ABSTRACT
The liver X receptors (LXRs) are ligand-activated nuclear receptors with established roles in the maintenance of lipid homeostasis in multiple tissues. LXRs exert additional biological functions as negative regulators of inflammation, particularly in macrophages. However, the transcriptional responses controlled by LXRs in other myeloid cells, such as dendritic cells (DCs), are still poorly understood. Here we used gain- and loss-of-function models to characterize the impact of LXR deficiency on DC activation programs. Our results identified an LXR-dependent pathway that is important for DC chemotaxis. LXR-deficient mature DCs are defective in stimulus-induced migration in vitro and in vivo. Mechanistically, we show that LXRs facilitate DC chemotactic signaling by regulating the expression of CD38, an ectoenzyme important for leukocyte trafficking. Pharmacological or genetic inactivation of CD38 activity abolished the LXR-dependent induction of DC chemotaxis. Using the low-density lipoprotein receptor-deficient (LDLR−/−) LDLR−/− mouse model of atherosclerosis, we also demonstrated that hematopoietic CD38 expression is important for the accumulation of lipid-laden myeloid cells in lesions, suggesting that CD38 is a key factor in leukocyte migration during atherogenesis. Collectively, our results demonstrate that LXRs are required for the efficient emigration of DCs in response to chemotactic signals during inflammation.
INTRODUCTION
Dendritic cells (DCs) represent a heterogeneous population of professional antigen-presenting cells (APCs) that arise from the bone marrow (BM) and reside in peripheral tissues and lymphoid organs (1, 2). DCs play central roles in the initial host recognition of pathogens and in the induction of antigen-specific adaptive immune responses. As sentinels located in peripheral tissues, immature DCs (iDCs) express a plethora of pattern recognition receptors and exhibit a high endocytic capacity. Upon the recognition and capture of microbial products, DCs undergo a maturation process characterized by the upregulation of costimulatory molecules and proinflammatory cytokines (3). Furthermore, mature DCs (mDCs) exhibit increased expression of the chemokine receptor CCR7, a G-protein-coupled receptor critical for DC migration from peripheral tissues to lymphoid organs (4, 5). Thus, the proper maturation and emigration of DCs from injured tissues are crucial for the initiation of antigen-dependent immune responses.
The liver X receptors (LXRs) (LXRα and LXRβ, encoded by Nr1h3 and Nr1h2, respectively) are ligand-activated transcription factors that belong to the nuclear receptor superfamily. LXRs function as key regulators of lipid homeostasis by controlling the expression of several genes that are pivotal for cholesterol, fatty acid, and phospholipid metabolism (6–8). Both LXRs form obligate heterodimers with retinoid X receptors (RXRs) and positively regulate the expression of target genes through direct binding to promoter or enhancer regions containing specific sequences (DR4 elements or LXREs) (9). In addition to their role in lipid metabolism, LXRs also participate in the transcriptional regulation of inflammation and host defense (10–14). Ligand-activated LXRs are able to antagonize the expression of inflammatory genes in response to different insults, a process that has been extensively studied in macrophages (11, 13, 15).
In contrast to macrophages, the transcriptional responses controlled by LXR in other immune cells, such as DCs, are poorly understood in comparison. In this study, we applied LXR gain- and loss-of-function approaches to define a role for LXR in DC gene expression programs that are important for DC chemotaxis. We show that the ability of LXRs to regulate DC migration in response to chemotactic signals is accomplished mainly via the transcriptional regulation of CD38 expression. In addition, CD38 activity is important for the accumulation of lipid-laden myeloid cells in response to atherogenic inflammation. These results outline a previously unrecognized role for LXR signaling in the regulation of chemotactic responses in murine DCs through the transcriptional induction of CD38 expression.
RESULTS
Contribution of LXRs during DC differentiation and maturation.Previous work established that the transcriptional activity of LXRα and LXRβ in macrophages is important for the regulation of inflammation, phagocytosis, and innate immune homeostasis (16). However, the impact of LXR signaling on DC function has not been explored in depth. We performed whole-genome microarray studies with RNA samples obtained from in vitro cultures of hematopoiesis-derived human and mouse DCs (Fig. 1A to C). These experiments revealed that both LXRα and LXRβ were moderately to highly expressed in human and mouse DCs (Table 1). Furthermore, cluster analysis of nuclear receptor transcript levels revealed the prominent expression of both LXRα and LXRβ in purified mouse and human tissue DC populations (Fig. 1C). These experiments also revealed that bone marrow-derived DCs (BMDCs) and splenic or lymph node (LN) classic DCs express higher transcript levels of LXRα than of LXRβ (Fig. 1C). Public repositories of transcript data sets show expression levels of LXR subtypes in DCs that are consistent with our observations (17).
Transcription factor and nuclear receptor expression profiles in ex vivo-differentiated mouse and human DCs. (A) Expression levels of transcription factors, including nuclear receptors, in mouse BMDCs (left) and human monocyte-derived DCs (right) were compared to all probe sets of an Affymetrix microarray. If more than one probe set represents a certain gene, the probe set with the highest signal intensity is shown. (B) Microarray-based comparison of the numbers of highly, moderately, and not expressed transcription factors as well as nuclear receptors in mouse and human ex vivo-differentiated DCs. (C and D) Heat map illustrating differentially or commonly expressed nuclear receptors in different types of human and mouse DCs. Blue indicates low expression levels, and red indicates high expression levels. We identified both LXRα and LXRβ in a selected list of receptors that are moderately to highly expressed. BMD DCs, bone marrow-derived dendritic cells; SLN, sentinel lypmh node; MLN, mesenteric lymph node.
