Rheumatoid arthritis (RA) is characterized by autoimmune joint destruction with debilitating consequences. Despite treatment advancements with biologic therapies, a significant proportion of RA patients show an inadequate clinical response, and restoration of immune self-tolerance represents an unmet therapeutic need. We have previously described a tolerogenic phenotype of plasmacytoid dendritic cells (pDCs) in RA patients responding to anti–TNF-α agents. However, the molecular mechanisms involved in tolerogenic reprogramming of pDCs in RA remain elusive. In this study, guided by transcriptomic analysis of CD303+CD123+ pDCs from RA patients in remission, we revealed enhanced expression of IL-6R and its downstream signaling compared with healthy pDCs. Functional assessment demonstrated that IL-6R engagement resulted in marked reduction of TNF-α secretion by pDCs whereas intracellular TNF-α was significantly increased. Accordingly, pharmacologic inhibition of IL-6R signaling restored TNF-α secretion levels by pDCs. Mechanistic analysis demonstrated impaired activity and decreased lysosomal degradation of ADAM17 (a disintegrin and metalloproteinase 17) sheddase in pDCs, which is essential for TNF-α cleavage. Importantly, reduction of TNF-α secretion by IL-6–treated pDCs attenuated the inflammatory potential of RA patient–derived synovial fibroblasts. Collectively, these findings position pDCs as an important source of TNF-α in RA pathogenesis and unravel an anti-inflammatory mechanism of IL-6 by limiting the pDC-derived TNF-α secretion.

Rheumatoid arthritis (RA) is an autoimmune disease characterized primarily by chronic synovial joint inflammation leading to articular destruction with debilitating consequences (1). It is now accepted that the immunopathogenesis of the disease starts with the development of autoreactive T and B cells to posttranslationally modified proteins in genetic-susceptible individuals, which after an asymptomatic period of systemic autoimmunity contribute to synovial tissue inflammation. Eventually, synovial stromal cells are converted to autoaggressive effector cells, contributing thus to the development of chronic synovial inflammation (2). Apart from the established knowledge of the importance of inflammatory cytokines and cell-signaling pathways in the pathogenesis of diseases, novel high-throughput technologies have revealed the essential contribution of peripheral and synovial cell subpopulations in the pathogenesis of diseases (35). During the last decades we have witnessed the advent of novel biologic therapies mostly targeting inflammatory cytokines (such as inhibitors of TNF-α, IL-6R, and IL-1β), T or B cells (such as CTLA4Ig and anti-CD20), or small-molecule inhibitors of intracellular signaling (JAK-STATs) (6). Nevertheless, these agents have essentially anti-inflammatory properties, although therapies to re-establish self-tolerance are not available. Although biologics are clinically effective agents, up to 35% of patients treated with TNF-α inhibitors achieve remission (710), and numbers are comparable for non-TNF inhibitors such as anti–IL-6R (tocilizumab) (11). The molecular and cellular mechanisms contributing to clinical failure of potent biologic agents are not well understood.

Plasmacytoid dendritic cells (pDCs) are a unique cell population that bridges innate and adaptive immune responses, characterized by the expression of the type II C-lectin receptor CD303 and the IL-3R α-chain CD123 (12). They are responsible for the substantial production of type 1 IFN, present Ags, and induce naive T cell differentiation (13, 14). They also secrete an array of proinflammatory cytokines including TNF-α and IL-6 (15) or anti-inflammatory molecules such as IL-10 and IDO (16, 17), all of which contribute to the decision of immunity versus tolerance. Regarding the latter, it has been shown that pDCs exhibit strong tolerogenic functions by inducing CD8+ T cell deletion, CD4+ T cell anergy (18), and regulatory T cell (Treg) differentiation (1921). In the context of autoimmunity, aberrant activation of pDCs has been implicated in the pathogenesis of autoimmune inflammatory diseases such as systemic lupus erythematosus (22) and psoriasis (23), whereas tolerogenic skewing of pDC activation has been linked to achievement of remission in RA (24). Interestingly, pDC ablation induces recruitment of myeloid-derived suppressor cells, facilitating re-establishment of tolerance in the mouse model of multiple sclerosis (25). In RA, mature pDCs from RA patients with low disease activity express high levels of IDO and promote the differentiation of naive CD4+CD25 T cells into IL-10–secreting Tregs or T regulatory type-1 cells (24). Moreover, in vivo depletion of pDCs exacerbates joint pathology and augments cell-mediated and humoral immunity to type II collagen in experimental arthritis (26). However, the molecular pathways involved in prompting pDCs as a tolerogenic cell population in RA remain elusive.

In this study, guided by transcriptomic analysis of pDCs from RA patients in remission, we reveal an anti-inflammatory function of IL-6 on pDCs through limiting the secretion of TNF-α. This was accompanied by increased pDC accumulation of intracellular TNF-α and decreased activity of ADAM17 (a disintegrin and metalloproteinase 17) sheddase. Notably, reduction of TNF-α secretion by IL-6–treated pDCs attenuated the inflammatory potential of RA synovial fibroblasts (RASFs) as indicated by the expression of proinflammatory mediators. Collectively, to our knowledge, these results describe a novel negative feedback loop mechanism comprising IL-6 and pDCs for dampening inflammatory responses through reduction of TNF-α, which could be therapeutically exploited.

Patients and healthy individuals were recruited through the Rheumatology and Blood Donation Department, University Hospital of Heraklion. The study was approved by the Ethics Committee of the University Hospital of Heraklion (no. 1476, March 20, 2012), and informed consent was obtained from all patients and healthy individuals prior to sample collection.

