Inflammasome Activation of IL-18 Results in Endothelial Progenitor Cell Dysfunction in Systemic Lupus Erythematosus

Systemic lupus erythematosus (SLE) is an autoimmune disease with varied and often devastating organ involvement and a bimodal mortality pattern that stems from immune mediated organ damage and from an up to 50-fold increase in atherosclerotic cardiovascular (CV) risk, when compared with age- and sex-matched controls. This CV risk cannot be fully explained by traditional Framingham risk factors (1). Furthermore, immunosuppression may decrease CV risk, suggesting that immune dysregulation characteristic of lupus may contribute to premature vascular damage in this disease (2).

Our group has proposed that one mechanism central to the development of premature CV disease (CVD) in SLE is a profound imbalance between vascular damage and vascular repair that leads to endothelial dysfunction and promotes plaque development. We have previously shown that the phenotype and function of bone marrow-derived endothelial progenitor cells (EPCs) and myeloid circulating angiogenic cells (CACs), both crucial cell subsets in vascular repair and maintenance of an intact endothelium (3, 4), are abnormal in SLE patients and in animal models of SLE (5, 6). Indeed, not only are lupus EPC/CAC numbers decreased, but these cells are unable to properly differentiate into mature endothelial cells (EC) in vitro when exposed to proangiogenic stimuli. Furthermore, lupus EPCs/CACs have impaired incorporation into formed vascular structures on a Matrigel assay and decreased capacity to synthesize crucial proangiogenic molecules such as vascular endothelial growth factor (VEGF). In addition, we have recently reported that decreased angiogenesis in SLE is present in vivo, as evidenced by vascular rarefaction and antiangiogenic signatures in lupus tissues (7).

The dysfunction of lupus EPC/CACs is mediated by type I IFN, as normal in vitro EPC/CAC function can be restored in this disease by blocking IFN-α signaling. Furthermore, healthy control EPCs/CACs exposed to IFN-α develop a similar abnormal phenotype to SLE cells (5). In addition, EPCs/CACs isolated from the lupus-prone New Zealand Black/New Zealand White (F₁) mouse, a strain in which type I IFNs play a prominent role in disease development (8), also show a similar aberrant phenotype that correlates with impaired aortic endothelial function (6). Although it is clear that IFN-α hampers vascular repair in SLE, the pathways by which this cytokine exerts its antiangiogenic effects in progenitor cells have only recently begun to be characterized. Our group has reported that one of the mechanisms by which IFN-α mediates EPC/CAC dysfunction is through the downregulation of the proangiogenic molecules IL-1β and VEGF and upregulation of the IL-1 receptor antagonist (IL1-RN). Indeed, addition of activated IL-1β to SLE EPC/CAC cultures restores their capacity to differentiate into mature ECs (7).

Although IL-1β is proangiogenic in SLE, less is known about the role of IL-18, another cytokine also activated via cleavage by caspase-1 (9). IL-18 is a constitutively expressed cytokine that acts as an important bridge between innate immune responses and the development of adaptive immunity. In addition, IL-18 may have an important role in lupus pathogenesis and organ damage. Serum levels of this cytokine are elevated in SLE patients, and this correlates with disease severity and the presence of lupus nephritis (LN) (10–12). Moreover, levels of IL-18 transcript are elevated in glomeruli from patients with LN. This correlates with increased recruitment of plasmacytoid dendritic cells, which express high levels of the IL-18R, to nephritic glomeruli (10).

The inflammasome is a multimolecular platform whose assembly results in rapid activation of caspase-1, the enzyme responsible for generation of the active forms of IL-1β and IL-18. The central components of the inflammasome include a scaffold of the NLR family (NLRP1, NLRP3, or NLRC4) or absent in melanoma-2 (AIM2); an adaptor molecule, apoptosis-associated speck-like protein containing a CARD (ASC); and caspase-1 (13). Regulation of the inflammasome can occur at a transcriptional level, but the presence of an activation signal is the central switch to make this structure functional (13). Activation and assembly of the inflammasome occur downstream of varied stimuli, including cholesterol crystals, uric acid, intracellular bacteria and ATP (13–15).

Type I IFNs have been proposed to have a regulatory role in inflammasome activity, because the IFN-α responsive inflammasome scaffold absent in melanoma-2 (AIM2) activates caspase-1 in an ASC-dependent manner. AIM2 has recently been proposed to be a cytoplasmic receptor for dsDNA, and its mRNA levels are increased in leukocytes of SLE patients (16, 17). Caspase-1 also appears to be regulated by IFNs, because the IFN-regulated transcription factor IRF-1 is essential for caspase-1 transcription and activation of IL-18 in response to IL-12 administration (18, 19).

The role of the inflammasome and IL-18 in CVD has not been well characterized. Increased synthesis of IL-18 by an active inflammasome is seen in proatherosclerotic low-density lipoprotein (LDL) receptor-deficient mice (14). In addition, IL-18 levels are increased in patients with the metabolic syndrome and correlate with arterial stiffness (20). Importantly, endogenous IL-18 may play a major antiangiogenic role in postischemic injury and contribute to decreased EPC numbers in other conditions (21, 22).