Microarray-based comparison of all nuclear receptors expressed in mouse and human ex vivo-differentiated DCs
To investigate the impact of endogenous LXR activity on DC differentiation and maturation in vitro, we employed flow cytometry to assess the expression of classic DC activation markers in cultured monocyte-derived DCs (MoDCs) obtained from wild-type (WT) and LXR-deficient (lacking both the Lxrα and Lxrβ genes) (designated here LXR double knockout [LXR-DKO]) mice. The activation of MoDCs with the Toll-like receptor 4 (TLR4) agonist lipopolysaccharide (LPS) increased the expression levels of CD11c and of costimulatory molecules required for T-cell priming, including major histocompatibility complex class II (MHC-II), CD80, CD86, and CD69, in both WT and LXR-DKO MoDCs (Fig. 2A and B). To validate these results in vivo, spleen and LN classic MHC-IIhi/CD11chi DCs (which express high levels of LXRs in WT mice) were present at similar frequencies in samples obtained from WT and LXR-DKO mice (Fig. 2C). We also identified DCs by histological examination of spleen and lymph node cryosections immunolabeled with a CD11c antibody. Analysis of WT and LXR-DKO samples showed similar distributions of DCs in both tissues, with CD11c+ cells being localized primarily within the T-cell zone (Fig. 2C). Consistent with these results, analysis of MHC-II+ cells obtained from ear epidermal sheets did not reveal appreciable differences in the expression levels of the myeloid markers CD68 or langerin in samples from WT and LXR-DKO mice (Fig. 2D and data not shown). Together, these data suggest that endogenous LXR activity is not required for the development or differentiation of DCs in vitro or in vivo.
Differentiation and activation of DCs from WT and LXR-DKO mice. (A and B) WT and LXR-DKO MoDCs were differentiated in vitro. (A) Flow cytometry analysis of MHC-II and CD11c expression. (B) Flow cytometry analysis of the expression of the DC maturation markers MHC-II, CD80, CD86, and CD69. Mean fluorescence intensity (MFI) quantifications are graphed below each plot. (C, left) Flow cytometry analysis of classic MHC-II+/CD11c+ DCs in splenic and LN cell suspensions from WT and LXR-DKO mice. (Right) Immunofluorescence analysis of spleen and LN sections from WT and LXR-DKO mice showing combinations of double staining with antibodies that recognize CD11c+ DCs and CD4+/CD8+ cells within the T-cell zone (spleen) or CD11c+ DCs and CD169+ subcapsular sinus macrophages (LN). Bars, 100 μm and 50 μm for spleen and LN samples, respectively. (D) Immunofluorescence analysis of MHC-II and CD68 expression in epidermal sheets prepared from WT and LXR-DKO mice. For panels A to D, representative plots and images from two independent experiments with 3 or 4 mice per genotype are shown.
We next investigated whether the in vitro maturation of DCs promoted changes in established LXR target genes in cultured MoDCs. (Fig. 3A) (18–20). We observed that DC maturation induced by LPS led to the upregulation of some LXR targets and to the downregulation of others (Fig. 3A and B). These results indicate that LXR-dependent gene expression in DCs might be influenced by various factors, such as endogenous LXR ligand availability. Furthermore, although LXR activity has been studied in mature DCs with pharmacological activation approaches using synthetic agonists in vitro (17, 18, 21), we considered the possibility that additional LXR target genes important for DC immune functions arise using our LXR genetic loss-of-function system.
Influence of LXR deficiency during DC maturation. (A) Real-time qPCR analysis of Abca1 and Pltp gene expression in WT iDCs and mDCs in response to the synthetic LXR ligand GW3965 (1 μM) (24 h). Statistical analysis was performed via Student's t test. *, P < 0.05. Error bars represent means ± SD. (B) LXR target gene expression during DC maturation (mDCs versus iDCs) (24 h of LPS treatment at 100 ng/ml) and in response to GW3965 (1 μM for 24 h) in mDCs. (C) Transcriptional profiling of WT and LXR-DKO iDCs and mDCs. (Left) Venn diagram representation showing the overlap of upregulated genes (5-fold or more in mDCs versus iDCs) in WT and LXR-DKO DCs. (Middle) Heat map illustrating differentially or commonly regulated genes in WT and LXR-DKO mDCs versus iDCs. (Right) Top KEGG pathways obtained from GO analysis of exclusively or commonly induced genes in WT and LXR-DKO DCs during DC maturation. Examples of representative genes from each group are listed.
To study the influence of endogenous LXR signaling on DC gene expression programs in depth, we conducted global gene expression analysis with WT and LXR-DKO DCs stimulated for 24 h with LPS. Using a stringent cutoff threshold of 5-fold or higher, we concentrated on the subsets of genes that were highly induced by DC maturation. In agreement with data from previous studies (22), considerable proportions of genes whose expression was induced by LPS in both WT and LXR-null cells were known targets with direct functions in antimicrobial and inflammatory responses in DCs (Fig. 3C). Interestingly, the magnitude of changes in inflammatory gene expression during DC maturation was generally higher in LXR-DKO cells than in WT control DCs (Fig. 3C, heat map). These results are consistent with the anti-inflammatory role of LXRs in other cell types (15). In addition, LXR-DKO cells presented a substantial increase in the number of maturation-induced genes, likely reflecting the existence of several derepressed inflammatory pathways in the absence of LXR.
Identification of LXR-regulated genes during DC maturation.In an effort to identify LXR-regulated genes in DCs that might contribute to LXR functions in immunity, we analyzed a subgroup of genes whose expression was preferentially upregulated in WT but not in LXR-DKO cells during DC maturation. We identified a set of genes (<30 genes) whose expression was differentially induced in WT mature DCs (Fig. 3C). The gene set included those encoding proteins with previously defined roles in innate immunity, inflammation, and chemotaxis, such as the interferon-responsive proteins IFIT2 and GBP3, the chemokine CXCL16, and the ectoenzyme CD38 (Fig. 3C; see also Table S2 in the supplemental material). DC maturation with LPS also promoted the expression of the established LXR target Abca1 in an LXR-dependent manner (Fig. 3C and 4A).
Cd38 is an LXR-responsive gene in DCs. (A) mRNA expression levels of Abca1, Cd38, Srebf1, and Abcg1 in WT and LXR-DKO iDCs and mDCs. (B) Regulation of mRNA expression levels of Srebf1 and Cd38 in WT and LXR-DKO iDCs and mDCs in response to GW3965 (1 μm for 24 h). In panels A and B, error bars represent means ± SD. *, P < 0.05; **, P < 0.01; †, P < 0.05. (C) Immunofluorescence microscopy analysis of CD38 protein expression in WT and LXR-DKO mDCs in the presence of the LXR agonist GW3965 (100 ng/ml LPS for maturation and 1 μM GW3965 added simultaneously). (D) Flow cytometry analysis of CD38 protein expression in WT and LXR-DKO iDCs and mDCs compared to an isotype control antibody. In panels C and D, representative images and plots were obtained from three independent experiments.
