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Research Article
Resolvin D1 stimulates efferocytosis through p50/p50-mediated suppression of tumor necrosis factor-α expression
Ha-Na Lee, Joydeb Kumar Kundu, Young-Nam Cha, Young-Joon Surh
Journal of Cell Science 2013 126: 4037-4047; doi: 10.1242/jcs.131003

Summary

Phagocytosis of apoptotic neutrophils, termed efferocytosis, is essential for the resolution of inflammation as it prevents the tissues surrounding the inflamed site from being exposed to the toxic contents of lytic cells. Resolvin D1 (RvD1), endogenously generated from docosahexaenoic acid during resolution of inflammation, is known to stimulate efferocytosis. However, the molecular mechanism underlying RvD1-mediated enhancement of efferocytosis remains largely unresolved. In the present study, murine macrophage-like RAW264.7 cells treated with lipopolysaccharide (LPS) exhibited markedly reduced efferocytic activity, but this was restored by co-incubation with RvD1. RvD1-induced restoration of the efferocytic activity appears to be mediated by downregulation of LPS-induced TNF-α expression. The inhibitory effect of RvD1 on LPS-induced TNF-α expression was associated with enhanced nuclear localization of p50/p50 homodimer and concomitant reduction of p65/p50 heterodimer accumulation in the nucleus. RvD1 triggered phosphorylation and proteasomal degradation of nuclear factor κB1 (NF-κB1) p105 to generate p50, which was subsequently translocated to the nucleus as a p50/p50 homodimer. Knockdown of NF-κB p50 abolished the ability of RvD1 to suppress TNF-α expression and also to restore efferocytosis, suggesting that the replacement of p65/p50 with p50/p50 homodimer in the nucleus is crucial for RvD1-mediated stimulation of efferocytosis. In a murine peritonitis model, intraperitoneal administration of RvD1 abolished the zymosan-A-induced TNF-α production, thereby stimulating efferocytosis. Taken together, these findings indicate that RvD1 expedites resolution of inflammation through induction of efferocytosis by p50/p50-homodimer-mediated repression of TNF-α production.

Introduction

Acute inflammation is an essential process in defending the host against infection. During inflammation, circulating neutrophils infiltrate the inflamed site to eliminate the injurious stimuli, and subsequently undergo apoptosis. The removal of apoptotic neutrophils terminates inflammation whereby tissue homeostasis can be restored. The clearance of apoptotic neutrophils by macrophages, the process termed ‘efferocytosis’, is an important step in preventing tissue necrosis and chronic inflammation, which can be caused by disgorgement of toxic contents from apoptotic neutrophils (Serhan and Savill, 2005; Vandivier et al., 2006; Serhan et al., 2007). The process of efferocytosis has been reported to be controlled actively, in part, by the endogenously generated chemical mediators or local autacoids, which stimulate the activity of pro-resolving macrophages. Resolvin D1 (RvD1) is one of the pro-resolving lipid mediators formed from docosahexaenoic acid (DHA; C22:6), a representative omega-3 fatty acid in sequential reactions catalyzed by 15-lipoxygenase (15-LOX) and 5-LOX (Hong et al., 2003). It has been reported that RvD1 limits infiltration of polymorphonuclear leukocytes (PMN), but enhances the infiltration of monocytes to the inflammatory site. Furthermore, in a murine model of peritonitis, intraperitoneal administration of RvD1 has been shown to increase the efferocytic activity of macrophages (Sun et al., 2007). However, the molecular mechanisms responsible for the enhancement of efferocytic activity of macrophages by RvD1 have not been fully clarified.

Tumor necrosis factor-α (TNF-α) is one of the major pro-inflammatory cytokines that stimulates the release of other mediators of inflammation, thereby inciting further inflammatory responses. Hence, prolonged or elevated production of TNF-α triggers chronic inflammation. Recent findings have demonstrated that TNF-α perturbs complete resolution of inflammation by inhibiting efferocytosis (McPhillips et al., 2007; Borges et al., 2009). The TNF-α expression is mainly under the regulation of nuclear factor-κB (NF-κB). The promoter region of the TNF-α gene (Tnfa) harbors four NF-κB-binding sequences (κB1, κB2, κB2a and κB3) (Baer et al., 1998). The NF-κB family is composed of either hetero- or homodimers of five subunit members. These include p65 (RelA), p105/p50 (NF-κB1), p100/p52 (NF-κB2), c-Rel and RelB. Whereas NF-κB, present predominantly as a p65/p50 heterodimer, transactivates a battery of pro-inflammatory genes including Tnfa, an atypical NF-κB species (p50/p50 homodimer) induces transcriptional repression of target genes (Barchowsky et al., 2000; Wessells et al., 2004).

In resting cells, NF-κB1 (p105/p50) exists as an inactive complex in the cytoplasm, where the C-terminal domain of p105 (alternatively known as IκBγ) masks the nuclear localization signal domain of p50 and hampers its nuclear translocation (Moorthy and Ghosh, 2003). During resolution of inflammation, proteasomal cleavage of the C-terminal portion of p105 generates p50, which then forms a p50/p50 homodimer (Salmerón et al., 2001). It has been reported that the p50/p50 homodimer forms a complex with Bcl3 and hastens the resolution of inflammation by inducing the transcription of anti-inflammatory genes (Singh and Jiang, 2004).

Because TNF-α acts as a mediator of inflammation partly by blocking efferocytosis, we investigated whether RvD1-induced efferocytosis and subsequent resolution of inflammation are mediated through the negative regulation of TNF-α. Here, we report that RvD1 potentiates efferocytic activity of macrophages through inhibition of TNF-α production by modulating the NF-κB signaling by two distinct mechanisms: (1) inhibition of the classical NF-κB (p65/p50) pathway, and (2) activation of an atypical NF-κB (p50/p50) pathway. Our study reveals that RvD1 induces predominantly the formation of p50/p50 homodimer, while it inhibits lipopolysaccharide (LPS)-induced activation of p65/p50 heterodimer in RAW264.7 macrophages. Thus, the pro-resolving effect of RvD1 is preferentially mediated through modulation of the NF-κB-TNF-α axis.

