Research ArticleInflammation

γ-Secretase Limits the Inflammatory Response Through the Processing of LRP1

See allHide authors and affiliations

Science Signaling  25 Nov 2008:
Vol. 1, Issue 47, pp. ra15
DOI: 10.1126/scisignal.1164263

Abstract

Inflammation is a potentially self-destructive process that needs tight control. We have identified a nuclear signaling mechanism through which the low-density lipoprotein receptor–related protein 1 (LRP1) limits transcription of lipopolysaccharide (LPS)–inducible genes. LPS increases the proteolytic processing of the ectodomain of LRP1, which results in the γ-secretase–dependent release of the LRP1 intracellular domain (ICD) from the plasma membrane and its translocation to the nucleus, where it binds to and represses the interferon-γ promoter. Basal transcription of LPS target genes and LPS-induced secretion of proinflammatory cytokines are increased in the absence of LRP1. The interaction between LRP1-ICD and interferon regulatory factor 3 (IRF-3) promotes the nuclear export and proteasomal degradation of IRF-3. Feedback inhibition of the inflammatory response through intramembranous processing of LRP1 thus defines a physiological role for γ-secretase.

Introduction

γ-Secretase is an intramembranous protease that processes various transmembrane proteins, thus releasing their intracellular domains (ICDs) into the cytoplasm (1). Nuclear translocation of ICDs and their regulation of transcription were first shown for members of the Notch family, which regulate gene expression during cell-fate decisions and embryonic development (2). A similar γ-secretase–dependent cleavage event was recently described for the receptor tyrosine kinase ErbB4 (3), in which the ICD functions as a transcriptional repressor of astrocytic genes, thereby inhibiting glial differentiation of neural precursor cells (4). Here, we define a previously unknown role for γ-secretase in the transcriptional regulation of inflammation through processing of low-density lipoprotein (LDL) receptor–related protein 1 (LRP1).

LRP1 is a ubiquitously expressed, multifunctional member of the LDL receptor family (5) and serves as a modulator and integrator of several distinct signaling pathways in the vascular wall (6, 7), the blood-brain barrier (8), neurons (9), and macrophages from various tissues (1012). We have shown that after pharmacological activation of protein kinase C (PKC) and metalloproteinase-induced shedding of the extracellular domain (ECD) of LRP1, the ICD of LRP1 (LRP1-ICD) is released from the membrane by γ-secretase (13). Activation of PKC by phorbol esters stimulates the production of matrix metalloproteinases and other inflammatory mediators (14), early events in the inflammatory response to infection and injury (15). Self-limiting mechanisms that keep the production of harmful inflammatory mediators in check are essential to avoid excessive cell damage and tissue destruction. We therefore hypothesized that the LRP1-ICD serves as a mediator of this negative feedback. To test our hypothesis, we examined the role of proteolytic processing of LRP1 in lipopolysaccharide (LPS)–induced inflammatory signaling in vitro and in vivo.

LPS molecules are components of the outer membranes of Gram-negative bacteria that induce a strong host defense response in infected organisms, which includes the increased production of proinflammatory cytokines (16). LPS signals through a receptor complex that contains Toll-like receptor 4 (TLR4). The binding of LPS to TLR4 induces two major branches of intracellular signaling cascades: Myeloid differentiation marker 88 (MyD88)–dependent signaling through the adaptor proteins MyD88 and MyD88 adaptor–like (Mal, also known as TIRAP) leads to the early activation of the transcription factor nuclear factor κB (NF-κB), whereas MyD88-independent signaling through the Toll interleukin-1 (IL-1) receptor–like (TIR) domain–containing adapter-inducing interferon-β (TRIF) and TRIF-related adaptor molecule (TRAM) results in the activation of the transcription factor interferon regulatory factor 3 (IRF-3) and late NF-κB signaling (17).

Here, we report a role for γ-secretase in the self-restriction of inflammation through proteolytic processing of LRP1 and modulation of nuclear signaling by the LRP1-ICD. Exposure of macrophages to LPS resulted in increased shedding of the LRP1-ECD. After γ-secretase–dependent cleavage, LRP1-ICD translocated to the nucleus where it bound to IRF-3 and facilitated its nuclear export. In the absence of LRP1, phosphorylated active IRF-3 accumulated in the nucleus, which resulted in the increased transcription of LPS-inducible genes in lrp1-deficient fibroblasts in vitro and in peritoneal macrophages of conditional lrp1 knockout mice. LPS-stimulated production of proinflammatory cytokines was also increased in lrp1-deficient macrophages. These findings reveal roles for γ-secretase and LRP1 in the inhibition of the inflammatory response, which suggests that these proteins may serve as potential therapeutic targets for the modulation of inflammation.

Results

Nuclear localization of LRP1-ICD

We have previously shown that LRP1 undergoes regulated proteolytic processing that culminates in the release of its ICD from the plasma membrane by the presenilin-containing γ-secretase complex (13). We stably transfected lrp1-deficient mouse embryonic fibroblasts (MEFs) with a complementary DNA (cDNA) that encodes the LRP1 cytoplasmic domain (LRP1-105) to investigate the subcellular localization of LRP1-ICD after its release by γ-secretase. Because free LRP1-ICD is subject to proteasomal degradation (13), the cells were treated with the proteasome inhibitor epoxomicin to allow LRP1-ICD to accumulate. Immunofluorescence analysis revealed a predominantly nuclear localization of LRP1-ICD in the stably transfected cell line (Fig. 1A, images c and f), whereas endogenous full-length LRP1 in wild-type (WT) fibroblasts exhibited a staining pattern compatible with the known endosomal localization of LRP1 (Fig. 1A, images a and d).

Fig. 1

Nuclear localization of free LRP1-ICD. (A) Epoxomicin-treated wild-type MEFs (a and d), LRP1-deficient MEFs (b and e), and MEFs stably expressing free LRP1-ICD (LRP1-105, c and f) were analyzed by immunofluorescence assays with an antibody directed against the C-terminus of LRP1. Nuclear counterstaining was performed with DAPI (4′,6-diamidino-2-phenylindole) (d to f). One experiment representative of five is shown. (B) Nuclear and cytosolic extracts were prepared from WT or LRP1-deficient (k.o.) MEFs after treatment with or without 1 μM epoxomicin (epox) for 12 hours and were subsequently analyzed by Western blotting with antibodies against the C terminus of LRP1, histone 2b (a nuclear marker), and Hsp90 (a cytosolic marker). One experiment representative of five experiments is shown.