Peripheral blood samples were obtained from individuals diagnosed with RA or from healthy individuals. Subjects with RA fulfilled the 2010 European League Against Rheumatism/American College of Rheumatology classification criteria and were followed up at the Rheumatology Clinic of the University Hospital of Heraklion (27) (clinical characteristics and treatments are shown in Supplemental Table I). At the time of sampling, all patients were treated with anti–TNF-α agents and classified either as being in remission/low disease activity (disease activity score of 28 joint counts [DAS28] < 3.2) or in active disease state (DAS28 ≥ 3.2) (28). Healthy age- and sex-matched volunteers from the Department of Transfusion Medicine of the University Hospital of Heraklion served as controls.

Heparinized blood (20 ml) was collected from healthy subjects and individuals with RA. Additionally, buffy coats were used from healthy donors. PBMCs were isolated by density-gradient centrifugation (Lymphosep). Erythrocytes were eliminated by hypotonic lysis. Viability was measured at 99% by trypan blue dye exclusion. The pDC population was magnetically isolated using the CD304 (BDCA-4/neuropilin-1) human MicroBead kit (Miltenyi Biotec), and purity was evaluated by flow cytometry (CD303+ and CD123+ cells/total live cells). Preparations of ≥85% purity were used in all experiments.

Total RNA was isolated from pDCs using the Qiagen RNeasy mini kit. For global transcriptome analyses, Affymetrix HG-U133 Plus 2.0 gene arrays (Affymetrix, Santa Clara, CA) were used. cRNA generation and array hybridization were done according to the instructions of the manufacturer (Affymetrix). Finally, 15 µg of cRNA was used for array hybridization. Microarray data analysis included data normalization and the generation of cell files by the Affymetrix GeneChip operating software. Cell files were uploaded in the BioRetis database (http://www.bioretis-analysis.de) for pairwise comparisons and the extraction of differentially expressed genes (29). The microarrays were deposited in ArrayExpress at the European Molecular Biology Laboratory–European Bioinformatics Institute (https://www.ebi.ac.uk/arrayexpress/) under accession number E-MTAB-11933 (https://www.ebi.ac.uk/arrayexpress/experiments/E-MTAB-11933). Heatmaps and hierarchical clustering was performed in the Biomedical Sciences Research Center “Alexander Fleming” (Athens, Greece) using the Morpheus application (https://software.broadinstitute.org/morpheus/). For protein-to-protein interaction, the network STRING database (PMID: 27924014) was used.

Magnetically isolated healthy pDCs (2.5 × 105) from buffy coats were plated in 96-well plates and stimulated with CpG-A (0.5 μM) and rIL-6 (100 ng/ml) for the indicated time points. In addition, to inhibit ADAM17 activity and thereby the release of cell-surface proteins, pDCs were preincubated with ADAM17/TNF-α–converting enzyme (TACE) inhibitor TAPI-1 (20 μM) for 30 min and then stimulated with CpG-A and rIL-6 for 18 h. Culture supernatants and cells were collected and stored at −80°C for further analysis. For all experiments, pDCs were cultured in RPMI 1640 supplemented with 10% FBS, 100 IU/ml penicillin, and 100 μg/ml streptomycin and incubated at 37°C and 5% CO2.

The pDC cell line CAL-1 (provided by B.R.) was grown in RPMI 1640 supplemented with 10% FBS, 100 IU/ml penicillin, 100 μg/ml streptomycin, and 1× nonessential amino acids. Then, 2 × 105 CAL-1 cells were plated in 96-well plates and stimulated with CpG-B (6.3 μM) and rIL-6 (100 ng/ml) for the indicated time periods (1.5, 3, and 5 h) before analyses.

For inhibition of the IL-6 signaling pathway, 2 × 105 CAL-1 cells were plated in 96-well plates and treated with the IL-6R antagonist tocilizumab (50 μg/ml, Roche) or LMT-28 (100 μΜ, Sigma-Aldrich), which targets the IL-6Rβ subunit gp130. After 30 min, cells were activated with CpG-B in the presence or absence of rIL-6 for 3 h. Supernatants were collected and TNF-α was measured using an ELISA kit (Invitrogen).

For lysosomal inhibition, 7 × 104 CAL-1 cells were seeded in coverslips or 2 × 105 CAL-1 cells were plated in 96-well plates. Cells were pretreated with chloroquine (CQ, 50 μM, Sanofi Aventis) or the autophagy inhibitor 3-methyladenine (3-MA, 10 μM, Sigma-Aldrich) for 30 min followed by activation with CpG-B for 3 h. Cells in coverslips were further analyzed by immunofluorescence. Supernatants from 96-well plates were collected and TNF-α was measured using an ELISA kit (Invitrogen).

RASFs (provided by C.O.) were isolated from RA patients’ synovial tissues (obtained from joint replacement surgery after informed consent with ethics approval from the local Ethics Committee). Patients fulfilled the 2010 classification criteria for RA (27) (Supplemental Table II). For cell culture, tissue samples were digested and RASFs were grown as previously described (30). Cultures of RASFs were subjected to experimental procedures at passages 4–9. RASFs were cultured for 48 h in a six-well plate at a density of 105 cells/well and then treated with freshly isolated pDCs or CAL-1 culture supernatants for 24 h. Finally, RASFs were collected and tested for MMP1, MMP3, IL-6, and CXCL8 mRNA levels.

For the ADAM17 activity assay, the SensoLyte 520 TACE activity assay kit (AnaSpec, Fremont, CA) with the fluorogenic peptide OXLTM 520/5-FAM (ADAM17 cleavage site from TNF-α) was used. Then, 2 × 105 CAL-1 cells/condition were seeded in a black 96-well plate with Opti-MEM (Thermo Fisher Scientific, Swindon, U.K.). CAL-1 cells were pretreated with TAPI-1 for 30 min and then CpG-B in the presence or absence of IL-6, and the peptide substrates were added to cultures. In the presence of TACE, the peptide was cleaved, and fluorescence of the cleaved substrate, which was directly proportional to the amount of TACE activity, was measured from baseline until 3 h after the addition of the treatments and substrate. Fluorescence measurement was performed at 528 nm (excitation, 480 nm) at 37°C (Victor X2 plate reader, PerkinElmer).