Results

RvD1 expedites the resolution of zymosan-A-induced murine peritonitis by suppressing TNF-α production

In zymosan-A-induced mouse acute peritonitis, the total leukocyte count in the peritoneal fluid normally reaches the maximum at about 12 hours (Bannenberg et al., 2005). However, the number of inflammatory leukocytes (especially PMNs) in zymosan-A-treated mouse peritoneal exudates sharply decreases during the resolution of peritonitis. In assessing its pro-resolving effect, RvD1 was given at the peak of peritoneal inflammation (12 hours after zymosan A injection). RvD1 dramatically stimulated the resolution of peritonitis by decreasing the total number of leukocytes in the peritoneal exudates, as compared to that observed in mice challenged with zymosan A alone (Fig. 1A). The reduced PMNs count, but not that of mononuclear cells, reflected the loss of total leukocytes in peritoneal exudates (Fig. 1B; supplementary material Fig. S1). Therefore, administration of RvD1 altered the cellular composition in peritoneal exudates with an increase in the proportion of mononuclear cells and a concomitant decrease in the number of PMNs. Whereas PMNs are the principal inflammatory cells that appear in the initial phase of inflammation, mononuclear cells are predominant during the resolution phase of inflammation. Hence, RvD1 accelerates resolution of zymosan-A-induced murine peritonitis. In addition to reducing the proportion of peritoneal neutrophils, RvD1 suppressed the secretion of pro-inflammatory cytokine TNF-α and IL-1β in the peritoneal exudates, whereas it stimulated the production of anti-inflammatory cytokine IL-10 (Fig. 1C; supplementary material Fig. S2). Intraperitoneal administration of TNF-α to mice reversed the pro-resolving effect of RvD1. Moreover, TNF-α prevented the RvD1-mediated decrease in accumulation of peritoneal leukocytes (especially PMNs; Fig. 1D; see also supplementary material Fig. S1D). Zymosan-A-injected mice co-treated with RvD1 and TNF-α retained the cellular composition, in terms of the proportion of monocytes and PMNs of the peritoneal exudates similar to that observed in mice given zymosan A alone (Fig. 1E). RvD1 accelerated the resolution of peritonitis in the zymosan-A-treated mice by increasing the proportion of mononuclear cells by up to ∼43%, at the expense of lowering the proportion of PMNs by ∼32%. However, when TNF-α was given, the ability of RvD1 to enhance the resolution of peritonitis was abolished (Fig. 1F).

Fig. 1.
Fig. 1.

RvD1 facilitates resolution of inflammation by inhibiting TNF-α production in zymosan-A-induced peritonitis. Mice were administered zymosan A (30 mg/kg) intraperitoneally for 12 hours followed by treatment with vehicle or RvD1 (300 ng). Some mice also received TNF-α (10 ng) intraperitoneally together with RvD1. Six hours later, peritoneal exudates were collected. (A,D) The number of total leukocytes in peritoneal exudates was determined. (C) The concentrations of TNF-α in the cell-free peritoneal lavage were measured by ELISA. (B,E,F) The proportions of mononuclear cells and PMNs in collected peritoneal exudates were determined by differential cell counting. Results are the means ± s.d. (n = 4), *P<0.05, **P<0.01, ***P<0.001.

Increased efferocytosis by RvD1 is mediated through suppression of TNF-α production

Compared with animals challenged with zymosan A alone, mice treated with RvD1 plus zymosan A showed an increase in the proportion of macrophages engulfing apoptotic PMNs (F4/80+Gr-1+). Thus, it is evident that RvD1 facilitates the clearance of apoptotic PMNs by macrophages. However, this RvD1-mediated enhancement of the efferocytic activity of macrophages was significantly diminished by treatment with TNF-α (Fig. 2A). We then examined the potential contribution of RvD1-mediated suppression of TNF-α production to macrophage efferocytosis in vitro and ex vivo. As a measure of in vitro efferocytic activity, murine macrophage RAW264.7 cells were allowed to engulf the FITC-stained apoptotic Jurkat T cells for 60 minutes. When RAW264.7 cells were pretreated with LPS, the ability of macrophages to take up apoptotic Jurkat T cells was suppressed, but the efferocytic ability was restored by RvD1 co-treatment. However, when TNF-α was added at 2 hours following the RvD1 treatment (the time when RvD1 effectively inhibited LPS-induced TNF-α expression; Fig. 3A), the RvD1-induced efferocytic activity was abolished (Fig. 2B; see also supplementary material Fig. S3A). These findings suggest that the increase of efferocytosis induced by RvD1 in macrophages challenged with LPS is mediated through downregulation of TNF-α expression. RvD1-induced restoration of efferocytosis was also confirmed by an ex vivo assay (Fig. 2C). To investigate whether TNF-α suppresses the efferocytic ability of macrophages by altering the expression of cell surface receptors, we examined the level of CD36, a scavenger receptor involved in efferocytosis. In RAW264.7 cells treated with LPS or TNF-α, CD36 expression was markedly decreased (Fig. 2D). The LPS-induced decrease in CD36 expression was restored by RvD1, but not in the presence of exogenous TNF-α. These findings suggest that LPS-induced TNF-α upregulation can suppress efferocytosis by downregulating the expression of the scavenger receptor on the macrophage surface, and that RvD1 reverses it.

Fig. 2.
Fig. 2.