Thus, in this study, we investigated the role of inflammasome activity in EPC/CAC dysfunction in SLE, a disease characterized by enhanced type I IFN synthesis and activity and specifically examined the role of inflammasome-driven IL-18 activation in the development of EPC/CAC dysfunction.

The University of Michigan institutional review board approved this study. Subjects gave informed consent in accordance with the Declaration of Helsinki. To obtain peripheral blood, patients fulfilled the revised American College of Rheumatology criteria for SLE (23) and were enrolled from the University of Michigan outpatient Rheumatology clinic. Age- and gender- matched healthy controls were recruited by advertisement. Lupus disease activity was assessed by the SLE Disease Activity Index (SLEDAI) (24). Patient and control demographics and clinical variables for EPC/CAC studies are reported in Table I. For renal microarray analysis, human renal biopsies were collected by the European Renal cDNA Bank (25). A total of 15 pretransplant healthy living donors (LD) and 32 LN patients were processed for microarray analysis. Patient and control demographics and clinical variables for renal microarray studies are available in Supplemental Table I.

Human PBMCs, known to contain both EPCs and CACs (3), were isolated and cultured under proangiogenic stimulation as described previously (7). To summarize, PMBCs contain bone marrow derived EPC and myelomonocytic CAC progenitors that will differentiate into EC when grown on extracellular matrix. These cells have been followed throughout development and have been found to differentiate from progenitors (CD34⁺,CD133⁺, VEGFR2⁺ for EPCs and CD14⁺, CD45⁺, CD115⁺ for CACs) to mature EC, which bind lectin, take up LDL, and express von Willebrand factor, caveolin, and NO synthetase (3, 26). EPC/CACs require the presence of other cells in culture for proper differentiation. Thus, culturing of PBMCs under proangiogenic conditions has become a standard for evaluating EPC/CAC function in vitro. As such, BMCs (2.27 × 10⁶/cm²) were cultured in EC-specific enrichment medium (EBM; Cambrex, East Rutherford, NJ) on fibronectin-coated wells (BD Biosciences, Franklin Lakes, NJ). In some of the experiments, human rIFN-α2b (Merck, Whitehouse Station, NJ), the caspase-1 inhibitor ac-YVAD-cmk (Enzo, Plymouth Meeting, PA), or the caspase-3 inhibitor ac-DEVD-cmk (CPC Scientific, Sunnyvale, CA) were added to the wells at the initiation of the culture, and medium was changed after 3 d and every 72 h thereafter, unless otherwise specified. Some of the molecules added to the culture were replaced with each media change. In experiments to assess the effects of IL-18 on EPCs/CACs, graded concentrations of IL-18 (BioVision, Mountain View, CA), IL-1β (Biovision), or 1 mg/ml of a neutralizing IL-18 Ab (R&D Systems, Minneapolis, MN) were added at the time of cell isolation and left in culture for the first 3 d.

To assess the capacity of peripheral blood EPCs/CACs to differentiate into mature ECs, on day 14 of culture, cells were incubated with 1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (diI)-acetylated LDL (ac-LDL; Biomedical Technologies, Stoughton, MA) and FITC–ulexeuropeus agglutinin (UEA)-1 (Vector Laboratories, Burlingame, CA). To assess EC morphology and expression of endothelial markers, cells were analyzed by fluorescent microscopy using a Leica DMIRB fluorescent inverted microscope (Bannockburn, IL). Images were acquired with a ×100 total magnification, at room temperature, using live cells in Ringer’s buffer. A total of four random fields of view were acquired for every triplicate well and images were analyzed using the CellC program (http://www.cs.tut.fi/sgn/csb/cellc/) to quantify mature ECs, which were considered as those that coexpress UEA-1 and ac-LDL. The numeric aperture for the objective lens of the fluorescent microscope was 0.3. Images were acquired with an Olympus DP30BW camera (Olympus Corporation, Tokyo, Japan) using the acquisition software Olympus-BSW (Olympus). Final processing was done with Adobe Photoshop CS2 (San Jose, CA).

To inhibit IFN-α signaling, the pan-Janus Kinase (JAK) inhibitor 2-(1,1-dimethylethyl)-9-fluoro-3,6-dihydro-7H-benz[h]-imidaz[4,5-f]isoquinolin-7-one (Pyridone 6) or the PI3K inhibitor (5-(4-Fluoro-2-hydroxyphenyl)furan-2-ylmethylene)thiazolidine-2,4-dione (Calbiochem, Gibbstown, NJ) were used. In brief, human EPCs/CACs were cultured under proangiogenic stimulation for 72 h, followed by change of media containing 50 μm pyridone 6, 50 nm PI3K inhibitor, or vehicle (DMSO) for 1 h before addition of 1000 U/ml rIFN-α. EPC/CACs were incubated for 6 additional hours, and then, total RNA was isolated, and real-time PCR was completed.