γ-Secretase–dependent proteolytic processing of LRP1 takes place constitutively in mammalian cells (fig. S4) (13). We therefore investigated the subcellular localization of endogenous LRP1-ICD in WT fibroblasts. As immunocytochemistry analysis was not sensitive enough for this purpose (Fig. 1A, images a and d), we generated nuclear and cytosolic extracts from epoxomicin-treated WT and lrp1-deficient (LRP1 KO) MEFs for Western blotting studies. A C-terminal fragment of the size of the LRP1-ICD (about 12 kD) accumulated predominantly in the nuclei of epoxomicin-treated WT (Fig. 1B, lane 6) but not LRP1 KO cells (Fig. 1B, lane 8), indicating that after separation from the cell membrane, the free LRP1-ICD translocated to the nucleus.

LPS enhances the proteolytic processing of LRP1

The nuclear localization of free LRP1-ICD suggested that it might function in transcriptional regulation. Increased γ-secretase–dependent production of LRP1-ICD occurs after activation of PKC and metalloproteinase-induced shedding of the LRP1-ECD (13). As the activation of PKC and the subsequent induction of metalloproteinases occur during the course of the inflammatory response, we hypothesized that the LRP1-ICD might function in transcriptional regulation during the inflammatory process. We therefore tested whether induction of the host defense response by LPS could modulate the processing of LRP1.

WT mouse peritoneal macrophages were pretreated with the γ-secretase inhibitor DAPT (N-[(3,5-difluorophenylacetyl)-l-alanyl-2-phenyl]glycine-1,1-dimethylethyl ester), and the abundance of the membrane-bound 25-kD LRP1 fragment generated after shedding of the LRP1-ECD was compared between cells that had been treated with or without LPS. Membrane fractions were isolated and analyzed by Western blotting with an antibody against the C-terminus of LRP1. Treatment with LPS led to the increased extracellular cleavage of LRP1 and the subsequent accumulation of the 25-kD fragment (Fig. 2A, lane 4) relative to that in untreated cells, which indicated that increased production of soluble LRP1-ICD occurred in the course of the inflammatory response evoked by LPS. Additional time course experiments were performed (fig. S5), and experiments performed with highly purified rLPS (rough LPS) gave similar results (fig. S6).

Fig. 2

LPS enhances the proteolytic processing of LRP1. (A) Wild-type primary macrophages were pretreated with the γ-secretase inhibitor DAPT (10 μM) for 2 hours. Cells were then treated with LPS (1 μg/ml) or left untreated for 9 hours, and cell membranes were prepared and analyzed by Western blotting with an antibody against the C-terminus of LRP1. One experiment, representative of four, is shown. (B) Schematic representation of LPS-induced signaling pathways. Binding of LPS to its receptor complex leads to the activation of MyD88- and Mal-dependent signaling that results in the early activation of NF-κB, and of MyD88-independent (TRIF and TRAM-dependent) pathways that lead to the activation of IRF-3 and late activation of NF-κB. TRIF, TIR domain–containing adaptor molecule; TRAM, TRIF-related adapter molecule; MyD88, myeloid differentiation marker 88; Mal, MyD88-like; TRAF6, tumor necrosis factor receptor–associated factor; TBK1, TANK-binding protein kinase; NEMO, NF-κB essential modulator; IKK, IκB kinase.

LRP1 modulates the activation of LPS-induced signaling pathways

As proteolytic processing of LRP1 was increased in response to LPS, we next examined whether downstream LPS signaling events, including the transcription of LPS target genes, were modulated by LRP1-ICD. Signaling and transcriptional regulation by LPS involve the activation of the transcription factors p65 NF-κB and IRF-3 [(17), see also Fig. 2B]. We therefore treated WT and LRP1 KO MEFs with LPS and examined the abundance of inhibitor of NF-κB (IκBα) protein, an inhibitor of NF-κB signaling, by Western blotting analysis. IκBα was slightly more abundant in LRP1 KO cells than in WT cells and the degradation of IκBα in response to LPS was delayed relative to that in WT cells (Fig. 3A and fig. S1A).

Fig. 3

LRP1-ICD modulates LPS-activated signaling through a direct interaction with IRF-3. (A) Whole-cell lysates from WT and LRP1-deficient fibroblasts were prepared after treatment with LPS (10 μg/ml) for the times indicated and from untreated controls (ut). Lysates were analyzed by Western blotting with an anti-IκBα antibody. β-Actin was used as a loading control. (For quantification of Western blotting data, see fig. S1A.) (B) Nuclear extracts were prepared from WT and LRP1-deficient fibroblasts, as well as from LRP1-deficient cells stably transfected with the LRP1-cDNA (k.o.-LRP1), the empty plasmid vector (k.o.-ctrl.), or the LRP1-β-chain (k.o.-LRP1-β-chain). The extracts were analyzed by Western blotting with an anti–phospho-IRF-3 antibody. β-Actin served as a loading control. Inset: Analysis of nuclear extracts from LRP1-deficient cells stably transfected with the LRP1-ICD-cDNA (k.o.-LRP1-105) or the empty plasmid vector (k.o.-ctrl.). One experiment representative of five is shown. (C) Total IRF-3 abundance in the cell types described in (B), was identical, as judged by analysis of Western blots of whole-cell lysates with an anti-IRF-3 antibody. β-Actin served as loading control. (D) Schematic representation of LRP1 and the LRP1 mutants transfected into LRP1-deficient fibroblasts. (E and F) Wild-type (WT) and LRP1-deficient (k.o.) MEFs and k.o. cells transfected with LRP1, the LRP1-β-chain, or the empty plasmid vector were analyzed by Western blotting with an antibody against the C terminus of LRP1. (G) Schematic representation of the GST-LRP1 fusion proteins used in a pull-down assay to detect the direct interaction between IRF-3 and LRP1. The LRP-ICD was N-terminally fused to a GST tag. In addition, a C-terminal LRP1-ICD deletion mutant and constructs with mutations in the first, second, or both NPxY motifs were used. The black box indicates a putative casein kinase II phosphorylation site in the distal LRP1-ICD. (H) GST-LRP1-ICD fusion proteins (see G) were used to pull down IRF-3 from whole-cell lysates of MEFs. Equal input of fusion proteins was visualized by Ponceau staining of the transfer membrane (fig. S7). One experiment representative of three experiments is shown.