Single-cell suspensions were prepared and cells were stained for extracellular markers for 20 min at 4°C in 5% FBS/PBS. Fluorochrome-conjugated Abs against human CD303 and CD123 (BioLegend) were used to identify the pDC population. Anti-HLA-DR–, anti-CD80–, anti-CD86–, anti-CD40–conjugated Abs (BioLegend) were used as maturation/activation markers. Additionally, cells were stained with mAbs specific for membrane IL-6R (BioLegend), and LysoTracker (Invitrogen) was used to label acidic organelles in the cells. For intracellular phosphorylated protein staining, cells were fixed with 1.5% formaldehyde, permeabilized with ice-cold methanol (31), and stained with conjugated Abs against human p-STAT1 and p-STAT3 (BioLegend). Samples were acquired on a FACSCalibur (BD Biosciences) and analyzed using FlowJo software (Tree Star).

For immunofluorescence, pDCs and CAL-1 cells were seeded in coverslips pretreated with poly-l-lysine (7–9 × 104 cells/slide). Cells were activated with CpG in the presence or absence of rIL-6 for 1.5–4 h. Then, slides were washed with PBS, fixed with 4% paraformaldehyde for 20 min at room temperature, rewashed three times with PBS, permeabilized with 0.5% Triton X-100 for 1 min, and blocked with PBS containing 1% BSA for 30 min. Cells were then stained with rabbit anti-ADAM17 (2 μg/ml, Atlas Antibodies) and adalimumab (Humira,1:100, AbbVie), followed by incubation with Alexa Fluor 555 anti-rabbit IgG (1:500, Molecular Probes) and Alexa Fluor 488 anti-human IgG (1:500, Molecular Probes). To monitor autophagy and lysosomal function, cells were stained with rat anti-lysosomal-associated membrane protein 1 (Lamp, 1:400, Abcam), rabbit anti-p62 (1:500, MBL International), and mouse transcription factor EB (TFEB, 1:100, R&D Biosystems) followed by incubation with CF555 anti-rat IgG (1:400, Biotium), CF488A anti-rabbit IgG, or anti-mouse IgG (1:500 or 1:100, Biotium). For visualization of the nuclei DAPI (Sigma-Aldrich) was used. Samples were coverslipped with Mowiol and visualized using the Leica SP8 inverted confocal live cell imaging system. Intensity per cell was calculated using the Leica software. Puncta per cell were calculated using a macro developed in Fiji software as described (32).

CAL-1 cells were lysed in RIPA buffer (50 mM Tris-HCl at pH 8, 150 mM NaCl, 0.5% sodium deoxycholate, 1% Nonidet P-40, and 0.1% SDS) supplemented with protease and phosphatase inhibitors (cOmplete, EDTA-free; Roche Applied Science). Equal amounts of proteins (40 μg) were subjected to SDS-PAGE electrophoresis on 8% gels and then transferred to Immobilon-PSQ membranes (Millipore). Membranes were blocked with 5% skimmed milk, 1% BSA in TBST and then incubated with anti-ADAM17 (Atlas Antibodies) and anti-tubulin Ab (1:5000, Millipore) as a loading control. Detection was performed using HRP-linked Abs, anti-rabbit HRP (1:8000) and anti-mouse HRP (1:3000) (Cell Signaling Technology), respectively. The images were resolved by an ECL system (Pierce and Millipore) and detected by ImageBlot (Bio-Rad). Relative intensity of bands was calculated using Image Lab analysis software.

Total RNA from cells was collected using the TRIzol extraction protocol or mini RNA isolation kit (Invitrogen) with Turbo DNAse (Ambion) treatment according to the manufacturer’s instructions. cDNA was prepared using a PrimeScript first-strand cDNA synthesis kit (Takara) according to the manufacturer’s protocol. Transcripts were quantified by incorporation of KAPA SYBR FAST qPCR kit master mix (2×) (Kapa Biosystems) with a Bio-Rad CFX Connect real-time system. Genes expression was normalized to GAPDH and calculated by the change-in-threshold method (2−ΔΔCt). Primer sequences used were as follows: IFN-α forward, 5′-GGTGACAGAGACTCCCCTGA-3′, reverse, 5′-CAGGCACAAGGGCTGTATTTCTT-3′; IL-6R forward, 5′-ACATTCACAACATGGATGG-3′, reverse, 5′-AGGACTCCTGGATTCTGTC-3′; TNF-α forward, 5′-GAGGCCAAGCCCTGGTATG-3′, reverse, 5′-CGGGCCGATTGATCTCAGC-3′; ADAM17 forward, 5′-GACTCTAGGGTTCTAGCCCAC-3′, reverse, 5′-GGAGACTGCAAACGTGAAACAT-3′; GAPDH forward, 5′-CATGTTCCAATATGATTCCACC-3′, reverse, 5′-GATGGGATTTCCATTGATGAC-3′; CXCL8 forward, 5′-ACTGAGAGTGATTGAGAGTGGAC-3′, reverse, 5′-AACCCTCTGCACCCAGTTTTC-3′; IL-6 forward, 5′-CAGATGAGTACAAAAGTCCTGA-3′, reverse, 5′-CTACATTTGCCGAAGAGCCC-3′; MMP1 forward, 5′-CTCGCTGGGAGCAAACACATC-3′, reverse, 5′-CTGCTTGACCCTCAGAGACCT-3′; MMP3 forward, 5′-GGACCTGGAAATGTTTTGGCCC-3′, reverse, 5′-ACCCAGGGAGTGGCCAATTT-3′.