RvD1-mediated repression of TNF-α production increases the efferocytic activity of macrophages in vivo and in vitro. (A) In the zymosan-A-induced peritonitis model, the proportion of macrophages with ingested PMN (F4/80+/Gr-1+) was determined by flow cytometry. Results are the means ± s.d. (n = 4) and expressed as a percentage increase of F4/80+/Gr-1+ macrophages. (B) Apoptosis of Jurkat T cells was induced by UVB (180 mJ/cm2) irradiation, followed by incubation for 8 hours. RAW264.7 cells treated with LPS (200 ng/ml), RvD1 (50 nM) and/or TNF-α (10 ng/ml) were co-incubated with FITC–annexin-V-stained-apoptotic Jurkat T cells for 1 hour. The number of macrophages engulfing apoptotic Jurkat T cells was determined by flow cytometry. The histogram shows the relative phagocytic index over control values (means ± s.d., n = 3); *P<0.05, **P<0.01. (C) For an ex vivo efferocytosis assay, peritoneal macrophages were treated with LPS, RvD1 and/or TNF-α for 4 hours, and co-incubated with apoptotic neutrophils for 1 hour. The engulfment of apoptotic neutrophils by macrophages was determined by immunostaining using anti-F4/80 (green; macrophage marker) and anti-Gr-1 (red; neutrophil marker) antibodies. A representative fluorescence micrograph shows macrophages (green) engulfing apoptotic neutrophils (red). (D) The cell surface expression of CD36 was analyzed by flow cytometry using FITC-conjugated anti-CD36 antibodies after treatment of RAW264.7 cells with LPS, RvD1 and/or TNF-α for 4 hours.

Fig. 3.
Fig. 3.

RvD1 suppresses LPS-induced TNF-α production by blocking the classical NF-κB pathway. (A) RAW264.7 cells were treated with LPS (200 ng/ml) in the absence or presence of RvD1 (10 or 50 nM) and harvested at 2 hours (top panel) or at the indicated time intervals (bottom panel). Semi-quantitative RT-PCR was conducted to measure Tnfa mRNA levels. The actin level was measured to ensure that equal amounts of mRNA were loaded. (B) Culture supernatants were collected at 4 hours after LPS or RvD1 treatment (50 nM), and TNF-α concentrations were measured by ELISA. Data are expressed as means ± s.d. (n = 3), ***P<0.001. (C) Nuclear protein from cells incubated with LPS or RvD1 for 30 minutes was prepared and incubated with the γ-32P-labeled oligonucleotides containing the NF-κB consensus motif. Protein–DNA complexes were separated from free probe by electrophoresis. (D) The RvD1-mediated suppression of the transcriptional activation of NF-κB was measured by the luciferase reporter gene assay. After overnight transfection with a luciferase reporter construct containing NF-κB response elements, cells were exposed to LPS or RvD1 for 30 minutes. Data are means ± s.d. (n = 3), ***P<0.001. (E) RAW264.7 cells were stimulated with LPS or RvD1 for 30 minutes. Localization of p65 in cytoplasm and nucleus was determined by western blot analysis. α-Tubulin and lamin B were used as cytoplasmic and nuclear markers, respectively. The histogram represents the relative level of nuclear translocated p65. Data are means ± s.d. (n = 3), ***P<0.001. (F,G) RAW264.7 cells were treated with RvD1 for 15 minutes in the presence or absence of LPS. The IKKβ kinase activity was measured by an immune complex kinase assay using GST-IκBα and [γ-32P]ATP. Immunoblot analysis was carried out to measure total IKKβ (F). Cell lysates were subjected to immunoblot analysis to measure the levels of phosphorylated IκBα and total IκBα (G). Data represent at least three independent experiments.

The inhibitory effect of RvD1 on LPS-induced TNF-α expression is mediated through inhibition of the classical NF-κB pathway

LPS-induced stimulation of Tnfa transcription in RAW264.7 cells was completely suppressed by co-treatment of 50 nM RvD1 (Fig. 3A). Likewise, the secretion of TNF-α from LPS-stimulated macrophages was reduced by RvD1 treatment (Fig. 3B). Since LPS-induced TNF-α expression is dependent primarily on the activation of the NF-κB, we assessed whether RvD1 could modulate NF-κB signaling. To confirm whether the LPS-induced overproduction of TNF-α was mediated by NF-κB, RAW264.7 cells were transfected with siRNA against p65, a functionally active subunit of NF-κB. As shown in supplementary material Fig. S4, LPS failed to upregulate TNF-α expression in p65 knockdown cells. We then examined whether the RvD1 could inhibit the LPS-driven NF-κB activation. The LPS-induced DNA binding (Fig. 3C) and transcriptional activity (Fig. 3D) of NF-κB was significantly inhibited by RvD1. Consistent with the inhibition of NF-κB transcriptional activity by RvD1, the nuclear translocation of p65 was markedly reduced by co-incubation with RvD1 in LPS-stimulated macrophages (Fig. 3E).

It is well established that IκBα is bound to p65 in physiological conditions and inhibits nuclear translocation of p65. In cells stimulated with LPS, IκBα is subjected to proteolytic degradation upon phosphorylation catalyzed by activated IKKβ. We noticed that RvD1 had an inhibitory effect on LPS-induced IKKβ phosphorylation and activity (Fig. 3F). As a consequence, phosphorylation and subsequent degradation of IκBα were substantially blocked by RvD1 co-treatment in LPS-stimulated macrophages (Fig. 3G). Taken together, these results suggest that RvD1 might block the LPS-induced activation of the classical NF-κB signaling pathway by suppressing IKKβ activation, and subsequently the phosphorylation and degradation of IκBα.

RvD1 blocked LPS-induced nuclear translocation of p65, but not p50

Although RvD1 inhibited the classical LPS-induced NF-κB activation pathways by blocking IKKβ-mediated IκBα phosphorylation/degradation and subsequently nuclear translocation and DNA binding of p65 (Fig. 3C–G), it failed to alter DNA binding and nuclear localization of p50 and degradation of its precursor p105 (Fig. 4A,B) under the same experimental conditions. Taken together, these results indicate that p50 protein can form a complex with either p65 or p50, but preferentially forms a p50/p50 homodimer in RvD1-treated cells because of a lack of p65 in the nucleus.

Fig. 4.
Fig. 4.