To examine activation of IRF-3, we prepared nuclear extracts of WT and LRP1 KO MEFs and analyzed the abundance of phosphorylated IRF-3 (pIRF-3) by Western blotting. We found that basal phosphorylation of IRF-3 was increased in the absence of LRP1 (Fig. 3B, lane 2 compared with lane 1). Expression of LRP1 (lane 3) or of the LRP1-ICD, either in the membrane-bound form (lane 5) or as the free cytoplasmic domain (inset, lane 2), was sufficient to reduce the abundance of pIRF-3. Total IRF-3 was comparable in all cell lines (Fig. 3C).

To further investigate the effect of the LRP1-ICD on the activation of IRF-3, we tested whether LRP1-ICD could directly interact with IRF-3. In experiments with glutathione-S-transferase (GST)–tagged full-length and mutant LRP1-ICD constructs (Fig. 3G), we detected binding of the ICD to IRF-3 (Fig. 3H, lane 8). This interaction was mediated by the second NPxY protein-protein interaction motif of the LRP1-ICD (Fig. 3H, lanes 9 to 12).

LRP1-ICD, part of the transcriptional complex on the interferon-γ promoter, limits LPS-induced transcription in vivo

To investigate the role of LRP1 in the activation of p65 NF-κB and IRF-3 in vivo, we generated primary peritoneal macrophages from mice with a conditional inactivation of the lrp1 gene in cells of the myeloid lineage (LysCre;LRPlox/lox) (fig. S8). LPS reduced the extent of both degradation of IκBα and phosphorylation of p65 in lrp1-deficient macrophages (a myeloid cell type) relative to that of LPS-treated cells from control (LRPlox/lox) mice (Fig. 4A, lane 5 compared with lane 6 and lane 7 compared with lane 8, respectively, and fig. S1B). In contrast, LPS-induced phosphorylation of IRF-3 was increased when LRP1 was lacking, which is consistent with our findings in fibroblasts (Fig. 4B, lane 6 compared with lane 5, and fig. S1C). LRP1 repressed both basal and LPS-induced transcription of interferon (ifn)-γ (Fig. 4C). Chromatin immunoprecipitation (ChIP) experiments performed with an antibody directed against the C-terminus of LRP1 showed the presence of LRP1-ICD in the ifn-γ transcriptional complex after treatment with LPS (Fig. 4E). Basal expression of ifn-β, by contrast, was only minimally enhanced in the absence of LRP1 (Fig. 4D), and no interaction of LRP1-ICD with the ifn-β promoter was detected (Fig. 4F). These findings indicate that LRP1-ICD repressed a subset of LPS-inducible genes. Because LPS enhanced the proteolytic processing of LRP1 (Fig. 2A), negative transcriptional regulation by LRP1-ICD generates a negative feedback loop of LPS-induced inflammatory signaling. This mechanism involves direct interaction with IRF-3, resulting in reduced activation of the protein and repression of target gene promoters by LRP1-ICD.

Fig. 4

The LRP1-ICD regulates the expression of a subset of LPS-induced genes by direct nuclear signaling and by limiting the nuclear localization of pIRF-3. (A) Whole-cell lysates were prepared from WT and LRP1-deficient macrophages after treatment with LPS (1 μg/ml) for the times indicated and from untreated controls (ut). The lysates were examined as described in Fig. 3A. Pooled macrophages from four k.o. and four WT mice were used for each repetition of the experiment (three independent experiments). (For quantification of Western blotting results, see fig. S1B.) (B) Whole-cell lysates were prepared from WT and LRP1-deficient (k.o.) macrophages treated with LPS (1 μg/ml) for the times indicated. Lysates were transferred to Western blots that were incubated with antibodies against LRP1, pIRF-3, IRF-3, and β-actin. Pooled macrophages from three k.o. and three WT mice were used or each repetition of the experiment (three independent experiments). (For quantification of Western blotting data, see fig. S1C.) (C) Total RNA was prepared from WT and LRP1-deficient macrophages treated with LPS (1 μg/ml) or untreated. Quantitative real-time RT-PCR was performed to assess the expression of ifn-γ. Bars represent the mean of 10 independent experiments; error bars depict SEM. Statistical analysis was performed with Student’s t test; *P < 0.05. For each of the 10 experiments, one conditional k.o. and one control mouse were used. (D) Total RNA was prepared from WT and LRP1-deficient macrophages either treated with LPS (1 μg/ml) or left untreated. Quantitative real-time RT-PCR was performed to assess the expression of ifn-β. Bars represent the mean of six independent experiments; error bars depict SEM. Statistical analysis was performed with Student’s t test; *P < 0.01. For each of the six experiments, one conditional k.o. and one control mouse were used. (E) WT and LRP1-deficient macrophages were treated with LPS (1 μg/ml) for 15 min or were untreated. DNA binding proteins were cross-linked to the chromatin and cell lysates were prepared. After shearing of DNA by sonification, an immunoprecipitation with the C-terminal LRP1 antibody was carried out. The precipitate was used in a PCR reaction with primers binding to the ifn-γ promoter. One experiment representative of three experiments is shown. For each of the three experiments, one conditional k.o. and one control mouse were used. (F) WT and LRP1-deficient macrophages were treated with LPS (1 μg/ml) for the times indicated or were untreated. Chromatin immunoprecipitation was performed as described above. Primers specific for the ifn-β promoter were used in the PCR reaction. One experiment representative of three experiments is shown. For each of the three experiments, one conditional k.o. and one control mouse were used. (G) WT and LRP1-deficient MEFs were treated with leptomycin B (4 ng/ml) for 19 hours or were untreated. Nuclear extracts were prepared and analyzed by Western blotting with an anti–pIRF-3 antibody. β-Actin served as a loading control. One experiment representative of three experiments is shown. (H) WT and LRP1-deficient MEFs were treated with 1 μM epoxomicin (epoxom) or 10 μM lactacystin (lactac) for 19 hours or were left untreated. Nuclear extracts were prepared and analyzed by Western blotting with an anti–pIRF-3 antibody. β-Actin served as a loading control. (I) WT and LRP1-deficient macrophages were treated with LPS (10 ng/ml) for 4 hours or were left untreated. Supernatants were then collected and TNF-α concentrations were determined by ELISA. Bars represent the mean of six independent experiments; error bars depict SEM. Statistical analysis was performed with Student’s t test; *P < 0.005. For each of the six experiments one conditional k.o. and one control mouse were used. (J) WT and LRP1-deficient macrophages were treated with LPS for 24 hours or were left untreated. Supernatants were collected and IL-6 concentrations were determined by ELISA. Bars represent the mean of six independent experiments; error bars depict SEM. Statistical analysis was performed with Student’s t test; *P < 0.05. For each of the six experiments, one conditional k.o. and one control mouse were used.