We further tested whether the expression of genes in this maturation-dependent cluster was acutely responsive to the activation of LXR/RXR heterodimers by synthetic ligands. Srebf1 expression was potently induced upon LXR activation, as expected (Fig. 4B). Interestingly, within this cluster of maturation-dependent genes, we found that mRNA levels of Cd38 were consistently upregulated by the synthetic LXR ligand GW3965 in WT DCs but not in LXR-DKO cells (Fig. 4B). In addition, the induction of Cd38 expression by LXRs was more prominent in mature DCs than in immature DCs (Fig. 4B).
The gene encoding the ectoenzyme CD38 was recently identified as an LXR target in macrophages in a separate report by some of the authors of the present study (14). Since Matalonga et al. evaluated the LXR-dependent regulation of CD38 expression in the context of macrophage antibacterial responses, we decided to further study the impact of LXR-CD38 signaling in DCs in more detail. CD38 is a type II transmembrane glycoprotein highly expressed by hematopoietic and nonhematopoietic cells. Also known as ADP-ribosyl cyclase 1, CD38 is a multifunctional enzyme that presents both extracellular and intracellular activities, including the ability to produce cyclic adenosine diphosphoribose (cADPR) and ADP-ribose (ADPR) from NAD+. Interestingly, CD38 actions were previously linked to leukocyte trafficking in response to inflammation, and Cd38−/− mice mount inefficient innate and adaptive immune responses (23, 24). The experiments described above indicated that CD38 expression is transcriptionally regulated by LXRs in murine DCs. Further analysis revealed that CD38 protein expression (analyzed by immunocytochemistry and flow cytometry) was severely compromised in mature LXR-DKO DCs in comparison to WT cells (Fig. 4C and D). Moreover, Cd38 mRNA expression was also induced in response to LXR agonists in human monocyte-derived DCs (Fig. 5), suggesting that the regulation of CD38 expression by LXRs is preserved across species.
mRNA expression levels of CD38 and established LXR targets in human monocyte-derived iDCs and mDCs were analyzed by real-time qPCR. Cells were obtained from CD14+ monocytes isolated from buffy coats, differentiated for 7 days with GM-CSF and IL-4, and further stimulated for 24 h with LPS in the absence or presence of 1 μM GW3965. Representative graphs of data from 3 independent experiments are shown. Error bars represent means ± SD of results from three experiments. **, P < 0.01; *, P < 0.05.
LXR is required for efficient CCR7-dependent chemotaxis in DCs.Previous work demonstrated that CD38 is important for host responses against pathogens including Listeria monocytogenes, Streptococcus pneumoniae, and Salmonella enterica serovar Typhimurium (14, 23, 24). Further studies concluded that migratory defects in bone marrow-derived cells underlie the increased susceptibility of Cd38−/− mice to infection. We therefore considered the possibility that LXR signaling contributes to chemotactic activity in DCs by regulating CD38 expression. To test this hypothesis, we used migration assays to analyze the chemotactic capacities of WT and LXR-DKO DCs in response to CCL21 and CCL19, which are ligands of the G-protein-coupled receptor CCR7. Importantly, equivalent mRNA and protein levels of CCR7 were observed in WT and LXR-DKO DCs in response to TLR activation signaling (Fig. 6A and data not shown). It is well documented that immature DCs present a weak migratory capacity in response to CCL21 and CCL19 (25, 26). Consistent with data from those studies, both WT and LXR-DKO immature DCs did not respond significantly to CCL19 and CCL21 stimulation (Fig. 6B, left). Remarkably, while mature WT DCs migrated robustly toward CCL21 or CCL19 gradients, a drastic decrease in chemotactic activity for both ligands was observed in LXR-DKO DCs (Fig. 6B, right).
Deficient chemotactic responses in LXR-DKO DCs in response to CCL19/CCL21. (A) mRNA levels of CCR7 in WT and LXR-DKO iDCs and mDCs analyzed by real-time qPCR. (B) Chemotaxis of WT and LXR-DKO iDCs and mDCs in response to CCL19 and CCL21 (100 ng/ml for 3 h) was analyzed in transwell migration assays. (C) Chemotaxis of isolated splenic DCs from WT and LXR-DKO in response to CCL19, CCL21, and GW3965. (D) Activation of signaling pathways by CCL19 in WT and LXR-DKO BM-derived mDCs. Cells were treated CCL19 for the indicated times, and protein extracts were analyzed by Western blotting with antibodies that recognize phospho-ERK (p-ERK), ERK1/2, phospho-Akt, and β-actin. The Western blot is representative of data from 3 independent experiments. Graphs are representative of data from 3 independent experiments with triplicate samples. **, P < 0.01; *, P < 0.05.
To further characterize the impact of LXR activity on DC migration, primary splenic DCs that had been pretreated with the vehicle or GW3965 were analyzed in transwell migration assays in response to CCR7 ligands. The activation of LXRs by GW3965 potentiated the CCL19- and CCL21-dependent migration of mature wild-type but not LXR-DKO DCs (Fig. 6C). These results indicate that while CCR7 expression levels in primary WT and LXR-DKO DCs are comparable, chemokine-induced migration is significantly regulated by LXR expression and activity in murine DCs.
In addition to promoting chemotaxis, the activation of CCR7 affects several additional functions of mature DCs, including survival (27, 28). These functions have been shown to be regulated by distinct downstream signaling pathways (29). While the CCR7-dependent survival of DCs is regulated mainly by phosphatidylinositol 3-kinase (PI3K)/Akt signaling, DC chemotaxis is controlled by mitogen-activated protein kinase (MAPK) signaling. Since both LXR and CD38 regulate DC chemotaxis, we considered whether the LXR-CD38 axis may participate in chemotactic signaling by regulating MAPK pathways downstream of CCR7. Stimulation of WT DCs with CCL19 resulted in a rapid activation of extracellular signal-regulated kinase 1/2 (ERK1/2) and Akt, as expected (30, 31) (Fig. 6D). In contrast, CCL19-induced phosphorylation of ERK1/2 was markedly inhibited in LXR-DKO DCs. Interestingly, the levels of Akt activation in response to CCL19 stimulation were similar in WT and LXR-DKO cells. These results indicate that LXR activity is important for the chemokine-induced activation of intracellular pathways that direct the migration of DCs but is dispensable for CCR7-dependent survival pathways.
