (NSC 74859; 100 μM) or PKA inhibitor (H89; 3 μM) together with RvD2 significantly reduced RvD2-stimulated phagocytosis (Fig. 5C). Similar results were obtained with BMDM, where H89 also reduced RvD2-enhanced phagocytosis of live E. coli. In comparison, an ERK inhibitor (FR 180204; 10 μM) did not significantly change RvD2-stimulated phagocytosis (Fig. 5C). Macrophage phagocytosis of STZ was also carried out. STAT3 and PKA inhibition, but not ERK inhibition, significantly blocked RvD2-stimulated phagocytosis (Fig. 5D). Taken together, our results demonstrate that DRV2 contributes to RvD2’s proresolving actions in macrophage clearance of live E. coli and this action is dependent on cAMP/PKA and STAT3 signaling pathways (Fig. 6).
RvD2 is a potent immunoresolvent and controller of leukocyte traffic and is protective in a wide range of disease models, including airway and gastrointestinal inflammation (8, 19). In the present study, we demonstrated that RvD2–DRV2 interaction protected mice from sepsis, preventing hypothermia, enhancing phagocytosis-based bacterial clearance, and increasing survival. In infectious exudates collected from sepsis, DRV2-KO gave increased levels of PG, LT, and TX, and reduced SPM (i.e., RvD1, RvD3, AT-RvD1, AT-RvD3) in infectious exudates (Fig. 2). These results are consistent with those found in E. coli peritoneal infections where selected SPM, including AT-RvD1, RvD2, RvD5, PD1, and AT-PD1, were significantly reduced in DRV2-KO mice compared with WT mice. DRV2-KO also gave increased amounts of TX (30), indicating that DRV2-KO is associated with heightened inflammatory status during bacterial infection. Of interest, mice deficient in an RvD1 receptor, namely ALX, also gave heightened disease severity, including hypothermia and cardiac dysfunction during sepsis (43). Conversely, ALX transgenic mice (i.e., mice overexpressing human ALX) gave reduced inflammatory status with markedly decreased PMN infiltrates in peritonitis (44). Along these lines, mice overexpressing the human RvE1 receptor, namely ChemR23/ERV1, also showed lower PMN infiltration in peritonitis and, additionally, reduced ligature-induced alveolar bone loss (reviewed in Ref. 8). Taken together, these results point to an endogenous role of SPM receptors (e.g., ALX, ERV1, and DRV2) as checkpoint controllers of resolution during inflammation and bacterial infections.
We monitored a panel of phospho-proteins using CyTOF. These include transcription factors CREB, STAT1, STAT3, and STAT5, and kinases such as ERK1/2, p38 MAPK, and Akt (Fig. 4A). We found that RvD2 significantly increased phosphorylation of CREB in WT but not DRV2-KO macrophages (Fig. 4A), suggesting that the RvD2/DRV2 axis could regulate a panel of CRE-containing genes in a PKA-dependent manner (42). In mouse macrophages, RvD2 enhanced cAMP levels in a DRV2-dependent manner (Fig. 5B). Additionally, cAMP-activated protein kinase (PKA)–mediated RvD2-stimulated phagocytosis (Fig. 5C, 5D). Earlier we reported that RvD2 activates recombinant human DRV2 that is sensitive to cholera toxin, suggesting receptor coupling to a Gαs-like protein. Also, in human macrophages, RvD2 does-dependently increases cAMP, which was abolished when DRV2 was knocked down using specific shRNA (30). Thus, the present results together with earlier findings indicate that RvD2–DRV2 interactions initiate Gαs protein coupling, leading to activation of the cAMP/PKA signaling pathway that enhanced macrophage phagocytosis (Fig. 6). Activation of PKA by cAMP enhances Rac1 activity, leading to increased efferocytosis with peritoneal resident macrophages (45). Also, a cAMP analog activates EPAC (a GTP/GDP exchange factor), which in turn increases the levels of GTP-Rap1, leading to F-actin formation and enhanced phagocytosis of STZ in RAW264.7 murine macrophages (46). These components, namely Rac1, EPAC, Rap1, and F-actin, might also contribute to RvD2-stimulated phagocytosis of E. coli and STZ reported in the present study (Fig. 5C, 5D). In this context, RvD1 rescues macrophages from oxidative stress–induced apoptosis during efferocytosis, suppressing activation of NADPH oxidase via cAMP–PKA signaling. This action is dependent on ALX (47). Thus, it is likely that the Gαs–cAMP–PKA signaling cascade in macrophages is a shared proresolving mechanism for RvD1 and RvD2 receptors.