EPCs/CACs were treated with or without 1000 U/ml of IFN-α or 10 ng/ml LPS O26:B6 (Sigma-Aldrich) overnight, washed once with PBS and lysed in SDS-PAGE buffer. The equivalent of 1 × 10⁶ cells were run on a 15% polyacrylamide gel and transferred to nitrocellulose. Membranes were incubated overnight with rabbit anti-human–caspase-1 (Cell Signaling Technology, Beverly, MA) at 1/1000 final concentration or with anti-β-actin (Cell Signaling Technology) at 1/5000 dilution, followed by HRP-conjugated goat anti-rabbit Ab (The Jackson Laboratory, Bar Harbor, ME) for 1 h at a dilution of 1/10,000. Ab detection was performed using Western Lightening (PerkinElmer, Waltham, MA), following manufacturer’s instructions.

For EPC/CAC studies, total RNA was isolated with Tripure (Roche, Indianapolis, IN) following the manufacturer’s recommendations. For human kidney tissues, cortical tissue segments were manually microdissected into glomeruli and tubulointerstitial compartments as previously published (27, 28). For microarray analysis, RNA was further purified and concentrated using an RNeasy micro kit and following the manufacturer’s recommendations (Qiagen, Valencia, CA). RNA samples were processed on an Agilent 2100 BioAnalyzer (Agilent Technologies, Santa Clara, CA) to assess RNA integrity.

Microarray samples and data results from control and lupus EPCs/CACs in the presence or absence of IFN-α have been previously published by our group (7). The Affymetrix raw data are available in the Gene Expression Omnibus microarray data repository under the reference GSE23203 (http://www.ncbi.nlm.nih.gov/geo).

Gene expression profiling from microdissected human kidney biopsies was performed as previously published using the Human Genome U133A Affymetrix Genechip arrays (Santa Clara, CA). The CEL files were collected and processed using the GenePattern analysis pipeline (www.genepattern.com). CEL file normalization was performed with the Robust Multichip Average method using the mouse and human Entrez-Gene custom CDF annotation from Brain Array version 10 (http://brainarray.mbni.med.umich.edu/Brainarray/default.asp). The normalized files were log₂ transformed. Of the 12,029 human genes, 11,285 and 11,429 were expressed above the 27 Poly-A Affymetrix control expression baseline (negative controls) in the glomerular and tubulointerstitial compartments respectively and were used for further analyses. Normalized data files are in the process of being uploaded on Gene Omnibus (http://www.ncbi.nlm.nih.gov/geo/).

Total RNA was transcribed into cDNA using oligodT and Moloney murine leukemia virus (Invitrogen, Carlsbad, CA) with 1 μg RNA using a MyCycler thermocyler (Bio-Rad, Hercules, CA), and levels of expression of the following genes were measured: caspase-1, 5′-GGA CTC TCA GCA GCT CCT CAG GCA-3′ (forward) and 5′-GCA AAG CTT GAC ATT CCC TTC TGA GCC-3′ (reverse); ASC, 5′-CCT ACG GCG CCG AGC TCA C-3′ (forward) and 5′-CTC CAG AGC CCT GGT GCG T-3′ (reverse); AIM2, 5′-TGG GCA TGC TCT CCT GAG TCC T-3′ (forward) and 5′-TCA GCC TCC TGA TCC CTG GGG C-3′ (reverse); β-actin, 5′-CAT CAC GAT GCC AGT GGT ACG-3′ (forward) and 5′-AAC CGC GAG AAG ATG ACC CAG-3′ (reverse); PRKR, 5′-CTT CCA TCT GAC TCA GGT TT-3′ (forward) and 5′-TGC TTC TGA CGG TAT GTA TTA-3′ (reverse); IFI44, 5′-CTC GGT GGT TAG CAA TTA TTC CTC-3′ (forward) and 5′-AGC CCA TAG CAT TCG TCT CAG-3′ (reverse); and IFIT, 5′-CTC CTT GGG TTC GTC TAT AAA TTG-3′ (forward) and 5′-AGT CAG CAG CCA GTC TCA G-3′ (reverse).

Real-time PCR was carried out using an ABI PRISM 7900HT (Applied Biosystems) with the following cycling conditions: an initial denaturing/activation at 95°C for 10 min, followed by 40 cycles of denaturing at 95°C for 30 s, annealing at 52°C for 30 s, and elongation at 72°C for 30 s. Samples were normalized to the housekeeping gene β-actin, and fold change was expressed as IFN-treated versus untreated or SLE versus control samples.

Serum human IL-18 was quantified by ELISA (eBioscience, Vienna, Austria), following the manufacturer’s instructions.

Patients’ serum was analyzed for autoantibodies via binding to fluoromagnetic beads using the BioPlex 2200 System (Bio-Rad) following the manufacturer’s instructions. Anti-dsDNA was quantified by Farr assay. Autoantibody quantification was performed at the University of Michigan Core Laboratories.