Signaling through p50/p50 homodimer is crucial for the inhibitory effects of RvD1 on LPS-induced TNF-α expression and restoration of efferocytosis. RAW264.7 cells treated with LPS in the absence or presence of RvD1 (50 nM) were harvested at 30 minutes. (A) p50 oligonucleotide binding in nuclear extracts was determined by the NF-κB TransAM assay. Data are means ± s.d. (n = 3), ***P<0.001 compared with the DMSO-treated group. NC, negative control; PC, positive control; Comp, competition. (B) Levels of p105 and p50 in the cytoplasm and nucleus were measured by western blot analysis. Histograms represent the relative expressions of p105 and p50 in cytoplasm, and p50 in the nucleus. Data are means ± s.d. (n = 3), *P<0.05, **P<0.01, ***P<0.001. (C) Cells were transfected with scrambled or nfκb1 (N-terminal) siRNA for 16 hours, and then LPS was treated in the absence or presence of RvD1 for an additional 2 hours. The mRNA levels of Tnfa p50 and actin were determined by RT-PCR. (D) RAW264.7 cells were transfected with scrambled or nfκb1 siRNA for 16 hours and the assay for efferocytosis was performed by incubating cells with FITC–annexin-V-stained-apoptotic Jurkat T cells for 1 hour after pre-incubation with LPS in the absence or presence of RvD1 for 4 hours. Representative flow cytometric dot plots demonstrating changes in the proportion of macrophages engulfing FITC–annexin-V-stained apoptotic Jurkat T cells are shown. Data are expressed as fold increases in the phagocytic index over the control values (means ± s.d., n = 3), **P<0.01, ***P<0.001.

p50/p50 formation by RvD1 accounts for inhibition of LPS-induced TNF-α, resulting in restoration of efferocytosis

To further investigate the role of nuclear p50/p50 homodimer in suppression of LPS-induced TNF-α, we utilized siRNA against nfκb1 (specifically targeting the p50 coding part). In p50 knockdown cells, Tnfa mRNA expression was not significantly induced despite LPS stimulation (Fig. 4C), because the DNA-binding capacity of p65 is compromised in the absence of the p50 subunit (Schmitz and Baeuerle, 1991). In our present study, RvD1 was found to lose its ability to suppress LPS-induced Tnfa mRNA expression in p50 knockdown cells (Fig. 4C), suggesting that not only inhibition of p65/p50 nuclear translocation but also facilitation of p50/p50 formation is essential for inhibition by RvD1 of LPS-induced transcriptional activation of TNF-α. Based on these findings, we speculate that the facilitated formation of p50/p50 homodimer and the repression of LPS-induced TNF-α expression by RvD1 are crucial for its induction of efferocytosis. In support of this assumption, the decline in the efferocytic activity of RAW264.7 cells caused by LPS stimulation was partially restored upon co-treatment with RvD1, but this pro-resolving effect of RvD1 was abolished in p50 knockdown cells (Fig. 4D; see also supplementary material Fig. S3B). Owing to weak induction of TNF-α in p50 knockdown cells, efferocytic activity of macrophages lacking p50 was not significantly attenuated upon stimulation with LPS alone. Thus, it is likely that the predominant formation of p50/p50 homodimer facilitated by RvD1 accounts for its restoration of efferocytosis in LPS-treated macrophages.

RvD1 increases production of the p50 subunit of NF-κB through p105 phosphorylation and degradation, thereby increasing nuclear translocation of p50/p50 homodimer

RvD1 treatment alone caused transiently increased nuclear translocation of p50 in RAW264.7 cells, as determined by immunoblot and immunocytochemical analyses (Fig. 5A,B). This was accompanied by a concomitant reduction in the cytoplasmic levels of p50 and its precursor p105. In contrast, RvD1 treatment did not cause any substantial translocation of p65 into the nucleus (Fig. 5A,B). Next, we determined whether RvD1-induced nuclear translocation of p50 occurred as a consequence of degradation of p105. Phosphorylation of NF-κB1 p105 on serine 927 is known to facilitate its proteolysis to produce p50 (Salmerón et al., 2001). RvD1 treatment time-dependently increased the phosphorylation of p105 at Ser 927 with concomitant reduction in the total p105 levels (Fig. 5C). This was accompanied by accumulation of p50. The full-length p105 expression plasmid, fused with the N-terminus of green fluorescent protein (GFP), was used to ascertain that p50 accumulation upon RvD1 treatment was derived from p105. With RvD1 stimulation, the level of ectopically expressed GFP-tagged p105 decreased, whereas GFP-tagged p50 markedly accumulated (Fig. 5D). These data clearly support that the p50 NF-κB subunit forming a homodimer upon RvD1 treatment is mainly derived from p105 degradation. To identify the kinase responsible for p105 phosphorylation and degradation, we measured the activation of some candidate mitogen-activated protein kinases (MAPK) after RvD1 treatment. Whereas RvD1 induced the phosphorylation of extracellular-signal related kinase 1/2 (ERK1/2) within 15 minutes, it did not change the phosphorylation of c-Jun-N-terminal kinase (JNK) and p38 MAPK (Fig. 5E). Treatment with U0126, a pharmacological inhibitor of ERK1/2, prevented RvD1-induced phosphorylation and degradation of p105, resulting in reduced accumulation of p50 (Fig. 5F). Moreover, treatment of RAW264.7 cells with the proteasome inhibitor MG132 prior to RvD1 stimulation completely blocked p105 degradation and consequently p50 generation, which was consistent with diminished nuclear translocation of p50 (Fig. 5G). Taken together, these findings suggest that RvD1 triggers proteasomal degradation of p105 through activation of ERK1/2 signaling.

Fig. 5.
Fig. 5.

RvD1-induced p105 degradation and concurrent p50 formation are dependent on proteasomal activity. (A) RAW264.7 cells were treated with RvD1 (50 nM) or vehicle for the indicated time periods, and both cytosolic and nuclear extracts were prepared. Levels of p105, p50 and p65 in the cytoplasm and nucleus were determined by western blot analysis. α-Tubulin and lamin B were measured to ensure separation of cytosolic and nuclear fractions, respectively. (B) RAW264.7 cells were treated with RvD1 for 30 minutes and nuclear translocation of p50 and p65 was determined by immunocytochemical analysis. (C) Total proteins isolated from RvD1-treated cells were subjected to immunoblot analysis for the measurement of phosphorylated p105 and total p105 as well as p50. Actin was used as an equal loading control for normalization. Histograms represent the relative levels of phosphorylated p105, total p105 and p50. Data are means ± s.d. (n = 3), *P<0.05, **P<0.01, ***P<0.001. (D) RAW264.7 cells were transfected with N-terminal GFP-tagged p105 vector, followed by incubation with RvD1 or vehicle for 30 minutes. Total protein isolated from cell lysates was subjected to immunoblot analysis for the measurement of p105 and p50 levels. (E) Lysates from RvD1-treated cells were subjected to western blot analysis. RvD1-induced activation of ERK, JNK and p38 was assessed using phospho-specific antibodies. (F) Cells were pre-incubated with the ERK inhibitor U0126 (10 µM) or DMSO for 2 hours and then treated with RvD1 for an additional 30 minutes. Levels of phosphorylated p105, total p105 and p50 were measured by western blot analysis. (G) Cells were pre-incubated with or without the proteasome inhibitor MG132 (10 µM) for 2 hours and then treated with RvD1 for 30 minutes. Levels of p105 and p50 in the cytoplasm and nucleus were measured by western blot analysis. Data represent at least three independent experiments.