The opposing NF-κB activation that we observed in lrp1-deficient cells thus likely reflects a counterregulatory response evoked by dysregulated inflammatory signaling, for example, through increased activity of IRF-3. This may represent an adaptive mechanism similar to that reported in tolerant macrophages that are exposed to repeated or prolonged stimulation with LPS (18).

LRP1-ICD facilitates nuclear export of pIRF-3

The LRP1-ICD interacted with both IRF-3 (Fig. 3H) and the promoter of at least one LPS- and LRP1-regulated gene (Fig. 4E). We therefore hypothesized that nuclear accumulation of pIRF-3 in the absence of the LRP1-ICD (Fig. 3B) was the result of reduced inactivation of pIRF-3 rather than increased phosphorylation of IRF-3 because phosphorylation of IRF-3 occurs in the cytosol. The activity of pIRF-3 is controlled by its nuclear export through the shuttling receptor chromosome region maintenance/exportin 1 (CRM1) and its proteasomal degradation (19, 20). To test whether LRP1 modulates either process, we treated WT and LRP1 KO MEFs with the CRM1 inhibitor leptomycin B or the proteasome inhibitors epoxomicin and lactacystin. Inhibition of nuclear export led to the nuclear accumulation of pIRF-3 in WT cells to the same extent as that seen in LRP1 KO cells (Fig. 4G). In contrast, inhibitors of proteasomal degradation led to the progressive accumulation of pIRF-3 in both cell types (Fig. 4H). Together, these results indicate that LRP1-ICD reduced the abundance of active pIRF-3 by facilitating its nuclear export and therefore its shuttling toward proteasomal degradation.

Increased secretion of proinflammatory cytokines by lrp1-deficient macrophages

To assess the physiological impact of the inhibitory mechanism that we have identified, we measured the secretion of cytokines by LPS-stimulated WT and LRP1 KO peritoneal macrophages. LPS-induced production of tumor necrosis factor–α (TNF-α) (Fig. 4I) and IL-6 (Fig. 4J) were both increased in the absence of LRP1, indicating that loss of LRP1-dependent inhibition of the transcription of LPS target genes results in an exaggerated inflammatory response in the affected cells.

Transcriptional profiling of LRP1-dependent LPS-inducible genes

To identify additional LRP1-ICD target genes, we performed a microarray screen of WT and LRP1 KO fibroblasts and of transfected cells expressing various mutant forms of LRP1 (Fig. 3D and fig. S3). Increases or decreases in gene expression in the various cell lines were compared with those in WT fibroblasts, and the data were stratified for genes whose expression was corrected both by transfection of full-length LRP1 (k.o.-LRP1) and the truncated receptor construct containing LRP1-ICD (k.o.-LRP1-β-chain), but not by expression of the LRP1-ECD (k.o.-LRP1-ECD) (table S1). We confirmed the expression patterns of the potential target genes we identified in the cell lines by quantitative real-time reverse transcription polymerase chain reaction (RT-PCR) assays (Fig. 5A). There was good concordance between the data obtained from the microarray assays and those from the RT-PCR assays for all genes tested. Moreover, in the cell line that stably expressed only the free LRP1-ICD (k.o.-LRP1-105), messenger RNA expression of Rsad2, Usp18, Tyki, and Oas1a, all of which are induced during inflammation (2126), were equal to or lower than those in WT cells, which supports the idea that regulation of their transcription is LRP1-dependent (Fig. 5A). We next tested the effect of LPS (2 μg/ml for 3 hours) on these previously unidentified candidate LRP1 target genes. Transcription of all of these genes was increased in response to LPS relative to that in untreated cells (Fig. 5B), which further confirms that LRP1-ICD controls the repression of LPS-inducible genes.

Fig. 5

Identification of a subset of LPS-inducible genes that are repressed by the LRP1-ICD. (A) Total RNA from WT and LRP1-deficient MEFs and from LRP1-deficient fibroblasts stably transfected with LRP1, the membrane-bound LRP1-ICD (LRP-β-chain), the LRP1-ECD, the free LRP1-ICD (LRP1-105), or the empty plasmid vector were analyzed by quantitative real-time RT-PCR for the expression of Usp18, Rsad2, Tyki, and Oas1. Bars represent the mean of six independent experiments; error bars depict SEM. Statistical analysis was performed with Student’s t test; *P < 0.001. (B) WT fibroblasts were treated with LPS (2 μg/ml) for 3 hours or were left untreated. RNA was prepared and examined by real-time RT-PCR for expression of Usp18, Rsad2, Tyki, and Oas1 as described. **P < 0.05, five independent experiments.