All animal protocols were reviewed and approved by the University of Michigan’s committee on use and care of animals. To evaluate the role of caspase-1 inhibition in lupus-prone mice, 9 mo old New Zealand Mixed (NZM) 2328 or NZM 2328 lacking the type I IFNR (INZM) (obtained initially as a gift from Dr. C. Jacob (University of Southern California, Los Angeles, CA) (29), and subsequently bred at the University of Michigan) were used. For IFN signaling studies, 15-20 wk old NZM (predisease onset) and BALB/c mice (The Jackson Laboratory) were used. Femurs and tibias were washed and flushed with ice-cold MACS buffer (Miltenyi Biotec, Auburn, CA). Mononuclear cells were then isolated on a Histopaque 1083 density gradient (Sigma-Aldrich), and residual RBCs were lysed with RBC lysis buffer (BioLegend, San Diego, CA). Following subsequent wash in PBS, cells were plated onto fibronectin-coated plates (BD Biosciences) at a density of 1 × 10⁶ cells/cm² in EGM-2 Bulletkit media (Lonza, Allendale, NJ) supplemented with 5% heat-inactivated FBS. At the time of plating, vehicle (DMSO) or ac-YVAD-cmk (Enzo, Plymouth Meeting, PA) were added to the media, which was replaced every 3 d. After 6 d in culture, live cells were incubated with FITC-conjugated Bandeiraea (Griffonia) Simplicifolia Lectin I (Isolectin B4) (BS-1; Vector Laboratories) and ac-LDL for 3 h. Cells were imaged as mentioned above for human studies. Mature ECs were designated as those cells costaining with BS-1 and ac-LDL and were quantified in triplicate in four random fields per well of a 48-well plate.

For Affymetrix microarray studies, the significance microarray analysis was used to define genes significantly differentially regulated between the studied groups; a q-value below 0.05 was considered significant. For correlation of EPC numbers with IL-18 and other variables, linear regression was performed. For correlation of the presence of autoantibodies with IL-18 levels, a Mann–Whitney U test was performed. For all other studies, a paired Student t test was performed. A p value <0.05 was considered significant.

All patient and control demographics are listed in Table I. Our group has demonstrated that EPCs/CACs isolated from SLE patients, or the exposure of healthy control EPCs/CACs to recombinant IFN-α, results in their increased apoptosis and decreased ability to differentiate into mature ECs (5). These abnormalities are mediated in part by transcriptional repression of IL-1β and transcriptional upregulation of IL1-RN (7). However, additional analysis of previously published microarray data (7) indicates that, while IFN-α represses IL-1β levels, mRNA of various inflammasome components are significantly upregulated by this cytokine (Table II) in SLE and control patients. With few exceptions, real-time PCR confirmed the microarray analysis. Up to a 5-fold increase of AIM2 mRNA and 3-fold increase in CASP1 mRNA was detected in healthy control or SLE EPC/CACs exposed to recombinant IFN-α (Fig. 1A). In contrast to some of the microarray data, a 3-fold increase in AIM2, PYCARD (ASC), and CASP1 transcripts was observed in untreated EPC/CACs from SLE patients when compared with untreated control cells (Fig. 1B). These increases correlated with the type I IFN signature in each patient analyzed, suggesting that SLE patients with higher basal levels of IFN-α have greater upregulation of both caspase-1 and AIM2 in their EPCs (Fig. 1C). Thus, despite our previous observations that IFN-α treatment of EPC/CACs results in repression of IL-1β pathways, AIM2 and caspase-1 are upregulated. This suggests that inflammasome activity may contribute to IFN-α–mediated effects on EPC/CAC dysfunction. Confirmation of increased induction of caspase-1 at the protein level upon IFN-α exposure was confirmed by Western blot analysis on control early EPCs/CACs (Fig. 1E). A less robust upregulation of caspase-1 in IFN-α–treated lupus EPCs/CACs was noted, which may reflect already increased procaspase-1 protein levels in SLE-untreated lysates, when compared with untreated controls (Fig. 1D). These results indicate that the inflammasome machinery is upregulated in SLE patients and that this phenomenon is mediated, at least in part, by their enhanced exposure to type I IFNs.

We previously reported that repression of IL-1β and upregulation of IL-1 receptor antagonist by IFN-α requires signaling through JAK/STATs, but not through PI3K, downstream of the type I IFN receptor (7). To assess whether the same signaling pathways are involved in upregulation of inflammasome components by type I IFNs, EPCs/CACs from both controls and SLE patients were preincubated with a pan-JAK or PI3K inhibitors prior to stimulation with IFN-α. Real time PCR analysis demonstrated that loss of JAK signaling downstream of type-I IFN receptor completely abrogated the transcriptional upregulation of the inflammasome components AIM2 and CASP1 by IFN-α. In contrast, PI3K inhibition did not alter AIM2 mRNA levels but there was a non-significant trend toward partial inhibition of caspase-1 transcript upregulation (Fig. 1E). These results indicate that the regulation of AIM2 and CASP1 transcripts are a direct downstream result of type I IFN receptor activation and JAK/STAT signaling and that CASP1 transcripts may also be influenced by PI3K activation.