DC migration in vivo is impaired in LXR-deficient mice.To investigate whether LXRs play a role in DC migration in vivo, we employed two widely accepted models. First, we used the classic approach of fluorescein isothiocyanate (FITC) skin painting to monitor the migration of endogenous skin antigen-presenting cells (26). The epicutaneous application of FITC under inflammatory conditions stimulates the activation and migration of DCs carrying the FITC antigen to draining lymph nodes. Immunofluorescence analysis of LNs obtained from FITC-painted mice showed that LXR-DKO LNs contained few CD11c+/FITC-positive (FITC+) DCs compared to WT LNs (Fig. 7A, left). Consistent with this result, flow cytometry analysis of LN cell suspensions revealed decreased numbers of migrated FITC+ DCs 24 to 48 h after FITC painting in LXR-DKO versus WT mice (Fig. 7A, bottom). Thus, LXR signaling is important to guide the migration of endogenous DCs to LNs in response to an antigenic stimulus in vivo.
Impaired migration of LXR-DKO DCs in vivo. (A) Skin contact sensitizer-induced DC migration to draining LNs. FITC+ DCs were identified by immunofluorescence analysis of consecutive sections of WT and LXR-DKO LNs isolated 24 to 48 h after FITC painting. CD11c+/FITC+ DCs were localized within the T-cell zone (CD3+). The graph at the bottom represents the frequency of CD11c+/FITC+ DCs analyzed by flow cytometry of LN cell suspensions obtained from mice painted with FITC for 24 and 48 h. (B) A total of 2 × 106 WT and LXR-DKO BMDCs were labeled with CellTracker red and green, respectively, and coinjected into footpads of WT mice. Inguinal LNs were isolated at 24 h postinjection, and tissue and cell suspensions were analyzed by immunofluorescence microscopy and flow cytometry. Immunofluorescence microscopy results are representative of data from three independent experiments with 3 mice. Graphs represent means ± SD of data from three experiments. **, P < 0.01; *, P < 0.05.
To determine whether the migration deficit observed in LXR-DKO DCs was cell intrinsic, we used a second model, in which we coinjected differentially labeled activated DCs from WT and LXR-DKO mice into footpads of recipient mice. Draining LNs were collected at 24 h postinjection, and the presence of migrated DCs was assessed by flow cytometry and immunofluorescence microscopy. While WT DCs reached the draining LNs after subcutaneous injection, a consistent decrease in the frequency of LXR-DKO DCs was detected by both experimental approaches (Fig. 7B). These results confirm that LXR-DKO DCs have an impaired migratory capacity in vivo due to a cell-intrinsic defect.
CD38 is required for LXR-dependent DC chemotaxis.Although CD38 catalyzes the production of several nucleotide-based metabolites from NAD(P)+, the generation of cADPR has been shown to be particularly relevant for the immunoregulatory functions of CD38 (32). Previous studies using the cADPR antagonist 8-Br-cADPR demonstrated that CD38 regulates calcium flux and the migration of chemokine-activated DCs in a cADPR-dependent manner (33). To determine the impact of CD38 catalytic activity on LXR-dependent DC chemotaxis, we analyzed the migration responses of WT and LXR-DKO DCs that were pretreated with GW3965 in the presence or absence of 8-Br-cADPR. Chemotaxis in response to CCL19 was greatly reduced in cells cultured with 8-Br-cADPR, in agreement with data from previous reports (Fig. 8A). Interestingly, inhibition of the activity of CD38 enzymatic products by 8-Br-cADPR blocked the GW3965-dependent induction of chemotaxis in wild-type DCs but had little effect on LXR-DKO cells (Fig. 8A). These results indicate that ligand-activated LXRs contribute to the regulation of DC chemotaxis through the generation of CD38-dependent enzymatic products.
CD38 activity is required for LXR-dependent regulation of DC chemotaxis. (A) Analysis of in vitro transwell migration of WT and LXR-DKO mDCs in the presence of a CD38 inhibitor. (B) Genetic absence of CD38 abolishes LXR-dependent DC chemotaxis. Chemotaxis of WT, CD38-KO, and LXR-DKO mDCs in response to CCL19 and GW3965 was analyzed by transwell migration assays. Error bars represent means ± SD. **, P < 0.01; *, P < 0.05; n.s., not significant.
To definitively assess the importance of LXR-CD38 signaling in the regulation of DC chemotaxis, we used a CD38 genetic loss-of-function model. As shown in Fig. 8B, the stimulation of CCR7-dependent DC chemotaxis by the LXR ligand was abolished in Cd38−/− cells, indicating that the LXR-dependent induction of Cd38 expression is functionally relevant during DC activation and migration. Interestingly, LXR-deficient DCs showed a decreased chemotaxis capacity compared to that of Cd38−/− cells (Fig. 8B), suggesting that LXR signaling participates in DC migration pathways through both CD38-dependent and -independent mechanisms.
We also analyzed the impact of CD38 deficiency on the frequency of circulating leukocytes under homeostatic conditions. Blood count comparisons indicate that CD38 activity does not affect substantially the number of circulating myeloid/lymphoid populations under homeostatic conditions (Fig. 9A). Furthermore, we tested the frequencies of peripheral blood cells and CD11c+ cells in spleen, bone marrow, and skin of irradiated WT mice (CD45.1) reconstituted with WT or CD38−/− (both CD45.2) progenitors. WT and CD38−/− donor-derived cells were present in these tissues of transplanted mice with similar efficacies (Fig. 9B and data not shown). Therefore, CD38 deficiency does not influence the frequency of circulating cells under homeostatic conditions or the differentiation of lymphoid/myeloid cells in response to a regimen of bone marrow transplantation.