Further analysis of WT and LRP1 KO MEFs by unbiased transcriptional profiling yielded a larger set of differentially expressed genes (table S2) that encode proteins with functions in signal transduction, membrane trafficking, and lipid metabolism. Several candidates were detected by Northern blotting analysis, thus confirming the results of the microarray experiments (Fig. 6A). Specificity was verified by transfection of full-length LRP1 cDNA into the LRP1 KO fibroblasts, which normalized gene expression to that of WT cells (Fig. 6B). Transcriptional regulation of these genes likely involves different mechanisms that reflect the multifunctionality of LRP1 and its importance in the regulation of diverse signal transduction pathways (7, 2729).

Fig. 6

LRP1 is involved in multiple signaling pathways. (A) RNA from WT and LRP1-deficient MEFs was analyzed by Northern blotting with cDNA probes for the genes indicated. Cyclophilin served as a loading control. A representative experiment of three experiments is shown. Dkk-3, dickkopf-3; C3, complement factor 3; sfrp-1, secreted frizzled-related protein-1; AHR, aryl hydrocarbon receptor. (B) RNA from WT and LRP1-deficient MEFs and from LRP1-deficient fibroblasts stably transfected with an expression plasmid encoding LRP1 (control: empty plasmid vector) were examined for the expression of 25-cholesterol-hydroxylase. One experiment representative of three experiments is shown.

Discussion

We have uncovered a mechanism by which γ-secretase–dependent proteolytic processing of the lipoprotein receptor LRP1 limits the inflammatory response through nuclear signal modulation and repression of the expression of LPS-inducible genes (Fig. 7). Exposure of cells to LPS led to increased shedding of the LRP1-ECD followed by release of the LRP1-ICD from the plasma membrane. The soluble ICD translocated to the nucleus, where it participated in LPS-induced transcriptional complexes. LRP1-ICD directly interacted with the LPS-activated transcription factor IRF-3 and promoted its nuclear export and proteasomal degradation, thereby limiting its transcriptional activity. As a result, a subset of LPS-inducible genes was repressed by the LRP1-ICD in a negative feedback loop.

Fig. 7

Proposed model for the negative feedback regulation of LPS-induced inflammatory signaling by the γ-secretase–dependent generation of LRP1-ICD. Expression of a subset of LPS-induced genes is activated by IRF-3. Activation of IRF-3 and other LPS-induced signaling pathways leads to increased production of metalloproteases. In addition, LPS-dependent activation of PKC occurs (55), which augments the proteolytical processing of LRP1. Increased shedding provides more substrate for the γ-secretase–mediated cleavage step that leads to release of LRP1-ICD. The LRP1-ICD interacts with IRF-3 and displaces it from its binding to CBP/p300, thereby unmasking its nuclear export signal (yellow) and facilitating its nuclear export. Subsequently, IRF-3-target gene expression is reduced.

Nuclear translocation of LRP1-ICD

We showed by immunofluorescence studies that untagged, recombinant, soluble LRP1-ICD efficiently translocated to the nucleus (Fig. 1A). Cellular fractionation and Western blotting assays revealed the nuclear localization of endogenous LRP1-ICD (Fig. 1B), which suggested that LRP1-ICD might play a role in the regulation of gene transcription. Earlier work from our laboratory and by our colleagues had shown that a tagged LRP1-ICD translocates to the nucleus and modifies transcription from a heterologous reporter (13, 30, 31). However, the tags that were used in those studies can themselves mediate nuclear translocation; thus, the trafficking, localization, and turnover of endogenous LRP1-ICD have remained uncharacterized.

LPS enhances the proteolytic processing of LRP1

Bacterial LPS molecules significantly enhanced the proteolytic processing of LRP1 (Fig. 2A and figs. S5 and S6). After pretreatment with the γ-secretase inhibitor DAPT, the 25-kD C-terminal fragment of LRP1 (LRP1-CTF) that is produced by shedding of the ECD accumulated in macrophages exposed to LPS, which indicated that LPS increased the cleavage of the ECD from LRP1. We reported earlier that this is the rate-limiting step in the production of the LRP1-ICD and that it is followed by further constitutive γ-secretase–dependent intramembranous processing of LRP1 (13, 31).

Shedding of the LRP1-ECD is mediated by a metalloprotease (32), and activation of PKC enhances this extracellular cleavage of LRP1 (13). The activation of PKC, which occurs early in the inflammatory response, induces the production of matrix metalloproteases and of other mediators of inflammation (14), which suggests a possible role for the LRP1-ICD in the inflammatory process. Studies of the effects of LPS, which induces a strong host defense response and activates PKCε among other downstream mediators (33), have shown that the LRP1-ICD is released in the course of the inflammatory response. This allows the LRP1-ICD to function as a feedback regulator that limits damage to normal tissue by unopposed inflammatory signaling. A physiological role for the LRP1-ICD during inflammation is further supported by the γ-secretase–dependent processing of LRP1 in phagosomes, where phagocytosis of foreign material and the inflammatory mediator IFN-γ stimulate release of the LRP1-ICD (12).

LRP1-ICD interacts with IRF-3 and limits its activity by facilitating its nuclear export

In the absence of LRP1, LPS-induced phosphorylation and activation of the transcription factor IRF-3 was prolonged in primary macrophages (Fig. 4B) and pIRF-3 accumulated in the nucleus in MEFs (Fig. 3B). Expression of free LRP1-ICD in LRP1 KO MEFs was sufficient to reduce the abundance of nuclear pIRF-3 to that of WT cells (Fig. 3B), indicating that the ICD directly limited the availability of pIRF-3 in the nucleus independently of the LRP1-ECD. Binding of LRP1-ICD to IRF-3 was mediated through the second NPxY motif (Fig. 3H).