To understand whether the upregulation of caspase-1 in SLE negatively impacts EPC/CAC differentiation into mature ECs, control or lupus EPCs/CACs were cultured in the presence of the caspase-1 inhibitor ac-YVAD-cmk (YVAD) or vehicle for 2 wk under proangiogenic conditions. This inhibitor is able to block formation of the active p20 fragment of caspase-1 (Fig. 2D). As previously reported by us (5), SLE patients display significantly fewer mature ECs after two weeks of culture (Fig. 2A), when compared with healthy controls. Although YVAD treatment did not alter the number of mature ECs in control samples, it significantly increased cell numbers in SLE cultures (Fig. 2A, 2C). These results indicate that inhibition of the central inflammasome enzyme caspase-1 improves the ability of SLE EPCs/CACs to differentiate into mature ECs in culture. In contrast, incubation of control or SLE cells with the caspase-3 inhibitor ac-DEVD-cmk (DEVD) did not improve differentiation (Fig. 2B). These results indicate that inhibition of caspases involved in apoptosis does not significantly impact endothelial differentiation.

As caspase-1 is upregulated by type I IFNs, the requirement for functional type I IFN signaling for the beneficial effects of caspase-1 blockade on these cells was examined. To this effect, lupus prone mice, which have previously been shown to have dysfunctional EPCs, were examined (6). EPC-containing bone marrow cells from lupus prone NZM 2328 mice or from NZM 2328 mice lacking a functional type I IFN receptor (IZNM) were cultured in the presence or absence of YVAD during their proangiogenic differentiation in vitro. As shown in Fig. 2E, YVAD treatment of male and female bone marrow from NZM mice improved their capacity to differentiate into mature ECs, while this effect was not observed in female and male INZM mice (Fig. 2E). These results suggest that only EPCs with functional type I IFN signaling can improve their differentiation capacity through caspase-1 inhibition, suggesting that caspase-1 effects on EPCs lie downstream of IFN-α. The effect of YVAD was not as robust in the mouse bone marrow EPCs as compared with the human samples; this may reflect the fact that the bone marrow compartment could lack CACs which are aberrant in SLE and have been shown to contribute to endothelial differentiation (26).

As the murine experiments indicated that the effect of caspase-1 inhibition in human and murine SLE is dependent on the presence of functional IFN signaling, we examined whether YVAD could inhibit IFN-mediated EPC dysfunction. Control and SLE EPCs/CACs were incubated with IFN-α in the presence or absence of YVAD. As previously reported by our group, IFN-α treatment resulted in ∼50% inhibition of EC numbers in control and SLE cultures (5). Although IFN-αtreatment did not result in detectable active caspase-1 p20, as measured by Western blot (data not shown), the addition of YVAD abrogated the effects of IFN-α on EPC/CAC differentiation (Fig. 3A). In contrast, the caspase-3 inhibitor DEVD did not alter the IFN-mediated EPC/CAC dysfunction (data not shown). Similar findings were obtained when analyzing bone marrow-derived EPCs isolated from non–lupus-prone Balb/c or SLE-prone NZM2328 mice. In both strains, preincubation with YVAD significantly blocked the ability of rIFN-α to disrupt differentiation of EPCs in culture (Fig. 3B). These results indicate that inhibition of caspase-1 disrupts IFN-α-mediated EPC/CAC dysfunction in murine and human systems. Furthermore, although IFN-α does not directly activate caspase-1, its effects require downstream caspase-1 activation for inhibition of EPC/CAC differentiation.

We have previously shown that IL-1β is proangiogenic and is transcriptionally repressed by type I IFNs and in SLE patients. Because inhibition of caspase-1 has positive effects on SLE EPC/CAC differentiation, we hypothesized that this improvement occurred through inhibition of IL-18 production, as previous reports have indicated that IL-18 levels are increased in SLE serum and tissues (12). IL-18 transcripts, assessed by real-time PCR,were upregulated in SLE EPC/CAC. Furthermore, IL-18 levels correlated with type I IFN signatures in each patient (Fig. 1C). These results indicate that, unlike IL-1β levels, which are repressed by exposure to IFN-α (7, 30), IL-18 levels are upregulated by type I IFNs.

Quantification of serum IL-18 in SLE patients demonstrated that the presence of detectable IL-18 was significantly associated with decreased endothelial differentiation capacity of EPCs/CACs (Fig. 4A). In addition, serum IL-18 protein levels significantly associated with the degree of EPC/CAC dysfunction, as assessed by their differentiation into ECs (Fig. 4B). In contrast, IL-18 was not detected in any of the healthy control serum samples analyzed (not shown). IL-18 serum levels or EPC/CAC dysfunction were not associated with lupus disease activity, as assessed by SLEDAI (Fig. 4C, 4D). These results indicate that IL-18 may have detrimental effects on endothelial function, possibly by promoting dysfunction of EPC/CAC differentiation, and this impact of IL-18 on the endothelium is independent of disease activity.

To evaluate whether IL-18 could directly impair EPC/CAC differentiation, healthy control EPC/CAC cultures were incubated with recombinant IL-18. In a dose-dependent manner, IL-18 significantly impaired endothelial differentiation by ∼50% when concentrations equivalent to those seen in SLE serum were used (Fig. 5A, 5B). No further inhibition of endothelial differentiation was detected when higher concentrations of IL-18 were tested (data not shown). In addition, the levels of IL-18 in lupus serum significantly correlated with the ability of caspase-1 inhibition by YVAD to improve endothelial differentiation in EPC/CAC cultures (Fig. 4E). Taken together, these results indicate that inflammasome activation and enhanced synthesis of IL-18 results in EPC/CAC dysfunction.