Discussion

Efferocytosis is crucial for the successful resolution of acute inflammatory response. Prolonged pro-inflammatory insults suppress macrophage efferocytosis, leading to chronic inflammation (Vandivier et al., 2006). Several studies have demonstrated that LPS significantly inhibits the ability of mouse peritoneal macrophages to take up apoptotic neutrophils because of an overproduction of TNF-α (Michlewska et al., 2009; Feng et al., 2011). In agreement with these reports, our present study revealed that pro-inflammatory stimuli, such as zymosan A and LPS, suppressed efferocytosis through induction of TNF-α expression, and that blocking TNF-α production nullified LPS-mediated suppression of efferocytosis. In addition, TNF-α downregulated the expression of CD36, one of the principal scavenger receptors involved in efferocytosis, thereby suppressing the ability of macrophages to recognize and take up apoptotic neutrophils. TNF-α is recognized as one of the major cytokines responsible for chronic inflammation (Clark, 2007). We consider that TNF-α-mediated development of chronic inflammation is attributable, at least in part, to repression of efferocytosis.

Resolution of inflammation is an active and tightly regulated process controlled by anti-inflammatory and pro-resolving endogenous mediators, such as lipoxins and resolvins (Lawrence et al., 2002; Serhan, 2007). It has been reported that RvD1 inhibits Toll-like receptor-mediated activation of macrophages, limits infiltration of PMNs, enhances the recruitment of nonphlogistic monocytes and promotes the engulfment of apoptotic leukocytes by macrophages (Serhan and Chiang, 2008; Schif-Zuck et al., 2011). Although the role of RvD1 in the resolution of inflammation has been extensively investigated, the molecular events associated with RvD1-induced activation of efferocytosis are not clearly defined. Some studies have demonstrated that RvD1 stimulates phagocytosis via its receptor (e.g. ALX and GPR32) activation and M2 macrophage polarization (Krishnamoorthy et al., 2010; Titos et al., 2011). One of the salient features of our findings is that RvD1 regulates the efferocytic activity of macrophages by suppressing TNF-α production upon pro-inflammatory stimulation. In RvD1-treated macrophages, LPS-induced TNF-α production was impaired, resulting in restoration of efferocytosis. Moreover, addition of exogenous TNF-α prevented the restoration of efferocytosis by RvD1. Feng et al. reported that the blockade of TNF-α activity by use of a neutralizing antibody reversed the inhibitory effect of LPS on phagocytosis (Feng et al., 2011). This prompted us to determine whether RvD1 could counteract TNF-α activity. RvD1 restored the efferocytic ability of macrophages, which is prone to suppression by TNF-α (supplementary material Fig. S5). These findings suggest that the restoration of efferocytosis by RvD1 is likely to be related to suppression of pro-inflammatory TNF-α expression as well as its action.

The present study demonstrates that the RvD1-mediated inhibition of LPS-induced transcriptional activation of TNF-α RvD1-mediated involves modulation of at least two different NF-κB pathways; one being the suppression of nuclear translocation of p65/50 and the other being the preferential binding of p50/p50 homodimer (Gomez et al., 2005; Dai et al., 2007) to the κB consensus sequence present in the TNF-α gene promoter. p65/p50 heterodimer is known to be the predominant form of functionally active NF-κB with pro-inflammatory activity, whereas p50/p50 homodimer exerts anti-inflammatory and pro-resolving effects. p50/p50 homodimer is considered to compete with p65/50 heterodimer for DNA binding (Bohuslav et al., 1998; Ma et al., 2003). Unlike p65/p50 heterodimer, p50/p50 homodimer lacks the transactivation domain, and the binding of p50/p50 homodimer to DNA hence causes repression of NF-κB target gene expression. In the present study, RvD1 enhanced localization of p50 in the nucleus, while it suppressed dissociation from IκBα and concurrent nuclear translocation of p65. This led to an increased net nuclear accumulation of p50 and predominant formation of p50/p50 homodimer. Rather than simply stimulating the physical dissociation of p50 from the p65/p50 complex, RvD1 caused the nuclear accumulation of p50 by enhancing the degradation of p105. The proteolytic degradation of p105 is regulated by two pathways, a limited (processing to p50) and a complete degradation (releasing bound p50) (Moorthy and Ghosh, 2003). We note that p50 accumulation in the nucleus following RvD1 treatment is a consequence of the degradation of C-terminal of p105, indicating the limited processing of p105. The possibility that RvD1 could release a p50/p50 homodimer from an inactive p105/p50 heterodimer (a complete degradation of p105) was excluded by the results of the experiment utilizing GFP-tagged nfkb1. We found that GFP–p105 was mainly produced from GFP-tagged nfkb1 in normal conditions, but the processing of GFP–p105 to yield GFP–p50 was increased after RvD1 treatment. Signal-induced processing of p105 to p50 was found to be dependent on phosphorylation and proteasome-mediated degradation of IκBγ in RvD1-stimulated cells (Lawrence et al., 2001). We noted that RvD1-induced p105 phosphorylation on Ser 927, facilitated recruitment of the SCFβ-TrCP (β-transducin-repeat-containing protein) ubiquitin ligase complex (Cohen et al., 2001). Pretreatment with the proteasome inhibitor MG132 blocked RvD1-induced proteolysis of p105, indicating the involvement of 26S proteasomes in RvD1-induced degradation of IκBγ (C-terminal of p105). Besides stimulating conversion of p105 to p50, RvD1 exerted an inhibitory effect on LPS-induced IKKβ activity, which is responsible for degradation of IκBα.