(A) Analysis of blood leukocyte populations obtained from WT and CD38−/− mice under homeostatic conditions (n = 5 each group). WBC, white blood cells. (B) Lethally irradiated C57BL/6 CD45.1+ hosts were reconstituted with either C57BL/6 CD45.2+ or CD38-KO CD45.2 donor cells. Twelve weeks after reconstitution, peripheral blood (PB) myeloid and lymphoid populations were analyzed by flow cytometry.
CD38 activity in hematopoietic cells affects atherosclerosis development.LXR-dependent gene expression in bone marrow-derived myeloid cells has been shown to impact the development of atherosclerosis through various mechanisms (34). We hypothesized that the regulation of Cd38 expression by LXRs in myeloid cells may contribute to the migratory capacity of phagocytic cells during atherosclerosis. To test this idea, we transplanted bone marrow progenitor cells obtained from WT or Cd38−/− mice into lethally irradiated Ldlr−/− mice and analyzed lesion formation after a regimen of Western diet feeding. We confirmed efficient bone marrow engraftment after transplantation by analyzing Cd38 mRNA expression (not shown). We did not observe changes in body weights or plasma cholesterol levels between Cd38−/− and WT progenitor-reconstituted mice (not shown). Remarkably, however, quantification of atherosclerosis by en face analysis after 18 weeks of the Western diet revealed a significant reduction in the atherosclerotic burden in Ldlr−/− recipients reconstituted with Cd38−/− bone marrow compared to WT bone marrow-reconstituted Ldlr−/− mice (n = 15 to 20 mice per group) (Fig. 10A). Immunohistochemical analysis with a CD68 antibody revealed a considerable reduction of total macrophage infiltration within lesions of Cd38−/− bone marrow-transplanted mice (Fig. 10B and C). Analysis of the relative signal of CD68 antigen per atherosclerotic lesion also revealed a trend toward reduction; however, it did not reach statistical significance (Fig. 10D). Overall, these results indicate that CD38 activity in myeloid cells is a determinant of atherosclerosis susceptibility and further suggest that CD38-dependent chemotaxis mechanisms play an important role in the infiltration of mononuclear cells during early atherosclerosis development. Collectively, our results outline a novel pathway by which LXRs participate in myeloid-derived DC migration through the direct regulation of CD38 expression.
CD38 expression in bone marrow cells is important for atherosclerosis development. (A) Percentage of the aorta surface area with atherosclerotic plaques in transplanted LDLR−/− mice after 18 weeks on a Western diet. Horizontal lines indicate means ± standard errors of the means. The right panels show representative photographs from en face analysis. A total of 15 to 20 mice in each group were analyzed. (B) Representative micrographs of frozen sections from the aortic roots of transplanted WT and CD38−/− mice that were stained with CD68 antibody (n = 20 for WT mice, and n = 15 for CD38−/− mice). (C) Quantification of the total CD68 signal within atherosclerotic lesions from each group (n = 20 for WT mice, and n = 15 for CD38−/− mice). (D) Quantification of the CD68 signal relative to each atherosclerotic lesion from both transplanted WT and CD38−/− mice (n = 7 per group). NS, not significant.
DISCUSSION
LXRs are crucial regulators of lipid metabolism that exert important functions in inflammation and host immunity (12, 15, 34, 35). Although the LXR pathway has been extensively studied in macrophages under inflammatory conditions, little is known about LXR transcriptional programs in DCs. Indeed, to our knowledge, this work is the first to report LXR actions in DCs using a complete genetic LXR deficiency (double deficiency of LXRα and LXRβ) in vitro and in vivo. We present a comprehensive analysis of LXR transcriptional activity in primary DCs by combining pharmacological and genetic manipulations of the LXR pathway with global gene expression analysis in models of DC activation. Our results revealed that LXR activity potentiates DC chemotaxis in vitro and in vivo. We furthermore showed a plausible mechanism by which LXRs modulate DC chemotaxis through the transcriptional regulation of CD38 expression.
Immune responses are initiated in secondary lymphoid organs, where DCs migrate to present antigens to naive T and B cells (2). The ability of DCs to migrate requires the expression of the lymphoid homing receptor CCR7 by maturing DCs (4). Using two in vivo models of stimulus-induced chemotaxis, we demonstrated that the migration of DCs to draining lymph nodes requires an intact LXR signaling pathway. It is possible that if lymphocytes encounter reduced numbers of antigen-presenting cells due to impaired chemotaxis, the development of an acquired immune response might be compromised in LXR-deficient mice. Although in vivo adaptive immunity has not been addressed in depth in LXR-null mice, these mice present abnormal lymphocyte proliferation and develop autoimmunity with age (36, 37). Therefore, additional studies will be required to determine the role of the LXR-dependent migration of APCs in the context of adaptive immunity. For example, through the generation of new mouse models of an LXR conditional deletion in APCs, it should be possible to directly test the contribution of LXR-dependent DC migration in the context of adaptive immunity.
In view of the impaired migratory capacity of LXR-DKO DCs, we examined the level of activation of chemokine-dependent signaling pathways in our DC culture system. Our findings revealed that although the magnitudes of intracellular Akt activation were equivalent, the induction of the MAPK pathway by chemokines was defective in LXR-DKO DCs compared to WT cells. Previous studies reported that pharmacological LXR activation can influence, both positively and negatively, the expression of the chemokine receptor CCR7 in different cultured cells in vitro or in vivo in models of disease, such as atherosclerosis regression or tumor progression (38–41). The reasons for these contrasting results are not entirely clear but could be due to differences in basal CCR7 expression levels that may not be similarly regulated upon stimulation among the particular models of cellular activation. Alternatively, the employment of different doses of various activating agonists, either synthetic ligands or natural oxysterols, under different culture conditions may account for these contrasting results (38–41). Nevertheless, our results using genetic LXR deficiency indicate that primary LXR-null DCs express CCR7 levels equivalent to those expressed by WT cells during DC maturation. Thus, a deficiency of LXR renders DCs hyporesponsive to CCL19/CCL21 chemokines, despite the normal expression of CCR7 and several other maturation markers. Importantly, the maturation of DCs promoted the expression of several LXR targets, including Abca1 and Cd38, while inhibiting others, such as Srebf1. Interestingly, although LXR is not the sole transcription factor involved in the expression of these targets, the upregulation of Abca1 and Cd38 by mature DCs was found to be dependent largely on LXR expression. This suggests that endogenous LXR activators might be generated during DC maturation by inflammatory signals. Alternatively, given that other LXR targets were downregulated by DC maturation, gene-specific epigenomic changes induced in the setting of inflammation may affect LXR and/or coregulator binding at target promoters/enhancers. Further work will be required to delineate the specific genomic locations of LXR binding during the inflammatory activation of DCs.