As IL-18 levels correlate with a decreased differentiation capacity of EPCs/CACs and exogenous IL-18 interferes with differentiation of control EPC/CACs, we determined whether endogenous production of this cytokine in SLE contributes to dysfunction of these cells. By ELISA, IL-18 was not detected in the supernatants of EPC/CAC cultures (data not shown). Thus, a neutralizing Ab to IL-18 was used to disrupt paracrine or low-level production that could be contributing to EPC/CAC dysfunction. Addition of an anti–IL-18 neutralizing Ab, but not an isotype control, to SLE EPCs/CACs for the first 72 h of culture restored normal endothelial differentiation (Fig. 6A, 6B). This improvement was equivalent to improvement seen with neutralization of IFN-α (Fig. 6A), as has been previously demonstrated by our group (5). These results suggest that production of IL-18 by lupus EPC/CAC cultures contributes to their poor differentiation into mature ECs. In contrast, when similar experiments were performed using healthy control EPCs/CACs, IL-18 neutralization did not modify their differentiation capacity (Fig. 6A). These results indicate that upregulation of the inflammasome and IL-18 production in EPC/CAC cultures is a hallmark of SLE cells.

To understand the relationship between the effects of IL-18 versus IL-1β on EPC differentiation, healthy control EPCs/CACs were incubated with rIL-18 in the presence or absence of rIL-1β. Treatment with a combination of both cytokines was able to overcome the inhibitory effects of IL-18 (Fig. 6C). In addition, as mentioned above and as previously reported (7), SLE EPCs/CACs displayed improvement in their differentiation capacity with both IL-1β stimulation and IL-18 blockade, and this effect was increased when a combination of IL-1β and IL-18 neutralization was tested (Fig. 6D). These results suggest that both blockade of IL-18 pathways and activation of IL-1β pathways enhance endothelial differentiation. The effects of both cytokines likely lie on convergent pathways, because IL-1β can overcome the detrimental effects of IL-18. Furthermore, divergent pathways may also be involved, given the enhanced differentiation of SLE EPCs/CACs observed in the presence of IL-18 blockade and concomitant IL-1β stimulation. In SLE, however, the dominant effect appears to be that of inhibitory IL-18 signaling, because IL-1β pathways are repressed (7).

Recent reports indicate that the presence of anti-Ro and anti-dsDNA Abs positively correlates with increased serum IFN-α activity in SLE patients (31). Nucleoprotein-containing immune complexes can stimulate IFN-α production via TLR7 and TLR9 signaling, both of which can serve as priming signals for inflammasome activation (15, 32, 33). As such, we examined whether specific autoantibody production correlated with increased serum IL-18 levels. As shown in Fig. 7A, the presence of an anti-Ro Ab significantly correlated with increased IL-18 levels, whereas other nucleoprotein-containing autoantibodies, such as anti-Smith and anti-RNP did not. Neither the presence (Fig. 7A) nor titers of anti-dsDNA (data not shown) were significantly associated with IL-18 levels. Additionally, the presence of anti-Ro Abs negatively correlated with lupus EPC/CAC differentiation, whereas the presence of anti-Sm and anti-RNP did not, suggesting a putative link among anti-Ro, IL-18 levels, and EPC dysfunction (Fig. 7B).

Given the above findings, evidence for aberrant inflammasome activity was sought in vivo. Others have reported elevated IL-18 protein in class IV and V LN specimens (11). Using microdissected specimens from kidneys from control and LN patients (see Supplemental Table I for patient demographics), microarray analyses was completed and inflammasome components were examined. We observed significant upregulation of PYCARD (ASC), CASP1, and IL-18 transcripts in both glomerular and tubulointerstitial compartments of LN specimens. In addition, glomeruli from nephritis patients also demonstrated significant IL-18R transcript upregulation (Table III). The transcriptional upregulation of PYCARD and CASP1 was more pronounced in World Health Organization classes III and IV nephritis biopsies (data not shown) as compared with class II, suggesting a possible link with more aggressive disease. As we had previously noted vascular rarefaction in glomeruli from LN biopsies (7), and given our findings of the effects of IL-18 on EPC/CAC differentiation, these data suggest that elevated IL-18 in LN may contribute to decreased vascular density in vivo in this disease.

The experiments reported above demonstrate for the first time, to our knowledge, a potential role for inflammasome activation in the development of type I IFN-mediated EPC/CAC dysfunction in SLE. As such, the inflammasome may play a crucial role in the development of aberrant vascular repair and increased CV risk in this disease. Caspase-1 and the inflammasome scaffold AIM2 are upregulated by IFN-α. Specific inhibition of caspase-1 improves EPC/CAC differentiation in murine and human SLE, suggesting that ongoing activation of this inflammasome-associated enzyme contributes to dysfunction of cells that are crucial in vascular repair. It appears that the deleterious effects of caspase-1 activation in EPC/CAC function are mediated by enhanced IL-18 activity, as neutralization of this cytokine restores lupus EPC/CAC differentiation capacity. This is further confirmed by the observed induction of healthy control EPC/CAC dysfunction by exogenous IL-18, as well as by the association of increased circulating levels of this cytokine with EPC dysfunction in SLE. Overall, these observations indicate a model in which the effects of IFN-α on EPC/CACs are complex and result in skewing of inflammatory cytokine production to favor increased IL-18 activation and repression of IL-1β, which may lead to profound dysfunction in vasculogenesis (Fig. 8). Given the important role of EPCs/CACs in vascular repair and CV disease prevention (reviewed in Ref. 3), these abnormalities may be crucial in the development of premature vascular disease in SLE.