Detection of human TNF-α and IL-6R (Invitrogen) in freshly isolated pDCs and CAL-1 culture supernatants harvested at the indicated time points was performed by sandwich ELISA following the manufacturer’s recommendations (Invitrogen). Light absorbance at 450 nm was measured using an ELx800 BioTek plate reader. All samples were assessed in duplicates.

All experiments were repeated at least three times, and representative data are shown. A nonparametric Mann–Whitney U test or one-way ANOVA was applied in experiments, and all data were analyzed using GraphPad Prism v8 software. Differences were considered statistically significant at p < 0.05.

Previous studies demonstrated that CD303+CD123+ pDCs exhibit a tolerogenic phenotype and contribute to remission in anti-TNF-α–treated RA patients (24, 26). However, the molecular mechanisms underlying the contribution of pDCs in RA remission are poorly defined. To address this, pDCs were isolated from peripheral blood of RA patients in remission and healthy individuals and subjected to gene expression analysis; however, pDCs from active RA patients were excluded from this study due to very low frequencies (Fig. 1A) (33). Interestingly, transcriptomic analysis of pDCs revealed >6000 coding transcripts to be differentially expressed in RA patients compared with healthy controls (data not shown), suggesting an intense transcriptomic reprogramming. Hierarchical clustering, using Genes@Work with Pearson correlation and center of mass, pointed to an enrichment in biological regulation, metabolic, and immune system pathways (Fig. 1B). Of interest, significant clusters of cytokine/chemokine and transcription factor transcripts were enriched in pDCs from the two groups (Fig. 1C).

FIGURE 1.

Transcriptional reprogramming of CD303+CD123+ pDCs in RA patients responding to therapy. (A) Gating and number of CD303+CD123+ cells in PBMCs (3 × 105 cells) from healthy donors (n = 10), RA patients in the active disease state (n = 19), and patients responding to therapy (remission) (n = 22). Healthy donor versus RA active, *p = 0.041; RA active versus RA remission, *p = 0.0137. ns, not significant. Results are expressed as mean ± SEM. Statistical significance was obtained by one-way ANOVA, followed by a post hoc pairwise test: F(2,48) = 3.87, *p = 0.027. (B) Top 16 biological processes enriched in the set of deregulated genes and their associated p values. (C) Heatmap of the differentially expressed genes related to cytokine and transcription factor group of genes. (D) Network of deregulated molecules on the IL-6 signaling pathway. (E) Representative histograms and IL-6R expression levels gated on pDCs from healthy donors (n = 9) and RA patients in remission (n = 11). *p = 0.01 (unpaired t test). (F) Representative histograms and expression levels of STAT1, STAT3, p-STAT1, and p-STAT3 gated on pDCs from healthy donors (n = 4) and RA patients in remission (n = 4). *p = 0.02 (unpaired t test). Dot plots represent geometric mean fluorescence intensity (gMFI). For (D), red nodes represent the upregulated genes and blue nodes the downregulated genes in RA. Node size has been adjusted according to the differential expression level.

FIGURE 1.

Transcriptional reprogramming of CD303+CD123+ pDCs in RA patients responding to therapy. (A) Gating and number of CD303+CD123+ cells in PBMCs (3 × 105 cells) from healthy donors (n = 10), RA patients in the active disease state (n = 19), and patients responding to therapy (remission) (n = 22). Healthy donor versus RA active, *p = 0.041; RA active versus RA remission, *p = 0.0137. ns, not significant. Results are expressed as mean ± SEM. Statistical significance was obtained by one-way ANOVA, followed by a post hoc pairwise test: F(2,48) = 3.87, *p = 0.027. (B) Top 16 biological processes enriched in the set of deregulated genes and their associated p values. (C) Heatmap of the differentially expressed genes related to cytokine and transcription factor group of genes. (D) Network of deregulated molecules on the IL-6 signaling pathway. (E) Representative histograms and IL-6R expression levels gated on pDCs from healthy donors (n = 9) and RA patients in remission (n = 11). *p = 0.01 (unpaired t test). (F) Representative histograms and expression levels of STAT1, STAT3, p-STAT1, and p-STAT3 gated on pDCs from healthy donors (n = 4) and RA patients in remission (n = 4). *p = 0.02 (unpaired t test). Dot plots represent geometric mean fluorescence intensity (gMFI). For (D), red nodes represent the upregulated genes and blue nodes the downregulated genes in RA. Node size has been adjusted according to the differential expression level.

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Guided by the transcriptomic analysis, together with the mechanisms implicated in RA pathogenesis, a prominent module of deregulated genes involved in IL-6 signaling was observed (Fig. 1D). Indeed, in line with the transcriptomic results, IL-6R expression levels were significantly increased in pDCs from RA patients in remission compared with healthy controls (Fig. 1E). Furthermore, expression of PSTAT1 and PSTAT3 and their phosphorylated proteins in pDCs from RA patients revealed increased expression levels of STAT1, p-STAT1, and p-STAT3, but not STAT3 (Fig. 1F), also confirming the transcriptomic analysis data (Fig. 1D). Collectively, these findings demonstrate an extensive reprogramming of pDCs in RA patients in remission characterized by engagement of IL-6R–mediated signaling.