In addition to RvD1, LPS stimulation also induced p105 degradation, nuclear translocation of p50 and formation of the p50/p50 homodimer, presumably as an adaptive cellular response to pro-inflammatory insult. However, p65/p50 heterodimer is initially predominant over p50/50 homodimer after LPS treatment, thereby provoking the pro-inflammatory state. Interestingly, LPS triggered transient nuclear translocation of p65 at this time, but p50 accumulation in the nucleus was sustained for up to 12 hours (supplementary material Fig. S6), allowing sufficient time for the resolution of inflammation. These results indicate that the binding preference of p50 monomer determines the inflammatory status.

Efferocytosis during resolution of inflammation is influenced by the surrounding environment in the inflamed site. At the onset of inflammation, pro-inflammatory cytokines, chemokines and lipid mediators are produced, and these molecules suppress nonphlogistic phagocytosis of apoptotic neutrophils by macrophages. In addition to TNF-α, prostaglandin E2, a representative pro-inflammatory mediator, is also known to inhibit phagocytosis by macrophages (Aronoff et al., 2004). However, at the late phase of inflammation, the balance of cytokines, chemokines and lipid mediators shifts towards anti-inflammatory and pro-resolving mediators, facilitating the termination of inflammation. For example, previously published data demonstrate that the anti-inflammatory cytokine IL-10 augments efferocytosis and stimulates resolution of inflammation (Michlewska et al., 2009). In this study, we also observed that the levels of pro-inflammatory cytokines including TNF-α and IL-1β were decreased, whereas the level of anti-inflammatory cytokine IL-10 was elevated, rendering macrophages active for efferocytosis. Therefore, lipid mediator class switching and the balance between pro-inflammatory and anti-inflammatory cytokines are key factors regulating macrophages to undergo efferocytosis.

In summary, RvD1-derived suppression of TNF-α expression is responsible for complete resolution of inflammation. As proposed in Fig. 6, RvD1 suppresses TNF-α expression and restores efferocytosis during resolution of inflammation through two distinct mechanisms. Even though TNF-α exerts the beneficial function by triggering the immune response to bacterial infection or other harmful stimuli, sustained production of TNF-α is implicated in the pathogenesis of a variety of human diseases, such as rheumatoid arthritis (Cavazzana et al., 2007), inflammatory bowel disease (El Mourabet et al., 2010), Alzheimer's disease (Swardfager et al., 2010) and cancer (van Horssen et al., 2006). These disorders are linked to imperfect resolution of inflammation, caused by disturbance of efferocytosis, which often results from overproduction of TNF-α. Therefore, timely blockade of TNF-α overproduction should be essential for resolution of inflammation and prevention of chronic inflammatory diseases. It is evident that endogenously produced RvD1, generated during resolution of inflammation, is one of the key molecules in the first line of cellular defense against persistent inflammatory responses. As exogenous administration of RvD1 stimulates efferocytosis by suppressing TNF-α overproduction, this molecule might have therapeutic potential in the management of chronic inflammatory diseases associated with impaired efferocytosis.

Fig. 6.
Fig. 6.

The proposed mechanisms underlying RvD1-mediated suppression of TNF-α expression and restoration of efferocytosis during resolution of inflammation. Sustained production of TNF-α at inflamed sites and subsequent inhibition of efferocytosis of apoptotic neutrophils can cause chronic inflammation. In our study, it has been demonstrated that RvD1 stimulates macrophages to engulf apoptotic neutrophils during resolution of inflammation by inhibiting TNF-α expression through two different NF-κB pathways: (1) suppression of nuclear translocation of p65/p50 by downregulating IKKβ activity, and (2) promotion of nuclear translocation of p50/p50 through p105 degradation.

Materials and Methods

Materials

RvD1 was purchased from Cayman Chemical Co. (Ann Arbor, MI, USA). LPS (Escherichia coli O111:B4) was obtained from Sigma-Aldrich (St Louis, MO, USA). Recombinant mouse TNF-α was produced by R&D systems (Minneapolis, MN, USA). Dulbecco's modified Eagle's medium (DMEM), RPMI 1640 and fetal bovine serum (FBS) were purchased from Gibco BRL (Grand Island, NY, USA). Primary antibodies against p105/p50, phospho-NF-κB p50, phospho-IKKα/β, IKKβ, IκB-α, ERK1/2, phospho-ERK (Tyr 204), phospho-JNK (Tyr 183/Tyr 185), JNK, phospho-p38 (Tyr 182) and p38 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA), and antibodies against p65 and phospho-IκB-α were obtained from Cell Signaling (Beverly, MA, USA). The anti-rabbit and anti-mouse horseradish peroxidase-conjugated secondary antibodies, and anti-lamin B1 were purchased from Zymed Laboratories (San Francisco, CA, USA).

Cell culture

RAW264.7 macrophages and Jurkat T cells were purchased from American Type Culture Collection (ATCC, Manassas, VA, USA). Cells were cultured in DMEM (for RAW264.7 cells) and RPMI 1640 (for peritoneal macrophages and Jurkat T cells) with 10% FBS, 100 µg/ml streptomycin and 100 U/ml penicillin in humidified 5% CO2 at 37°C.

Zymosan-A-induced peritonitis

Institute of Cancer Research (ICR) mice (8 weeks of age) were purchased from Central Lab Animal Inc. (Seoul, South Korea). All the animals were maintained according to the Institutional Animal Care Guidelines. Animal experimental procedures were approved by the Institutional Animal Care and Use Committee at Seoul National University. Zymosan A (30 mg/kg; Sigma, St Louis, MO, USA) was administered intraperitoneally 12 hours before giving DMSO, RvD1 (300 ng/mouse, intraperitoneally) alone or together with TNF-α (10 mg/mouse, intraperitoneally), and mice were sacrificed 6 hours later. Peritoneal leukocytes were harvested by washing with 3 ml of phosphate-buffered saline (PBS) containing 3 mM ethylenediaminetetraacetate (EDTA).