Using models of LXR pharmacological activation and LXR deficiency in DCs, we showed that LXRs directly upregulate CD38 expression in primary DCs. CD38 is a multifunctional enzyme that belongs to the ADP-ribosyl cyclase family and has both ectoenzyme and receptor functions. Previous studies demonstrated that CD38-deficient mice are unable to mount an effective immune response against bacterial infections (14, 23, 24). The inability of myeloid cells to directionally migrate to sites of infection was reported to be a possible explanation for the defective antimicrobial responses in these mice. Interestingly, both LXR- and CD38-deficient mice present immune defects against bacterial infections (12, 14, 23, 42). Because LXR regulates the expression of CD38, the migration of specific subsets of APCs in vivo to sites of infection in the context of LXR deficiency could be an interesting angle to explore in future studies. However, such studies could be also confounded by the fact that LXR-null mice are defective in splenic marginal-zone macrophages (35), a specific subset of macrophages important for the systemic capture of circulating antigens. Thus, new mouse models with an APC-specific deficiency of LXR in which the splenic marginal zone is not affected would be potentially eligible for analyses of the recruitment of APCs in response to systemic infections.
Importantly, other studies have documented the cellular localization of CD38 in lipid-rich membrane domains. An association of CD38 with membrane signaling receptors, including CCR7, CD83, and CD11b, has been reported for human MoDCs (43). Since CD38 ensures efficient migration in response to CCR7 ligands (32, 44), it is therefore possible that the ligation of CCR7 induces interactions with CD38 and other signaling receptors within lipid rafts, in which membrane cholesterol fluidity is important (45). Thus, cooperation between CCR7, CD38, and perhaps other receptors in lipid rafts may regulate innate and adaptive immune responses by modulating DC migration and survival. We found that the influence of LXR signaling on this DC chemokine signaling cross talk was dependent largely on CD38 expression. The loss of CD38 abolished the LXR-dependent induction of CCR7-dependent chemotaxis. Interestingly, LXR-deficient DCs presented a more profound impairment in DC chemotaxis than did CD38−/− DCs, suggesting that LXR signaling participates in multiple pathways that control DC chemotaxis, both CD38 dependent and independent. One possibility is that the accumulation of excess cholesterol in membrane microdomains in LXR-DKO DCs could be altering the interaction between chemokine receptors and coreceptors that is important for migration signaling.
The migration of myeloid cells into the artery wall in response to chemotactic molecules is one of the key steps in early atherosclerotic lesion formation (46). In this regard, although the recruitment of monocytes may initially serve as a protective mechanism to remove excess low-density lipoprotein (LDL) cholesterol and reduce inflammation, the progressive accumulation of cells in the artery wall in the context of hypercholesterolemia ultimately leads to atheroma formation. Many studies have demonstrated that interference with myeloid chemotaxis retards the development of atherosclerosis (47). On the other hand, in surgical models of plaque regression in which aortas of hypercholesterolemic mice are transplanted into WT mice, a reduction of cholesterol loading in lesional myeloid cells has been shown to induce their emigration from plaques and alleviate atherosclerosis (48). Atherosclerosis regression in these models was shown to be impaired in the absence of LXRα or LXRβ and associated with decreased CCR7 expression in myeloid cells (38). Accordingly, the CCR7-dependent chemotaxis of myeloid cells back to draining lymph nodes appears to play a beneficial role in the context of the regression of atherosclerosis in surgical models. However, other studies have implicated CCR7 expression and CCL19/CCL21 ligands in the progression of atherosclerosis (49, 50). Thus, the migratory capacity of myeloid cells has been associated with both the attenuation and exacerbation of atherosclerosis in different models.
Although studies with knockout mice of some individual LXR target genes have shown divergent effects on atherosclerosis, the net result of LXR activation is clearly atheroprotective (34). This is due to the fact that LXRs regulate the expression of a number of genes whose activity promotes the removal of cholesterol from plaque myeloid-derived foam cells. We showed that the connection between CD38 and DC migration has pathophysiological relevance in the low-density lipoprotein receptor-deficient (LDLR−/−) model of atherosclerosis. A loss of hematopoietic CD38 expression alleviates atherogenic lesion progression. The defective chemotactic capacity of CD38−/− myeloid cells is likely a factor that contributes to this phenotype. These observations suggest that novel CD38 inhibitors could have therapeutic benefit in the setting of atherosclerosis.
Collectively, our results uncover a previously unrecognized mechanism that operates in DCs in which LXRs modulate chemokine signaling in mature DCs, at least in part, through the potentiation of CD38 expression.
MATERIALS AND METHODS
Animals.LXR-deficient (Nhr1h3−/− Nhr1h2−/−) (LXR-DKO) mice on a mixed Sv129/C57BL/6 background were originally provided by David Mangelsdorf (University of Texas Southwestern) (51), and CD38-deficient mice (N-B6.129P2-Cd38 tm1Lnd) had been backcrossed to the C57BL/6 background for more than 10 generations. All mice were maintained under pathogen-free conditions in a temperature-controlled room and with a 12-h light-dark cycle in the animal facilities of the Universidad de Las Palmas de Gran Canaria (ULPGC) and Nationwide Children's Hospital (NCH)/Ohio State University (OSU). All animal studies were conducted in accordance with institutional participants' animal ethics research committees. WT and CD38-deficient mice (n = 5 per group) were euthanized, and blood was collected by cardiac puncture and stored in EDTA-coated tubes. Leukocyte counts were determined by using an Abacus JuniorVet hematologic counter (Diatron).