The activity of IRF-3 is determined by its subcellular localization (19). IRF-3 contains both nuclear import and export signals and shuttles between the cytoplasm and the nucleus. Following its phosphorylation by TANK-binding protein kinase (TBK1), it binds to cAMP response element–binding protein (CREB)–binding protein (CBP) or p300, which leads to its nuclear sequestration (19). The activity of IRF-3 is terminated through its nuclear export by CRM1 and its subsequent proteasomal degradation (20). Treatment of WT and LRP1 KO fibroblasts with the CRM1 inhibitor leptomycin B equalized the abundance of nuclear pIRF-3 in both cell types (Fig. 4G). Inhibition of the proteasome, however, led to further accumulation of nuclear pIRF-3 in both WT and LRP1 KO cells (Fig. 4H). Together, these results indicated that the LRP1-ICD restricted transcriptional activation by IRF-3 through facilitating its nuclear export. It is conceivable that the direct interaction between the LRP1-ICD and IRF-3 interferes with the binding of IRF-3 to CBP/p300 and thus unmasks the nuclear export signal, which allows IRF-3 to be shuttled to the cytoplasm by CRM1. Further studies will have to be performed to show whether this regulatory mechanism also applies when IRF-3 is activated by signals other than the LPS-stimulated TLR4 signaling pathway. In a preliminary analysis, we found that treatment of cells with polyinosinic polycytidylic acid (pIpC), a potent activator of IRF-3 that stimulates TLR3, failed to induce proteolytic processing of LRP1 and thus to activate the first branch of the γ-secretase–dependent negative feedback loop (fig. S6). LPS-stimulated generation of LRP1 ICD, on the other hand, allows the ICD to function as a negative feedback regulator of LPS signaling. This indicates some specificity in the regulatory pathway; however, it is still possible that the LRP1-ICD is produced in other inflammatory conditions, thus allowing for a more general role for LRP1-ICD in modulating the activity of IRF-3.

Degradation of the NF-κB inhibitory protein IκB was delayed in the absence of LRP1 (Figs. 3A and 4A). It was recently reported that the LPS-induced transcriptional activity of IRF-3 is dependent on NF-κB (34). If the actions of LPS-induced IRF-3 and NF-κB are coordinated, decreased activation of NF-κB in the absence of LRP1 might occur as a secondary counterregulatory mechanism evoked by increased IRF-3 activity, similar to the defective NF-κB signaling that is observed under conditions of “LPS tolerance,” that is, after repeated or prolonged stimulation of cells with LPS (18). In agreement with this model, transcriptional activation of NF-κB targets by platelet factor 4 and IL-1 is also dependent on LRP1 (35, 36). An additional direct role for LRP1 in the activation of NF-κB also remains possible, but so far, no molecular mechanism has been elucidated. Further support for the indirect model comes from a study in which modulation of NF-κB activity through down-regulation of cell-surface TNF receptor 1 (TNFR1) by LRP1 was reported. This effect seemed to stem partly from a decrease in total TNFR1 and thus might also be mediated through transcriptional regulation by LRP1 (37).

LRP1-ICD inhibits the transcription of genes in the inflammatory response

In experiments with ChIP assays, we showed that LRP1-ICD resided in the transcriptional complex that assembled on the promoter of ifn-γ (Fig. 4E), through which it repressed LPS-induced gene transcription in vivo (Fig. 4C), indicating that γ-secretase–dependent processing of LRP1 mediated transcriptional regulation. In contrast, ChIP assays failed to show LRP1-ICD on the ifn-β promoter, which suggests that only a subset of LPS-inducible genes is modulated by LRP1-ICD. Gene-specific modulation of LPS-induced transcriptional activation through interaction with differentially activated transcriptional cofactors has been described for nuclear receptors (38). Furthermore, chromatin modifications determine gene-specific control of transcription in inflammation (39). In addition, the exact time course of activation of a specific gene relative to the time course of LRP1-ICD release could determine whether modulation of transcription by the ICD occurs. All of these mechanisms, alone or together, could determine the ability of the LRP1-ICD to interact with the transcriptional complex at a given LPS-inducible gene.

Transcriptional profiling of cell lines expressing mutant forms of LRP1 identified several genes whose expression were modified specifically by the presence of LRP1-ICD (table S1). All of the LRP1-ICD targets that we identified in this manner are LPS-inducible genes (Fig. 5B) (2325), indicating that expression of a substantial subset of LPS-regulated genes is modulated by the LRP1-ICD. Increased release of proinflammatory cytokines from LPS-treated LRP1-deficient macrophages showed the physiological impact of the transcriptional control mechanism that we have identified (Fig. 4, I and J).

During our search for LRP1-target genes by transcriptional profiling of WT and LRP1 KO cells, we identified an additional series of differentially expressed transcripts (table S2). It is likely that the expression of many of these genes is altered by indirect mechanisms reflecting the multifunctionality of the receptor (40). For instance, the identification of TGF-β and Wnt signaling-related genes, such as schnurri, latent TGF-β–binding protein, sfrp1, and dkk-3 is consistent with the established role of LRP1 as a co-receptor for TGF-β (41) and with its interaction with Frizzled proteins, the transmembrane receptors for Wnts (42).

Physiological and pathophysiological implications

Regulation of LPS-induced signaling and gene expression represent previously unrecognized roles for γ-secretase and LRP1 as modulators of inflammation. Inflammation is an important pathogenetic factor of atherosclerosis, and the absence of feedback suppression may be responsible in part for the increased atherosclerosis that develops in mice lacking LRP1 in macrophages and in vascular smooth muscle cells (6, 43, 44). The development of atherosclerotic lesions induced by the peroxisome proliferator–activated receptor γ (PPARγ) agonist rosiglitazone was mitigated in conditional knockout animals lacking lrp1 in smooth muscle cells (7); rosiglitazone is also a negative regulator of LPS-inducible inflammatory genes (38, 45). Moreover, in vitro studies showed that the LRP1 ligand apolipoprotein E (apoE) limits inflammatory signaling by IL-1 in smooth muscle cells in an LRP1-dependent manner (35), further supporting a mechanism where LRP1 protects the vascular wall by locally controlling the inflammatory response.

In addition, evidence now shows that cleavage of LRP1 occurs in the penumbra of infarcted cerebral tissue, where it might modulate the occurrence of apoptotic cell death (46), which suggests that LRP1 has a more general role in the control of inflammation. This is potentially relevant to all conditions in which inflammation contributes to pathogenesis. Examples include pulmonary inflammation in the context of short-term lung injury, which is aggravated by TLR4 activation (47), and Alzheimer’s disease, in which inflammation contributes to pathogenesis, whereas polymorphisms of the LRP1 ligand apoE constitute the most important known risk factor for sporadic disease (48). Moreover, control of inflammation by γ-secretase has to be taken into account when considering the therapeutic use of γ-secretase inhibitors, because loss of negative feedback may result in undesirable drug effects.