IL-6 is known to contribute to RA pathogenesis, and although IL-6R blocking constitutes a therapeutic target in RA (34), a significant proportion of RA patients still do not achieve remission, suggesting the existence of unknown mechanisms that contribute to therapeutic resistance (11). Moreover, the role of the IL-6/IL-6R signaling pathway in pDC function remains elusive. To address this, healthy pDCs were activated with CpG in the presence of rIL-6. Assessment of type I IFNα mRNA expression, a hallmark molecule of pDCs, showed no significant differences between IL-6–treated and nontreated cells (Fig. 2A). In addition, flow cytometry analysis showed no alterations on HLA-DR, CD80/CD86, and CD40 expression by IL-6–treated pDCs as compared with nontreated cells (Fig. 2B), suggesting that IL-6 does not impact the Ag-presenting properties of pDCs. Next, focusing on the transcriptomic data, we identified genes implicated in TNF-α production to be significantly deregulated, whereas expression of the TNFα gene was markedly decreased in pDCs from RA patients (Fig. 3A). Thus, considering that TNF-α is heavily involved in RA pathogenesis, constituting a major therapeutic target, we assessed TNF-α protein levels in IL-6–treated pDC culture supernatants. Notably, we found markedly decreased secretion of TNF-α in supernatants from IL-6–treated pDCs compared with supernatants from control pDCs (Fig. 3B). Of interest, confocal microscopy demonstrated that IL-6–treated pDCs were highly decorated with TNF-α, suggesting an IL-6–mediated impaired TNF-α cleavage (Fig. 3C). Strikingly, a similar pattern of TNF-α expression was observed in ex vivo–isolated pDCs from RA patients in remission (Fig. 3D). Finally, IL-6 treatment of CAL-1 leukemic cells, which share phenotypic and functional features with pDCs, including TNF-α–producing capacity (35), showed increased p-STAT1 and p-STAT3 signaling (Fig. 3E) and significantly reduced TNF-α secretion in culture supernatants (Fig. 3F). In addition, TNF-α was significantly accumulated on IL-6–treated CAL-1 cells compared with control-treated cells (data not shown). Overall, these results indicate that IL-6 restrains the secretion of TNF-α by activated pDCs, although increased TNF-α levels accumulated in IL-6–treated pDCs, possibly due to impaired cleavage.

FIGURE 2.

No significant differences in IFNα mRNA levels and in surface activation markers between IL-6–treated and nontreated pDCs. (A) IFNα mRNA levels expressed by CpG-activated pDCs after 4.5- and 13-h treatment in the presence or absence of rIL-6 (n = 3–5 different experiments). ns, not significant. (B) Healthy CpG-activated pDCs treated with rIL-6 for 18 h (n = 3 different experiments). Expression levels of HLA-DR, CD80, CD86, and CD40 were evaluated by flow cytometry. Representative FACS histograms are shown. ns, not significant. Results are expressed as mean ± SEM (unpaired t test) and as fold over the untreated pDCs.

FIGURE 2.

No significant differences in IFNα mRNA levels and in surface activation markers between IL-6–treated and nontreated pDCs. (A) IFNα mRNA levels expressed by CpG-activated pDCs after 4.5- and 13-h treatment in the presence or absence of rIL-6 (n = 3–5 different experiments). ns, not significant. (B) Healthy CpG-activated pDCs treated with rIL-6 for 18 h (n = 3 different experiments). Expression levels of HLA-DR, CD80, CD86, and CD40 were evaluated by flow cytometry. Representative FACS histograms are shown. ns, not significant. Results are expressed as mean ± SEM (unpaired t test) and as fold over the untreated pDCs.

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FIGURE 3.

IL-6R signaling diminishes TNF-α production by pDCs. (A) Network of deregulated molecules implicated in TNF-α production. (B) TNF-α levels in culture supernatants of healthy CpG-activated pDCs in the presence or absence of rIL-6 for 13 h (n = 4 different experiments) and 18 h (n = 8 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-activated pDCs. *p = 0.02, ***p = 0.0009. (C) Immunofluorescence confocal microscopy for TNF-α (green) and DAPI (blue) on isolated CpG-activated pDCs treated in the presence or absence of rIL-6 for 4 h. Representative fields. Scale bars, 5 μm. One representative experiment of 6 is shown. TNF-α intensity/cell is depicted. **p=0.001. (D) Immunofluorescence confocal microscopy for TNF-α (green) and DAPI (blue) on ex vivo isolated pDCs from healthy donor and RA patient in remission. Representative fields are shown. Scale bars, 5 μm. One representative experiment of six is shown. TNF-α intensity/cell is depicted. ****p < 0.0001. (E) Representative histograms and expression levels of p-STAT1 and p-STAT3 produced by CpG-activated CAL-1 cells in the presence or absence of rIL-6 for 10 min (n = 4 different experiments). *p = 0.028. Dot plots represent geometric mean fluorescence intensity (gMFI). (F) TNF-α levels in culture supernatants of healthy CpG-activated CAL-1 cells in the presence or absence of rIL-6 for 18 h (n = 5 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-activated CAL-1 cells. **p = 0.0079. For (A), red nodes represent the upregulated genes and blue nodes the downregulated genes. Node size has been adjusted according to the differential expression level. (B–D) ns, not significant. For (B)–(F), results are expressed as mean ± SEM (unpaired t test). For (C) and (D), 30–50 pDCs were examined per experiment.

FIGURE 3.