The regulation of caspase-1 and AIM2 by functional JAK activation is consistent with current dogma regarding activation of the type I IFNR (34). In addition, the trend of inhibition of caspase-1 activation by IFN-α in the presence of PI3K inhibitors suggests an additional role for the antiviral IRS-dependent regulation of caspase-1.

IL-18 is an inflammatory cytokine with an emerging role in endothelial dysfunction and atherosclerosis. Its levels have previously been shown to correlate with decreased circulating EPC numbers in Graves’ disease (22). Additionally, elevated levels of IL-18 correlate with increased vascular stiffness and intima media thickness in men without evidence of coronary artery disease, indicating that this cytokine mayplay a role in early stages of atherosclerosis development (35). Further, IL-18 levels correlate with instability of atherosclerotic plaque and strongly serve as a predictor of CV death in patients with subclinical atherosclerosis (36, 37). Certainly, its effects on ECs could be proposed to be proatherosclerotic and proinflammatory, as this cytokine also upregulates IL-6, IL-8, and adhesion molecules including ICAM (9). Thus, we propose that in SLE, chronic dysregulation of inflammasome activity, possibly induced by increased circulating IFN-α levels, results in elevated IL-18 that leads to downstream detrimental effects on vascular repair.

In other model systems, such as inflamed synovium, IL-18 has a proangiogenic role (38). These discrepancies of IL-18 effects on the vasculature are not necessarily contradictory. Indeed, many of the cytokines upregulated by IL-18 can in turn be proangiogenic in highly inflammatory environments, increase inflammatory cell migration and alter endothelial repair, which may also promote accelerated atherosclerosis development.

The cellular source of IL-18 in SLE remains unknown, but likely candidates would include monocytes, macrophages, and other myeloid cells that represent the classic source of inflammasome-produced cytokines (13). In EPC/CAC cultures, myeloid cells are present, especially early on, and may be the source of continued IL-18 production in vitro. This hypothesis would agree with previous findings that active IL-18 colocalizes with mononuclear phagocytes in atherosclerotic plaques, while the endothelium is rich in the IL-18R (39).

Our observations suggest that an environment characterized by enhanced type I IFN levels and activity, such as seen in SLE, skews the inflammasome machinery toward IL-18 over IL-1β activation,through alteration of substrates available to the inflammasome. First, IL-1β is tightly regulated and IFN-α exposure leads to significant transcriptional downregulation of this cytokine in human and murine systems (7, 30, 40). Second, IL-18 is more constitutively expressed than IL-1β (9), and we have shown that IL-18 transcript levels positively correlate with type I IFN signatures in SLE patients. Thus, we can propose a model where, under the influence of type I IFNs, IL-1β transcript and protein are repressed while IL-18 is upregulated. In this environment, activation of caspase-1 will result in preferential processing of IL-18 as the cytokine most available to the inflammasome machinery. Although others have proposed that alternative proteases can also cleave and activate IL-18 (41–43), our data suggest that caspase-1 inhibition improves differentiation of EPC/CAC cultures, supporting the inflammasome as the trigger of enhanced production of this cytokine in SLE. If the cellular source of IL-18 is indeed monocytes, a cell subset known to have constitutively active caspase-1 (44), then it is possible that upregulation of this enzyme by IFN-α may contribute significantly to processing and activation of IL-18 in SLE.

Additional steps may also contribute to the activation of the inflammasome in SLE. One possibility for inflammasome activation may be the production of neutrophil-associated extracellular traps (NETs). Lupus neutrophils are capable of releasing NETs containing DNA/antimicrobial peptide complexes that enter plasmacytoid dendritic cells and stimulate TLR9 (45). In addition, low-density granulocytes, which produce abundant NETS in vivo (46) and are found in PBMC fractions in SLE, but not in control patients (47), externalize the cathelicidin LL37 and other immunostimulatory molecules through NETosis. LL37 can stimulate inflammasome activation via induction of the P2X7 receptor (48). Priming of the inflammasome in SLE could occur via signaling through TLR7 or TLR9 via immune complex uptake (15, 33). TLR7 is typically expressed in plasmacytoid dendritic cells, but its expression is inducible in monocytes (49). As such, signaling through TLR7 could contribute to enhanced IL-18 production in the presence of immune complexes, both via induction of IFN-α synthesis in plasmacytoid dendritic cells and inflammasome priming in monocytes. In addition, as the presence of IFN-α increases neutrophil netting in response to autoantibodies (50), this could explain the increased association with elevated IL-18 levels in the presence of anti-Ro Abs. Given the link we report between the presence of anti-Ro Abs, IL-18 levels and EPC/CAC dysfunction, it is possible that Ro-containing immune complexes prime monocytes to activate the inflammasome. Further research into the mechanisms involved in this potential phenomenon should be performed.