Reagents.Recombinant murine CCL19, CCL21, and granulocyte-macrophage colony-stimulating factor (GM-CSF) were obtained from Peprotech (London, United Kingdom). The synthetic LXR ligand GW3965 was provided by J. Collins (GlaxoSmithKline). LPS serotype 055:B5 and 8-Br-cADPR were obtained from Sigma. CellTracker green (5-chloromethylfluorescein diacetate [CMFDA]) and the fluorescent dye Cell Tracker red (CMTPX; Molecular Probes C34552) were obtained from Molecular Probes.
Cell isolation and culture.Monocyte-derived DCs (MoDCs) were prepared as described previously (52). In brief, bone marrow (BM) monocytes were purified from cell suspensions through the depletion of T cells, B cells, granulocytes, NK cells, and DCs with antibodies that recognize B220, MHC-II, Thy1.2, CD43, and CD24. Averages of 90 to 95% of monocytes were collected from negatively selected cells. MoDCs were obtained by culturing monocytes with GM-CSF for 24 h. In another set of experiments, BM-derived DCs (BMDCs) were generated in vitro, as described previously (53), with modifications. BM cells were cultured for 6 to 7 days in RPMI 1640 medium containing 10% fetal bovine serum (FBS) supplemented with mouse GM-CSF (20 ng/ml) every 2 days. Nonadherent cells were collected and further enriched in DCs by positive selection with CD11c microbeads (Miltenyi Biotec). For the isolation of DCs from lymphoid tissues, spleen or lymph nodes were digested with 1 mg/ml Liberase CI (Roche), 40 mg/ml DNase I (Roche), and 1% (vol/vol) FBS for 30 min at 37°C in RPMI medium. Cell suspensions were enriched by positive selection with CD11c microbeads (Miltenyi Biotech) and further purified by fluorescence-activated cell sorting using CD11c and MHC-II antibodies. The purity of the DC population based on CD11c and MHC-II expression was >90% by flow cytometry analysis. For DC activation, MoDCs, BMDCs, or purified DCs from lymphoid tissues were stimulated with LPS (10 ng/ml from Escherichia coli; Sigma-Aldrich) for 24 h.
Flow cytometry.Single-cell suspensions (1 × 106 cells) were washed twice in staining buffer (phosphate-buffered saline [PBS] with 0.1% bovine serum albumin [BSA] and 0.1% sodium azide) and incubated with Fc block (anti-CD16/32; Sigma-Aldrich) for 20 min at 4°C. Cells were incubated with labeled antibodies for 30 min at 4°C. Intracellular CCR7 staining was performed according to the manufacturer's instructions (eBioscience). Cells were then analyzed on a BD FACSCalibur or FACSCanto II instrument (Becton Dickinson) with FlowJo software (TreeStar, Inc.) (for a detailed description of antibodies used for flow cytometry, see Table S1 in the supplemental material).
RNA and protein analysis.Total RNA was obtained with TRIzol reagent (Invitrogen). RNA was reverse transcribed with an iScript reverse transcription kit (Bio-Rad). Real-time quantitative PCR (qPCR) was performed with a Bio-Rad iQ5 detector and SYBR green assays, as described previously (35). Expression levels were normalized to the 36B4 expression level (see the supplemental material for primer and probe sequences).
For Western blot analysis, DCs were stimulated with CCL19 for the indicated times. Stimulation was terminated by solubilizing the cells in 100 μl ice-cold radioimmunoprecipitation assay (RIPA) buffer supplemented with protease and phosphatase inhibitors (Sigma). Lysates were resolved by SDS-PAGE and transferred to nitrocellulose membranes (Bio-Rad Laboratories). Membranes were incubated with the indicated antibodies (Table S1). Blots were washed and visualized with the appropriate horseradish peroxidase (HRP)-conjugated secondary antibodies (Bio-Rad), an ECL kit (ECL-Plus; Amersham Biosciences), and a Bio-Rad ChemiDoc imaging system (for a detailed description of the antibodies used, see Table S1 in the supplemental material).
Histology and immunofluorescence staining.Lymph nodes and spleens were embedded in OCT compound (Tissue-Tek) and snap-frozen in dry ice and isopentane. Four- to eight-micrometer frozen sections were air dried, fixed with 4% paraformaldehyde, blocked with 6% BSA and 2% preimmune serum in PBS, and stained with fluorescence-conjugated antibodies diluted in blocking solution.
In vitro migration assay.In vitro chemotaxis assays were performed by using migration chambers. Briefly, DCs were cultured in Transwell chambers (5-μm pore size; Costar) with 100 ng/ml of CCL19 and CCL21 or a control solvent in the lower chamber. After 3 h of incubation at 37°C, migrated cells were quantified by using an automated cell counter (TC-20; Bio-Rad). Where indicated, cells were pretreated with 8-Br-cADPR (10 μM) for 30 min or with GW3965 (1 μM) for 18 h. Each experiment was performed in triplicate. The results are expressed as the mean chemotactic index (CI) ± standard deviation (SD) for triplicate wells. The CI represents the fold increase in the number of migrated cells in response to chemoattractants over the number of cells that spontaneously migrated under control medium conditions.
In vivo DC migration assays.Mouse abdominal and inguinal areas were gently shaved. Fluorescein isothiocyanate (FITC) (0.5%) was dissolved in acetone-dibutylphthalate (1:1) and applied to the exposed areas of mice (54). After 24 to 48 h, draining inguinal LNs were harvested and processed. To test the in vivo migration of ex vivo-differentiated cells, WT and LXR-deficient mature BMDCs were labeled with 1 μM CellTracker green (CMFDA) or CellTracker red (CMTPX), respectively, and resuspended in PBS. A total of 1 × 106 DCs at a 1:1 ratio were injected subcutaneously into the hind footpads of control mice (55). Twenty-four hours after injection, popliteal LNs were harvested. Representative LN samples from each mouse were processed for immunofluorescence analysis or flow cytometry.