Total and differential leukocyte counts

Total peritoneal leukocyte counts were carried out using Turk's solution (0.01% Crystal Violet in 3% acetic acid) in a hematocytometer. For the differential count, peritoneal exudate cells were spun in a cytocentrifuge at 400 g for 5 minutes onto a slide and stained with Wright-Giemsa stain.

Efferocytosis assay

To assess the percentage of macrophages engulfing apoptotic PMNs in vivo, peritoneal exudate cells were exposed first to anti-mouse CD16/32 blocking antibody (eBioscience, San Diego, CA, USA) for 5 minutes and then labeled with the FluoroTag fluorescein isothiocyanate (FITC)-conjugated anti-mouse F4/80 antibody (eBioscience) for 20 minutes. The labeled cells were permeabilized for 10 minutes using 0.1% Triton X-100 and then labeled further for 20 minutes with PE-conjugated anti-mouse Gr-1 (Ly-6G) antibody (eBioscience). The proportion of macrophages containing neutrophils (F4/80+/Gr-1+) was determined by employing flow cytometry or immunocytochemistry. For an ex vivo efferocytosis assay, mouse peritoneal macrophages were incubated in six-well flat-bottomed microtiter plates for 24 hours. Non-adherent cells were collected and incubated for additional 24 hours to induce apoptosis. After washing with medium, adherent monolayer cells were co-incubated for 1 hour with apoptotic non-adherent cells, which were mostly composed of neutrophils. Peritoneal macrophages engulfing apoptotic cells were evaluated as described above.

To determine the in vitro efferocytic activity of macrophages, RAW264.7 cells were co-incubated for 1 hour with Jurkat T cells (stained with FITC-conjugated annexin V) undergoing apoptosis. To remove the non-engulfed apoptotic Jurkat T cells, RAW264.7 cells were washed three times with PBS and the proportion of RAW264.7 cells containing apoptotic Jurkat T cells (FITC-positive cells) was assessed by flow cytometry and fluorescent microscopy. Apoptosis of Jurkat T cells was induced by serum withdrawal and UVB (180 mJ/cm2) irradiation, followed by incubation for 8 hours at 37°C in an atmosphere of 5% CO2.

Measurement of TNF-α, IL-1β and IL-10

The concentrations of TNF-α, IL-1β and IL-10 in cell-free peritoneal lavage and in culture supernatant were determined by using mouse TNF-α, IL-1β and IL-10 ELISA kits (KOMA BIOTECH Inc., Seoul, Korea) according to the manufacturer's instructions.

Flow cytometry

RAW264.7 cells were fixed with 10% buffered formalin solution (20 minutes), washed in PBS twice, and blocked with 2% BSA in PBS (30 minutes). FITC-conjugated anti-CD36 antibodies (Abcam, Cambridge, UK), diluted 1∶50 in 2% BSA in PBS, were kept for 1 hour on ice. Cells were analyzed using FACSCalibur Flow Cytometer (BD, Franklin Lakes, NJ, USA).

Reverse transcription-polymerase chain reaction (RT-PCR)

Total RNA was isolated from RAW264.7 cells using TRIzol® reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's protocol. One microgram of total RNA was reverse transcribed by using reverse transcriptase of murine leukemia virus (Promega, Madison, WI, USA). PCR was carried out in a thermo-cycler using specific primers for Tnfa, p50 and actin. The primers employed were (forward and reverse, respectively): Tnfa, 5′-TGAACTTCGGGGTGATCGGTC-3′ and 5′-AGCCTTGTCCCTTGAAGAGAAC-3′; p50, 5′-ATGTTCACAGCCTTCCTCCC-3′ and 5′-GGTAAATCTCCTCCCCTCCC-3′; actin, 5′-AGAGCATAGCCCTCGTAGAT-3′ and 5′-CCCAGAGCAAGAGAGGTATC-3′. PCR products were resolved by 2% agarose gel electrophoresis, stained with ethidium bromide, and photographed under ultraviolet light.

NF-κB reporter gene analysis

RAW264.7 cells were grown to 80% confluency in six-well plates and transfected with 1 µg of the NF-κB reporter construct, along with 0.5 µg of pSVGal plasmid using LipofectAMINE 2000 (Invitrogen) in Opti-MEM medium (Gibco). After 24 hours of transfection, cells were treated with LPS or RvD1 for an additional 2 hours, and then lysed using the reporter lysis buffer (Promega). Luciferase assays were performed using 20 µl of cell extract and 100 µl of luciferin substrate (Promega), and the luciferase activity was measured using a luminometer (AntoLumat LB953, EG and G Berthold, Bad Widbad, Germany). The obtained luciferase activity was normalized by comparing with β-galactosidase activity, which was carried out according to the manufacturer's instructions (Promega β-Galactosidase Enzyme Assay System).

Protein extraction and western blot analysis

Cytosolic extracts were obtained by suspending the cells in hypotonic buffer A [10 mM hydroxyethyl piperazineethanesulfonic acid (HEPES; pH 7.9), 10 mM KCl, 2 mM MgCl2, 1 mM dithiothreitol (DTT), 0.1 mM EDTA and 0.1 mM phenylmethanesulphonylfluoride (PMSF)] and Nondiet P-40 solution (0.1%). The mixture was then centrifuged for 5 minutes at 12,000 g to obtain the cytosolic fraction, and the pellet was washed once with buffer A. To obtain the nuclear fraction, cell pellets were then suspended in hypertonic buffer C [50 mM HEPES (pH 7.9), 50 mM KCl, 300 mM NaCl, 0.1 mM EDTA, 1 mM DTT, 0.1 mM PMSF and 10% glycerol]. Whole cells extracts were prepared by suspending the cells directly in the RIPA lysis buffer [20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1 mM Na2EDTA, 1 mM ethylene glycol tetra-acetic acid (EGTA), 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na3VO4, 1 µg/ml leuptin, 1 mM PMSF] for 1 hour on ice and this was followed by centrifugation for 15 minutes at 12,000 g. Protein lysates (15 µg) were electrophoresed using sodium dodecyl sulphate (SDS)-PAGE and the separated proteins were transferred to polyvinyl difluoride (PVDF) membrane (0.22 µm thickness; Gelman Laboratory, Ann Arbor, MI, USA). To block the non-specific binding of proteins with primary antibodies, the blots were incubated in a 5% non-fat dry milk–PBST buffer [PBS containing 0.1% Tween-20] for 1 hour at room temperature. The membranes were then incubated with the primary antibody suspended in 3% non-fat milk PBST buffer overnight at 4°C. This was followed by washing with 1 PBST and incubation using appropriate secondary antibody coupled to horseradish peroxidase. Proteins tagged with specific primary antibodies were visualized with an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, Buckinghamshire, UK).