In summary, our findings define a mechanism by which lipoprotein receptors modulate nuclear signaling, show a role for γ-secretase in inflammation, and uncover an interdependence of lipoprotein receptor and inflammatory signaling. Modulation of the inflammatory response by γ-secretase and LRP1 thus offers a new potential target for therapeutic intervention and may allow conceptually novel treatment in a variety of inflammatory syndromes.

Materials and Methods

Chemicals

LPS (L4391) was purchased from Sigma Aldrich (Germany). rLPS was provided by M. Freudenberg and C. Galanos (MPI for Immunobiology, Freiburg, Germany). pIpC (P1530) was obtained from Sigma Aldrich. DAPT and epoxomicin were obtained from Calbiochem (USA), Leptomycin B was from Biomol (Germany), and lactacystin was obtained from Merck (Germany).

Cell culture and stable transfection of MEFs

MEFs derived from WT or lrp1-deficient mouse embryos were maintained in Dulbecco’s modified Eagle’s medium (DMEM) with glucose (4.5 g/liter), 2 mM l-glutamine (Cambrex, Belgium), penicillin (100 U/ml), streptomycin sulfate (100 μg/ml; Cambrex), and 8% (v/v) fetal calf serum (FCS) (Sigma Aldrich). Cells were transfected with FuGene (Roche, Germany) and stable transfectants were selected with zeocin (Invitrogen, Germany) at a concentration of 700 μg/ml followed by subcloning of emerging colonies. Information on plasmids can be found in the Supplementary Materials.

Immunofluorescence

MEFs or macrophages cultured on glass coverslips were fixed with 4% paraformaldehyde, permeabilized with 0.2% Triton X-100, quenched with 0.1% NaBH4, and blocked with 10% donkey serum and 1% albumin in Tris-buffered saline (TBS), pH 7.4. Incubation with primary antibody [affinity-purified rabbit polyclonal anti-LRP1 (49) or 8G1 mouse monoclonal anti-LRP1, Progen, Germany, or rabbit anti-Iba1, Wako Chemicals, Germany] was performed overnight at 4°C. Alexa 488–labeled donkey anti-rabbit or Alexa 555–labeled anti-mouse antibodies (Invitrogen) were used at a 1:200 dilution for the detection of bound primary antibody. Coverslips were mounted in aqueous mounting medium (Molecular Probes, USA) and examined with a Zeiss Axioplan 2 imaging microscope with ApoTome.

RNA preparation and Northern blot analysis

Total RNA was prepared from MEFs with RNA-STAT60 (Tel-Test, USA). Twenty micrograms of total RNA were used for Northern blotting. Briefly, RNA samples were separated on a 1% agarose, 5.5% formaldehyde gel and transferred to a Hybond N+ membrane (Amersham, USA) by upward capillary transfer in 10× SSC (1.5 M NaCl, 0.15 M sodium citrate, pH 7.0). Nucleic acids were cross-linked to the membrane by UV irradiation. RapidHyb buffer (Amersham) was used for prehybridization and hybridization steps. For hybridization, 1 ng of probe was labeled with [32P]deoxycytidine triphosphate (Amersham) by the Rediprime Labeling System (Amersham). Probe bound to the membrane was detected by autoradiography. Information on probes is provided in the Supplementary Materials.

Microarray experiments

Total RNA from MEFs was prepared with RNA-STAT60 (Tel-Test) or TRIzol reagent (Invitrogen). RNA was used for labeling as described in the Affymetrix technical bulletin (www.affymetrix.com/support/downloads/manuals/expression_analysis_technical_manual.pdf). Hybridization, washing, scanning, and analysis of the Affymetrix GeneChip Murine Genome 430 A 2.0 arrays (Affymetrix, USA) were carried out as described (50) in the core facility of University of Texas Southwestern Medical Center or by the Affymetrix service provider Kompetenzzentrum für Fluoreszente Bioanalytik, Regensburg, Germany. Data obtained from the microarray hybridizations were processed with Microarray Suite 5.0 (Affymetrix) software.

Quantitative real-time RT-PCR

RNA was extracted from MEFs grown to 95 to 100% confluence or from mouse peritoneal macrophages with TRIzol (Invitrogen) and was treated with RNase-Free DNase I (Fermentas, Germany). For cDNA synthesis, random primers (Promega, Germany), M-MLV (Moloney murine leukemia virus) reverse transcriptase (Promega), RNase inhibitor (Promega), and deoxynucleoside triphosphates (Genaxxon, Germany) were used. The real-time PCR reaction was performed with 2× Absolute QPCR SYBR Green Mix with Fluorescein (Abgene) on a single-color real-time PCR detection system (BioRad MyiQ with MyIQ Optical System Software Vers. 1.0). The sequences of the primers used are provided in the Supplementary Materials. The ΔΔCt method was used to compare increased or decreased expression (51). Ct values were standardized with respect to those of cyclophilin or gapdh. For all experiments, samples were assayed at least in duplicate and the mean Ct value was used for further calculations. Experiments were repeated independently at least five times. Data were averaged and statistically analyzed as described in the figure legends.

Preparation of whole-cell lysates and Western blot analysis

Cells were harvested in ice-cold phosphate-buffered saline (PBS) with 1 mM phenylmethylsulfonylfluoride, 1 mM NaVO4, 25 mM β-glycerolphosphate, and 10 mM NaF; RIPA buffer (see Supplementary Materials) was used for lysis. MEF cell lysate (30 μg) or macrophage lysate (50 μg) was subjected to SDS–polyacrylamide gel electrophoresis (PAGE) and Western blotting according to standard procedures. Primary antibodies used are listed in the Supplementary Materials. After incubation with a horseradish peroxidase (HRP)–conjugated secondary antibody, bound antibodies were visualized by enhanced chemiluminescence with SuperSignal CL-HRP Substrate (Perbio, Germany).