In addition, the identification of the inflammasome adaptors involved in EPC dysfunction needs to be characterized. Others have shown that the NLRP3 inflammasome is repressed by type I IFNs in bone marrow-derived macrophages, in an IL-10-dependent fashion (30). Thus, in environments with high type I IFN levels, such as in SLE, the AIM2 inflammasome may be the more relevant scaffold molecule. AIM2 is upregulated by type I IFNs and is activated by dsDNA. Others have shown increased AIM2 transcript in peripheral leukocytes from SLE patients, but no increase was found in nephritis biopsies (16). We have now confirmed that IFN-α regulates AIM2 transcript in EPC/CAC cultures. This suggests that the AIM2 inflammasome may be relevant to increased IL-18 production in SLE. Future studies will address the role of various inflammasome components in SLE-related vascular disease.

The upregulation of inflammasome components in LN biopsies suggests that inflammasome activation of IL-18 in the kidney may play a pathogenic role. Little is known about the role of caspase-1 in LN, but IL-18 has been shown to possibly play a pathogenic role in both humans and mice (11, 12, 51, 52). Vascular rarefaction, which is demonstrated in SLE but not in ANCA-associated vasculitis nephritis biopsies (7), and has been associated with progression of renal disease (53), may be reflective of aberrant vasculogenesis in LN. Given our data showing the role of caspase-1 in EPC/CAC dysfunction in SLE and the detrimental effects of IL-18 on endothelial differentiation, we speculate that inflammasome activity in LN may contribute to vascular rarefaction at the level of glomeruli and renal blood vessels and contribute to progressive renal failure. Future studies should address the role of the inflammasome components in mediating LN, and given our data on caspase-1 regulation by IFN-α, further consideration should be given to models in which type I IFNs play a role.

In conclusion, we have demonstrated a novel role for the inflammasome in contributing to EPC/CAC dysfunction, and potentially to premature vascular damage, in SLE. Enhanced levels or sensitivity to type-I IFNs may skew the inflammasome toward production of IL-18 by repression of IL-1β and upregulation of caspase-1 and AIM2. Given its effects on EPC/CAC function, enhanced IL-18 may play a deleterious role in vascular repair mechanisms in SLE and contribute to accelerated CVD, progression of kidney damage and premature mortality in this disease.

We thank Srilakshmi Yalavarthi for excellent technical assistance, Luigi Franchi for helpful discussions, and Caroline Mohai, Marnie Hyzy, Alyssa Yang, and physicians from the Division of Rheumatology at the University of Michigan for assistance in recruitment of patients for this study. We also thank Dr. Chaim Jacob for providing breeding pairs of NZM and INZM mice. In addition, we thank Dr. Anna Henger for the processing of the human renal biopsies and the University of Michigan Microarray Core Facility (Cancer Center) for human DNA chip hybridizations. Members of the European Renal cDNA Bank-Kroener-Fresenius biopsy bank at the time of the study: Clemens David Cohen, Holger Schmid, Michael Fischereder, Lutz Weber, Matthias Kretzler, Detlef Schlöndorff, Munich, Germany/Zurich, Switzerland/Ann Arbor, MI/New York, NY; Jean Daniel. Sraer, Pierre Ronco, Paris; Maria Pia Rastaldi, Giuseppe D’Amico, Milano, Italy; Peter Doran, Hugh Brady, Dublin, Ireland; Detlev Mönks, Christoph Wanner, Würzburg, Germany; Andrew Rees, Aberdeen, Scotland/Vienna, Austria; Frank Strutz, Gerhard Anton Müller, Göttingen, Germany; Peter Mertens, Jürgen Floege, Aachen, Germany; Norbert Braun, Teut Risler, Tübingen, Germany; Loreto Gesualdo, Francesco Paolo Schena, Bari, Italy; Jens Gerth, Gunter Wolf, Jena, Germany; Rainer Oberbauer, Dontscho Kerjaschki, Vienna, Austria; Bernhard Banas, Bernhard Krämer, Regensburg, Germany; Moin Saleem, Bristol, U.K.; Rudolf Wüthrich, Zurich, Switzerland; Walter Samtleben, Munich, Germany; Harm Peters, Hans-Hellmut Neumayer, Berlin; Mohamed Daha, Leiden, The Netherlands; Katrin Ivens, Bernd Grabensee, Düsseldorf, Germany; Francisco Mampaso, Madrid; Jun Oh, Franz Schaefer, Martin Zeier, Hermann-Joseph Gröne, Heidelberg, Germany; Peter Gross, Dresden, Germany; Giancarlo Tonolo, Sassari, Italy; Vladimir Tesar, Prague, Czech Republic; Harald Rupprecht, Bayreuth, Germany; Hermann Pavenstädt, Münster, Germany; Hans-Peter Marti, Bern, Switzerland; and Peter Mertens, Magdeburg, Germany.

The authors have no financial conflicts of interest.