Electrophoretic mobility-gel shift assay (EMSA)

Customized double-stranded oligonucleotide containing the NF-κB binding domains was obtained from Promega, and 100 ng of the oligonucleotide was labeled with [γ-32P]ATP by employing T4 polynucleotide kinase and was purified on a Nick column (Amersham Pharmacia Biotech, Buckinghamshire, UK). The nuclear protein (10 µg) was mixed with 4 µl of concentrated incubation buffer [10 mM Tris-HCl (pH 7.5), 100 mM NaCl, 1 mM DTT, 1 mM EDTA, 4% glycerol and 0.1 mg/ml sonicated salmon sperm DNA] and the hypertonic buffer was added to make up the final volume 20 µl. After pre-incubation at room temperature for 15 minutes, the labeled oligonucleotide (400,000 cpm) was added to the nuclear fraction and incubation was continued for an additional 50 minutes at room temperature. To ensure the specific binding of the labeled oligonucleotide to nuclear protein, a competition assay was carried out with the excess amounts of unlabeled oligonucleotide. After the incubation, 0.1% Bromophenol Blue (2 µl) was added, and NF-κB–DNA complexes were separated from the unbound free probe by electrophoresis on 6% nondenaturing polyacrylamide gel in 1× TBE buffer [90 mM Tris base, 90 mM boric acid and 0.5 mM EDTA (pH 8.0)] at 140 V for 3 hours. Gels were dried and exposed to X-ray film. For the antibody supershift assay, 1 µl of antibody specific either for NF-κB p50 or p65 was incubated with the nuclear protein extract for 30 minutes on ice prior to addition of the reaction mixture containing radiolabeled nucleotide.

In vitro IKK activity assay

Precleared cytosolic extracts (200 µg) were subjected to immunoprecipitation using anti-IKKβ antibody overnight. 20 µl of protein-G–agarose beads (Santa Cruz Biotech) were then added to the mixture, which was rotated for 4 hours at 4°C. The immunoprecipitate thus obtained was suspended in 50 µl of reaction mix containing 47 µl 1× kinase buffer [25 mM Tris-HCl (pH 7.5), 5 mM glycerolphosphate, 2 mM DTT, 0.1 mM Na3VO4 and 10 mM MgCl2], 1 µg GST-IκBα substrate protein and 10 µCi [γ-32P]ATP and incubated at 30°C for 45 minutes. The kinase reaction was stopped by the addition of 2× SDS loading dye, boiled at 99°C for 5 minutes. The supernatant was subjected to SDS-PAGE analysis, and the gel was stained with Coomassie Brilliant Blue and destained with destaining solution (glacial acetic acid∶methanol∶distrilled water = 1∶4∶5, v/v). The destained gel was dried at 80°C for 1 hour and visualized by autoradiography.

Evaluation of DNA-binding activity of p50

The DNA-binding activity of p50 was measured in nuclear protein extracts (20 µg) by the TransAM™ NF-kB p50 protein assay (Active Motif, Carlsbad, CA, USA), an ELISA-based method designed to detect and quantify NF-κB p50 subunit activation. The assay was performed according to the manufacturer's protocol.

Construction of p105

The cDNA of p105 was PCR amplified using the primers 5′-CCGCTCGAGATGGCAGACGATGATC-3′ and 5′-CCGGAATTCCTAAATTTTGCCTTCAATAGG-3′. The PCR product was sub-cloned into the pEGFP-C3 vector, and was validated by sequence analysis.

Immunocytochemistry

RAW264.7 macrophages were seeded at 3×104 cells per well in an eight-chamber plate and cultured in serum-starved medium for 4 hours. The cells were incubated for 30 minutes in the absence or presence of RvD1 and then fixed with 10% buffered formalin solution (20 minutes). The cells were then washed in PBS (twice for 5 minutes each), permeabilized with 0.2% Triton X-100 (5 minutes), washed in PBS (twice for 5 minutes each), and blocked with 5% BSA in PBST (30 minutes). Polyclonal rabbit anti-p50 or -p65, diluted 1∶100 in 1% BSA in PBST, were applied overnight at 4°C. This was followed by washing cells in PBS (twice for 5 minutes each) and then incubation for 1 hour at room temperature with FITC-conjugated anti-rabbit IgG secondary antibody diluted at 1∶1000 in 1% BSA–PBST. After washing (twice for 5 minutes each), cells were treated with propidium iodide. The p50 and p65 signals were detected using a confocal microscope (Nikon, Tokyo, Japan).

Footnotes

  • Author contributions

    H.N.L. designed and performed the experiments, analyzed data and wrote the manuscript; J.K.K. helped with the animal study; Y.N.C. revised the manuscript; Y.J.S. discussed experimental designs and revised the manuscript. All authors discussed the data.

  • Funding

    This work was supported by a Global Core Research Center grant from the National Research Foundation, Republic of Korea [grant number 2012-0001184].

  • Supplementary material available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.131003/-/DC1

  • Accepted May 29, 2013.

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Research Article
Resolvin D1 stimulates efferocytosis through p50/p50-mediated suppression of tumor necrosis factor-α expression
Ha-Na Lee, Joydeb Kumar Kundu, Young-Nam Cha, Young-Joon Surh
Journal of Cell Science 2013 126: 4037-4047; doi: 10.1242/jcs.131003
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