IL-6R signaling diminishes TNF-α production by pDCs. (A) Network of deregulated molecules implicated in TNF-α production. (B) TNF-α levels in culture supernatants of healthy CpG-activated pDCs in the presence or absence of rIL-6 for 13 h (n = 4 different experiments) and 18 h (n = 8 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-activated pDCs. *p = 0.02, ***p = 0.0009. (C) Immunofluorescence confocal microscopy for TNF-α (green) and DAPI (blue) on isolated CpG-activated pDCs treated in the presence or absence of rIL-6 for 4 h. Representative fields. Scale bars, 5 μm. One representative experiment of 6 is shown. TNF-α intensity/cell is depicted. **p=0.001. (D) Immunofluorescence confocal microscopy for TNF-α (green) and DAPI (blue) on ex vivo isolated pDCs from healthy donor and RA patient in remission. Representative fields are shown. Scale bars, 5 μm. One representative experiment of six is shown. TNF-α intensity/cell is depicted. ****p < 0.0001. (E) Representative histograms and expression levels of p-STAT1 and p-STAT3 produced by CpG-activated CAL-1 cells in the presence or absence of rIL-6 for 10 min (n = 4 different experiments). *p = 0.028. Dot plots represent geometric mean fluorescence intensity (gMFI). (F) TNF-α levels in culture supernatants of healthy CpG-activated CAL-1 cells in the presence or absence of rIL-6 for 18 h (n = 5 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-activated CAL-1 cells. **p = 0.0079. For (A), red nodes represent the upregulated genes and blue nodes the downregulated genes. Node size has been adjusted according to the differential expression level. (B–D) ns, not significant. For (B)–(F), results are expressed as mean ± SEM (unpaired t test). For (C) and (D), 30–50 pDCs were examined per experiment.

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Cleavage of the membrane-bound TNF-α precursor to its mature soluble form is mediated by the sheddase ADAM17 (3638). Of note, analysis of the transcriptomic data showed decreased expression of ADAM17 in pDCs from RA patients responding to treatment as compared with healthy pDCs (data not shown). Thus, we sought to examine whether reduction of TNF-α production in IL-6–treated pDCs is due to deregulated expression and/or function of ADAM17. First, we treated pDCs with TAPI-1, an ADAM17-specific inhibitor (39), and observed a significant reduction of soluble TNF-α in culture supernatants, confirming that ADAM17 catalyzes the release of TNF-α from pDCs (Fig. 4A). Thus, we assessed ADAM17 activity via quantification of a fluorogenic peptide cleaved by ADAM17, and our results showed significantly reduced activity of ADAM17 in IL-6–treated CAL-1 cells compared with control treated cells (Fig. 4B). To further support the diminished ADAM17 activity in IL-6–treated pDCs, we examined the shedding of IL-6R, another substrate of the sheddase. To this end, measurement of soluble IL-6R concentration by ELISA revealed significantly decreased levels in supernatants from IL-6–treated cells compared with those derived from control cells (Fig. 4C). Of interest, confocal microscopy revealed increased intracellular accumulation of ADAM17 in IL-6–treated pDCs (Fig. 4D), and this finding was confirmed in IL-6–treated CAL-1 cells (Fig. 4E). In support, Western blot analysis for ADAM17 and tubulin in CpG-activated CAL-1 lysates verified the increased expression of ADAM17 in the presence of rIL-6 (Fig. 4F).

FIGURE 4.

IL-6 inhibits ADAM17 sheddase activity, which mediated the cleavage of TNF-α by pDCs. (A) Expression levels of TNF-α in culture supernatants of healthy CpG-activated pDCs pretreated for 1 h in the presence or absence of ADAM17 inhibitor, TAPI-1 (n = 8 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-treated pDCs for 18 h. ***p < 0.0001. (B) ADAM17 shedding activity of CpG-activated CAL-1 cells measured after a 3-h treatment with rIL-6 (n = 6 different experiments). *p = 0.01. (C) Expression levels of IL-6R in culture supernatants of healthy CpG-activated pDCs treated in the presence or absence of rIL-6 (n = 6 different experiments), expressed as fold change over the levels of IL-6R produced by CpG-treated pDCs for 18 h. **p = 0.002. (D) Immunofluorescence confocal microscopy for ADAM17 (green) and DAPI (blue) on CpG-activated pDCs treated in the presence or absence of rIL-6 for 4 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of four is shown. ADAM17 intensity/cell is depicted. *p = 0.019. (E) Immunofluorescence confocal microscopy for ADAM17 (red) and DAPI (blue) on CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 for 4 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of six is shown. ADAM17 intensity/cell is depicted. *p = 0.013. (F) Western blot analysis for expression of ADAM17 and α-tubulin in CAL-1 lysates of indicated groups (n = 5/group). One representative experiment of five is depicted. Relative expression of ADAM17 is depicted. *p = 0.02. Results are expressed as mean ± SEM (unpaired t test). For (D) and (E), 30–50 cells were examined per experiment.

FIGURE 4.

IL-6 inhibits ADAM17 sheddase activity, which mediated the cleavage of TNF-α by pDCs. (A) Expression levels of TNF-α in culture supernatants of healthy CpG-activated pDCs pretreated for 1 h in the presence or absence of ADAM17 inhibitor, TAPI-1 (n = 8 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-treated pDCs for 18 h. ***p < 0.0001. (B) ADAM17 shedding activity of CpG-activated CAL-1 cells measured after a 3-h treatment with rIL-6 (n = 6 different experiments). *p = 0.01. (C) Expression levels of IL-6R in culture supernatants of healthy CpG-activated pDCs treated in the presence or absence of rIL-6 (n = 6 different experiments), expressed as fold change over the levels of IL-6R produced by CpG-treated pDCs for 18 h. **p = 0.002. (D) Immunofluorescence confocal microscopy for ADAM17 (green) and DAPI (blue) on CpG-activated pDCs treated in the presence or absence of rIL-6 for 4 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of four is shown. ADAM17 intensity/cell is depicted. *p = 0.019. (E) Immunofluorescence confocal microscopy for ADAM17 (red) and DAPI (blue) on CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 for 4 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of six is shown. ADAM17 intensity/cell is depicted. *p = 0.013. (F) Western blot analysis for expression of ADAM17 and α-tubulin in CAL-1 lysates of indicated groups (n = 5/group). One representative experiment of five is depicted. Relative expression of ADAM17 is depicted. *p = 0.02. Results are expressed as mean ± SEM (unpaired t test). For (D) and (E), 30–50 cells were examined per experiment.