Cell fractionation

Cells were harvested in ice-cold PBS and lysed in hypotonic buffer A (see Supplementary Materials). To prepare membrane fractions, lysates were centrifuged at 850g for 5 min at 4°C and the supernatants were centrifuged at 100,000g for 30 min, after which the membrane pellet was resuspended in SDS buffer. For the preparation of nuclear extracts, the pellet from the first centrifugation step was resuspended in buffer C, incubated at 4°C for 60 min, and centrifuged at 100,000g for 30 min. The supernatant was then used for further examination of nuclear proteins by Western blotting analysis.

Generation of GST fusion proteins

GST fusion plasmids of LRP1-ICD mutants were prepared by standard cloning procedures. The LRP1 sequences cloned into pGEX-4T1 (Amersham Pharmacia Biotech) are provided in the Supplementary Materials. Fusion proteins were expressed in BL21-CodonPlus bacteria (Stratagene) after induction by 1 mM isopropyl-thio-d-galactopyranoside for 5 hours. Proteins were recovered by Triton lysis [PBS containing 1% Triton X-100 and Protease Inhibitors (Roche)] and purified with glutathione-agarose beads (Sigma Aldrich, Germany).

GST pull-down assay

MEF lysates were incubated with 50 μl of glutathione-agarose and 10 μg of the respective purified GST fusion protein for 6 hours at 4°C. Glutathione beads were washed rapidly three times in 150 mM NaCl, 10 mM Tris-HCl, pH 7.5, 2 mM MgCl2, 2 mM CaCl2, and 2 mM MnCl2 for 10 min. SDS sample buffer was added to the supernatant or beads. Proteins were separated by SDS-PAGE and analyzed by immunoblotting with an antibody against IRF-3 (Santa Cruz, sc 15991).

LysCre;LRPlox/lox mice

Mice carrying a loxP-marked LRP1 allele were described previously (52). These mice were bred with animals transgenic for the viral Cre recombinase under the control of the lysozyme (lyz1) promoter (53). These mice were provided by I. Förster, Technical University, Munich, Germany. Age-matched 3- to 6-month-old mice were used in all experiments. Animals were kept under standard laboratory conditions and experiments were carried out according to the principles of good laboratory animal care and were approved by the Regierungspräsidium Freiburg, Ref. Veterinärwesen. Details of PCR analyses for genotyping these mice are found in the Supplementary Materials.

Preparation of thioglycollate-elicited peritoneal macrophages

Mice were injected intraperitoneally with 2 ml of thioglycollate solution (3.85 g of LB-thioglycollate in 100 ml of doubly distilled H2O, autoclaved for 30 min at 120°C and 1 bar). After 4 days, macrophages were collected in PBS containing 2% penicillin and streptomycin. After centrifugation for 5 min at 300g at 4°C, cells were resuspended in 500 μl of DMEM containing glucose (1 g/liter), 2 mM glutamine, 8% FCS, and 1% penicillin and streptomycin. Macrophages were seeded as required and were washed three times with prewarmed PBS after 4 hours.

ChIP assays

ChIP analysis was performed with the ChIP Assay Kit (Upstate Cell Signaling Solutions, UK). Briefly, mouse peritoneal macrophages were seeded at 3 × 106 cells per condition. Cells were either left untreated or were stimulated for 15 min with LPS (1 μg/ml). Cells were then harvested and DNA was sheared by sonification with a Branson Digital Sonifier (Model 450) and a 3-mm-diameter microtip probe (two cycles of 10 s with an amplitude of 15%). For each immunoprecipitation, 15 μl of anti-LRP1 serum was used. The following primers were used for the analysis of the ifn-γ promoter (54): Forward, 5′-ATCACCTCCATTGAAGGGCTTCCT-3′; reverse, 5′-AGTTTCCTTTCGACTCCTTGGGCT -3′. PCR conditions were 94°C for 2 min followed by 30 cycles of 94°C for 30 s, 55°C for 30 s, and 68°C for 30 s, and ending with incubation at 68°C for 5 min. Primers used for the detection of the ifn-β promoter were as follows: IFNβ forward, 5′-CCAGCAATTGGTGAAACTGTACAA-3′ and IFNβ reverse, 5′-CAGTGAGAATGATCTTCCTTCATGG-3′.

Cytokine measurements

Mouse TNF-α/TNFSF1A and mouse IL-6 Quantikine enzyme-linked immunosorbent assay (ELISA) kits (R&D Systems) were used to measure the concentrations of the respective cytokines in cell culture supernatants. Briefly, WT or LRP1 KO peritoneal macrophages were seeded into 24-well plates at 3 × 105 cells per well. Supernatants were collected after 4 hours (TNF-α) or 24 hours (IL-6) of stimulation with LPS for ELISA measurements. A FLUOstar Optima plate reader was used for absorbance measurements.

Acknowledgments

We thank S. Zenker and J. Göldner for excellent technical assistance, S. Goerke for help with the cloning of the GST fusion proteins, and M. Kirsch for scientific discussions. We are indebted to M. Frotscher and H. E. Blum for scientific advice and support. This work was supported by the Deutsche Forschungsgemeinschaft [Emmy Noether fellowship MA-2410/1-2 and 1-3 (P.M.), grant BO-1806/2-1 (H.H.B.)], by grants from the NIH [HL20948, HL63762, NS43408 (J.H.)], and by the Humboldt Foundation [Wolfgang Paul Award (J.H.)].

Supplementary Materials

www.sciencesignaling.org/cgi/content/full/1/47/ra15/DC1

Supplementary Materials and Methods

Fig. S1. Quantification of Western blots.

Fig. S2. Detection of LRP1-ECD in transfected LRP1-deficient fibroblasts.

Fig. S3. Schematic representation of the LRP1-ECD mutant transfected into LRP1-deficient fibroblasts.

Fig. S4. Effect of dominant-negative presenilin on the processing of LRP1.

Fig. S5. Time course of DAPT treatment.

Fig. S6. Effect of purified rLPS on the processing of LRP1.

Fig. S7. Analysis of fusion proteins.

Fig. S8. Analysis of macrophages from wild-type and conditional LRP1−/− mice.

Table S1. Effects of LRP1 deficiency on target gene expression.

Table S2. Differential gene expression in wild-type and LRP1-deficient fibroblasts.

References

References and Notes

View Abstract

Navigate This Article