Microarray analysis.Total RNA was isolated from iDCs and mDCs (stimulated with LPS at 100 ng/ml for 24 h) by using an RNeasy kit (Qiagen) according to the manufacturer's protocol. RNA quality was assessed on an Agilent Bioanalyzer 2100 instrument (Agilent Technologies). Transcriptional profiling was performed as follows. Mouse Affymetrix 430 2.0 microarray analyses were performed at the Genomics Core Facility, Universidad Complutense, Parque Científico de Madrid, Spain, and at the Center for Clinical Genomics and Personalized Medicine Microarray Facility (University of Debrecen, Hungary). Affymetrix Gene Chip Human Genome U133 Plus 2.0 microarray analyses were conducted at the Microarray Core Facility of the European Molecular Biology Laboratory (Heidelberg, Germany). Data were analyzed with GeneSpring and GeneChip analysis suite software (Affymetrix), as described previously (56). Only statistically significant differences in expression levels are presented. Raw signal intensities were normalized per chip (to the 50th percentile). We removed probe sets that failed to reach raw signal intensities of 50 (human monocyte-derived DCs) and 75 (mouse BMDCs) in all three samples. These values represented roughly the median of the signal intensity values for all probe sets. We defined the remaining probe sets as expressing genes. Next, we calculated the median raw values for the expressing genes and created two categories. The expressed genes with raw values over the median were categorized as highly expressed genes, while the expressed genes with raw values under the median were categorized as moderately expressed genes. All data sets are available through the NCBI GEO server or the ArrayExpress database (GEO accession numbers GSE15907 [public data set from the IMMGEN consortium] and GSE23618 [human DC data] and ArrayExpress accession number E-TABM-34 [ArrayExpress database DC subsets in human tonsils and blood]).
Isolation and culture of human DCs and mouse epidermal sheets.Human peripheral blood mononuclear cells (PBMCs) were isolated from healthy volunteers' buffy coats by Lymphoprep (NycomedPharma, Oslo, Norway) gradient centrifugation, according to standard procedures, followed by immunomagnetic separation with anti-CD14-conjugated MicroBeads (VarioMACS separation system; Miltenyi Biotec). Monocytes were cultured in 6-well plates at a density of 106 cells/ml in RPMI 1640 medium (Sigma-Aldrich) supplemented with 10% FBS (Invitrogen), 2 mM l-glutamine (Invitrogen), and penicillin-streptomycin (Sigma-Aldrich). For DC differentiation, we treated the freshly isolated monocytes with 800 U/ml GM-CSF (Leucomax; Gentaur Molecular Products) and 100 ng/ml interleukin-4 (IL-4; PeproTech) for 5 days. Epidermal sheets were obtained from ears of wild-type and LXR-deficient mice as described previously (4). Briefly, ears were split into dorsal and ventral halves and floated split side down for 2 h on 20 mM EDTA in PBS at 37°C. The epidermis was separated from the dermis with fine forceps, washed twice in PBS, and fixed in ice-cold acetone for 20 min at room temperature. After rehydration in PBS, sheets were processed as described above for lymph nodes and spleen. To detect the cell nucleus, samples were stained with 4′,6-diamidino-2-phenylindole (DAPI; Sigma-Aldrich). Sections were observed with an LSM 5 Pascal laser scanning microscope (Carl Zeiss).
Bone marrow transplant histological and lesion analyses.Recipient male LDLR−/− mice (B6.129S7-Ldlrtm1Her/J; Jackson Laboratory) or WT C57BL/6 CD45.1 mice were lethally irradiated with 900 rad and transplanted with 3 × 106 BM cells from donors that were 8 weeks of age or older (WT or CD38) via tail vein injection as previously described (57). After 4 weeks of recovery, transplanted LDLR−/− mice were fed a Western diet (catalog number D12079B; Research Diets) for 18 weeks. Mice were euthanized and perfused with 0.5 mM EDTA–PBS. Aortas were dissected, fixed (4% paraformaldehyde, 5% sucrose, 20 μM EDTA), pinned, and stained with Sudan IV. Images were captured with a charge-coupled-device (CCD) camera. Atherosclerosis in the aortic roots and the descending aortas (en face) was quantified by computer-assisted image analysis as described previously (56, 58). Lesion development is expressed as a percentage of the total aortic surface covered by lesions (56). The preparation and staining of frozen sections from aortas were performed as described above. Immunohistochemistry to detect CD68 antibody binding (catalog number MCA1957GA; AbD) at a 1:400 dilution with biotin with a 6-atom spacer [biotin-SP-conjugated AffiniPure goat anti-rat IgG(H+L) secondary antibody; Jackson Laboratories] was used.
Statistical analysis.Experimental groups included at least 4 to 5 mice. All experiments were performed at least 3 times. Data were expressed as means ± SD. Statistical analyses were performed with SPSS software (IBM). An analysis of variance (ANOVA)-Bonferroni test or Student's t test was used to determine statistical differences between multiple or paired comparisons with a normal distribution of the data.
ACKNOWLEDGMENTS
We thank David Mangelsdorf (University of Texas Southwestern) for the LXR-null mice, Jon Collins (GlaxoSmithKline SA North Carolina) for the LXR agonist GW3965, and Andres Hidalgo and Lisardo Bosca for reagents and comments.
This work was supported by the following grants: Spanish Ministerio de Ciencia y Tecnologia grants SAF2008-00057, Ministerio de Economia y Competitividad (MINECO) SAF2011-29244, and SAF2014-56819-R and Comunidad de Madrid grant S2010/BMD-2350 to A.C.; MINECO grant SAF2014-57856 to A.F.V.; a grant from the Spanish MINECO Nuclear Receptor Excellence Network NuRCaMeIn (SAF2015-71878-REDT to A.F.V. and A.C.); and NIH grants HL-066088 and HL-030568 to P.T. S.B. was supported in part by a fellowship from the Spanish Ministerio de Ciencia y Tecnologia (BES2006-12056), and J.M. received a fellowship from the Spanish Ministerio de Ciencia e Innovacion MICINN (FPI, BES-2009-014828).
FOOTNOTES
- Received 23 October 2017.
- Returned for modification 21 November 2017.
- Accepted 10 February 2018.
- Accepted manuscript posted online 5 March 2018.
Supplemental material for this article may be found at https://doi.org/10.1128/MCB.00534-17.
- Copyright © 2018 American Society for Microbiology.