Close modal

Taken together, these findings reveal that IL-6 signaling in pDCs attenuates TNF-α production and induces intracellular ADAM17 accumulation, an evidence that may lead to the impairment of ADAM17 activity.

It is established that ADAM17-mediated shedding takes place at the cell surface, and upon internalization ADAM17 can be either recycled or undergoes lysosomal degradation (40). Because our findings indicated increased ADAM17 accumulation intracellularly in IL-6–treated CAL-1 cells, we asked whether IL-6 could regulate ADAM17 lysosomal degradation in pDCs. To this end, immunofluorescence microscopy of CpG-activated CAL-1 cells in the presence of CQ, a blocker of lysosomal function, revealed increased intensities of ADAM17 and TNF-α, whereas using 3-MA, a PI3K inhibitor and autophagy inhibitor, no significant differences in ADAM17 and TNF-α expression were observed (Fig. 5A). Additionally, TNF-α protein levels were significantly reduced in the supernatants of CpG-treated CAL-1 cells in the presence of CQ but not 3-MA, corroborating the findings observed with IL-6–treated cells (Fig. 5B). Interestingly, assessment of lysosomal pH in IL-6–treated CAL-1 cells, via measuring the accumulation of a specific pH-sensitive probe LysoTracker, demonstrated increased pH compared with control cells, indicating a dysfunctional lysosome (Fig. 5C). Furthermore, immunofluorescence microscopy of IL-6–treated CAL-1 cells revealed increased expression of the Lamp-1 and the adaptor protein sequestosome-1 (SQSTM1 or p62) that targets ubiquitinated proteins for lysosomal degradation (41, 42), as compared with CpG-treated CAL-1 cells (Fig. 5D). In line with this, expression of TFEB in IL-6–treated CAL-1 cells was markedly increased, confirming the presence of compensatory mechanisms operating during lysosomal dysfunction (43) (Fig. 5E). Collectively, our findings suggest that IL-6 regulates the lysosomal function in CAL-1 cells, which impairs the degradation of ADAM17, and this mechanism may regulate trafficking of ADAM17 to the cell surface to exert its sheddase activity.

FIGURE 5.

Impaired lysosomal function and defective ADAM17 clearance in IL-6–treated CAL-1 cells. (A) Immunofluorescence confocal microscopy for ADAM17 (red), TNF-α (green), and DAPI (blue) on CpG-activated CAL-1 cells treated with chloroquine (CQ) or 3-methyladenine (3-MA) for 3 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of three is shown. ADAM17 and TNF-α intensities/cell are depicted. *p = 0.02, ****p < 0.0001. (B) Expression levels of TNF-α in culture supernatants of CpG-activated CAL-1 pretreated for 30 min in the presence or absence of CQ or 3-MA (n = 9 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-treated CAL-1 for 4 h. ****p < 0.0001. ns, not significant. (C) Representative histogram and MFI of LysoTracker Red of CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 (n = 5 different experiments) expressed as fold change over the levels of LysoTracker Red of CpG-treated CAL-1 cells for 3 h. *p = 0.015. (D) Immunofluorescence confocal microscopy for p62 (green), Lamp-1 (red), and DAPI (blue) on CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 for 3 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of three is shown. Lamp-1 (**p = 0.0047) and p62 (*p = 0.011) puncta/cell are depicted. (E) Immunofluorescence confocal microscopy for TFEB (green) and DAPI (blue) on CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 for 3 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of three is shown. TFEB puncta/cell are depicted (***p = 0.0001). Results are expressed as mean ± SEM. Statistical significance was obtained by a one-way ANOVA (A and B) or unpaired t test (C–E). For (A), (D), and (E), 30–50 CAL-1 cells were examined per experiment.

FIGURE 5.

Impaired lysosomal function and defective ADAM17 clearance in IL-6–treated CAL-1 cells. (A) Immunofluorescence confocal microscopy for ADAM17 (red), TNF-α (green), and DAPI (blue) on CpG-activated CAL-1 cells treated with chloroquine (CQ) or 3-methyladenine (3-MA) for 3 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of three is shown. ADAM17 and TNF-α intensities/cell are depicted. *p = 0.02, ****p < 0.0001. (B) Expression levels of TNF-α in culture supernatants of CpG-activated CAL-1 pretreated for 30 min in the presence or absence of CQ or 3-MA (n = 9 different experiments), expressed as fold change over the levels of TNF-α produced by CpG-treated CAL-1 for 4 h. ****p < 0.0001. ns, not significant. (C) Representative histogram and MFI of LysoTracker Red of CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 (n = 5 different experiments) expressed as fold change over the levels of LysoTracker Red of CpG-treated CAL-1 cells for 3 h. *p = 0.015. (D) Immunofluorescence confocal microscopy for p62 (green), Lamp-1 (red), and DAPI (blue) on CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 for 3 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of three is shown. Lamp-1 (**p = 0.0047) and p62 (*p = 0.011) puncta/cell are depicted. (E) Immunofluorescence confocal microscopy for TFEB (green) and DAPI (blue) on CpG-activated CAL-1 cells treated in the presence or absence of rIL-6 for 3 h. Representative fields are shown. Scale bars, 5 μm. One representative experiment of three is shown. TFEB puncta/cell are depicted (***p = 0.0001). Results are expressed as mean ± SEM. Statistical significance was obtained by a one-way ANOVA (A and B) or unpaired t test (C–E). For (A), (D), and (E), 30–50 CAL-1 cells were examined per experiment.