RvD2–DRV2 interactions also increased phosphorylation of STAT3, STAT5, ERK1/2, Akt, and ribosomal protein S6 (Fig. 4). S6 protein is a known downstream target of PI3K/Akt signaling pathway and the Raf/ERK pathway (48). In the case of RvE1, the ERK/Akt/S6 pathway is involved in RvE1-stimulated phagocytosis in an ERV1-dependent manner (49). Additionally, RvD1 and RvD2 increased phosphorylation of Akt and CREB in LPS-stimulated human monocytes (50). Inhibition of STAT3 phosphorylation decreases efferocytosis and M2 macrophage polarization in vitro (51). Additionally, RAW264.7 murine macrophage phagocytosis of Staphylococcus aureus leads to upregulation of genes involved in the JAK/STAT pathway, including STAT3 and STAT5. Phosphorylation of STAT3 and STAT5 proteins is required for phagosome acidification and maturation (52). Thus, it is likely that STAT3 phosphorylation is also required for phagosome maturation during human macrophage phagocytosis. Disruption of this pathway with STAT3 inhibition diminished RvD2-enhanced phagocytosis of E. coli and STZ (Fig. 5C, 5D). Along these lines, RvD1 upregulates STAT3 in human monocytes (53). Taken together, these results indicate that each SPM stereoselectively activates its own specific receptor (e.g., RvD2-DRV2, RvD1-DRV1, and RvE1-ERV1). JAK/STAT and ERK/Akt/S6 are likely to be the common downstream pathways following receptor activation by SPM that are involved in SPM-initiated phagocytosis.
In summary, we provide evidence for an RvD2/DRV2 resolution axis that protects microbial sepsis, increasing bacterial clearance and improving survival. Additionally, we identified DRV2 receptor–dependent signaling pathways in macrophages. Our present results suggest that these signaling components and pathways could represent “resolution signaling cassette/complex” in phagocytosis. Earlier reports established that other SPM activation of their receptors also mediate proresolving actions. For example, RvD1 regulates a panel of select micro-RNAs and their target genes involved in resolution of inflammation, including IκB kinase, IL-10, and 5-lipoxygenase in an ALX- and DRV1/GPR32-dependent manner (53). RvE1-ERV1 enhanced phagocytosis via the Akt/S6 pathway (49). Resolvins and the other proresolution mediators from n-3 fatty acids are agonists of the resolution response and are conserved structures across the animal kingdom (8). Taken together, these findings provide new proresolution mechanisms of the host that may be of interest, providing potential therapeutic opportunities for the control of unwanted inflammation and infection that accompanies sepsis and infectious inflammatory diseases. Thus, SPM and their receptors and signaling pathways documented in this study provide examples of the potential for resolution pharmacology.
We thank Mary Small for assistance with manuscript preparation.
This work was supported in part by National Institutes of Health Grants R01 GM38765 (to C.N.S.) and R01 GM38765-29S1 (to S.L.).
The online version of this article contains supplemental material.
Abbreviations used in this article:
bone marrow–derived macrophage
cecal ligation and puncture
cytometry by time-of-flight
multiple reaction monitoring
protein kinase A
resolvin D2 (7S,16R,17S-trihydroxy-docosa-4Z,8E,10Z,12E,14E,19Z-hexaenoic acid)
specialized proresolving mediator
serum-treated zymosan particle
The authors have no financial conflicts of interest.