Close modal

Taking into consideration the pivotal role of TNF-α in RA pathogenesis, we sought to assess the biological significance of the IL-6–dependent decreased production of TNF-α by pDCs. RASFs are the most common cell type in the inflamed synovium, and they contribute to joint destruction through production of cytokines, chemokines, and matrix-degrading molecules (44, 45). Thus, we first exposed SFs derived from RA patients to TNF-α–enriched supernatants of CpG-activated primary pDCs, in the presence or absence of adalimumab, a therapeutic monoclonal TNF-α blocker. We found significantly decreased expression of MMP1, MMP3, IL6, and IL8 in RASFs upon TNF-α neutralization compared with control-treated pDCs (Fig. 6A). Importantly, supernatants from IL-6–treated primary pDCs significantly inhibited the expression of MMP1, MMP3, and IL8 proinflammatory mediators from RASFs (Fig. 6A), whereas no significant differences in IL6 expression levels were observed, suggesting that IL-6–dependent attenuation of TNF-α secretion by pDCs exerts anti-inflammatory properties on RASFs. Accordingly, supernatants of CpG-activated CAL-1 cells reduced transcription levels of these genes upon TNF-α neutralization compared with non anti-TNF-α–treated or recombinant human TNF-α–treated CAL-1 cells, whereas no significant differences in IL6 expression levels were observed (Fig. 6B).

FIGURE 6.

pDC-derived TNF-α exerts proinflammatory functions in RA patient–derived fibroblasts. (A) MMP1, MMP3, IL6, and IL8 mRNA levels expressed by RASFs after a 24-h treatment with supernatants from healthy CpG-activated pDCs (pDC sup) in the presence or absence of rIL-6 or adalimumab (n = 5 different experiments). Values for each corresponding molecule are expressed as fold change over the mRNA levels produced by the pDC sup–treated RASFs. MMP1, **p = 0.006; MMP3, *p = 0.03 and 0.01, ****p < 0.0001; IL6, *p = 0.01, ***p = 0.0004. ns, not significant; IL8, *p = 0.03 and 0.01, **p = 0.002. (B) MMP1, MMP3, IL6, and IL8 mRNA levels expressed by RASFs after a 24-h treatment with supernatants from CpG-activated CAL-1 cells (CAL-1 sup) in the presence or absence of an anti–TNF-α agent, adalimumab (n = 4 different experiments). Values for each corresponding molecule are expressed as fold change over the mRNA levels produced by the untreated RASFs. rTNF was used as positive control. MMP1, *p = 0.02; MMP3, *p = 0.02, ***p = 0.0002; IL8, *p = 0.04, ****p < 0.0001. ns, not significant. (C) Expression levels of TNF-α in culture supernatants of healthy CpG-activated CAL-1 cells pretreated for 1 h with TCZ or LMT-28 (IL-6R blocker) and cultured in the presence or absence of rIL-6 for 3.5 h. Values are expressed as fold change over the control of the respective treatment (n = 5 different experiments). **p = 0.0079. ns, not significant. Results are expressed as mean ± SEM. Statistical significance was obtained by a one-way ANOVA (A and B) or unpaired t test (C).

FIGURE 6.

pDC-derived TNF-α exerts proinflammatory functions in RA patient–derived fibroblasts. (A) MMP1, MMP3, IL6, and IL8 mRNA levels expressed by RASFs after a 24-h treatment with supernatants from healthy CpG-activated pDCs (pDC sup) in the presence or absence of rIL-6 or adalimumab (n = 5 different experiments). Values for each corresponding molecule are expressed as fold change over the mRNA levels produced by the pDC sup–treated RASFs. MMP1, **p = 0.006; MMP3, *p = 0.03 and 0.01, ****p < 0.0001; IL6, *p = 0.01, ***p = 0.0004. ns, not significant; IL8, *p = 0.03 and 0.01, **p = 0.002. (B) MMP1, MMP3, IL6, and IL8 mRNA levels expressed by RASFs after a 24-h treatment with supernatants from CpG-activated CAL-1 cells (CAL-1 sup) in the presence or absence of an anti–TNF-α agent, adalimumab (n = 4 different experiments). Values for each corresponding molecule are expressed as fold change over the mRNA levels produced by the untreated RASFs. rTNF was used as positive control. MMP1, *p = 0.02; MMP3, *p = 0.02, ***p = 0.0002; IL8, *p = 0.04, ****p < 0.0001. ns, not significant. (C) Expression levels of TNF-α in culture supernatants of healthy CpG-activated CAL-1 cells pretreated for 1 h with TCZ or LMT-28 (IL-6R blocker) and cultured in the presence or absence of rIL-6 for 3.5 h. Values are expressed as fold change over the control of the respective treatment (n = 5 different experiments). **p = 0.0079. ns, not significant. Results are expressed as mean ± SEM. Statistical significance was obtained by a one-way ANOVA (A and B) or unpaired t test (C).

Close modal

Finally, CAL-1 cells were pretreated with IL-6R blockers prior to their activation. For that, we used tocilizumab (RoActemra), a recombinant humanized anti-human IL-6R mAb, or the IL-6R blocker LMT-28, which binds to the IL-6R β subunit gp130, and then activated CAL-1 cells in the presence of rIL-6. Notably, assessment of TNF-α in culture supernatants demonstrated that blocking of IL-6 signaling restored the TNF-α protein levels, indicating a direct role of IL-6/IL-6R in the suppression of TNF-α release by CAL-1 cells (Fig. 6C). Taken together, these findings demonstrate an important contribution of pDC-derived TNF-α in RA inflammatory processes and highlight the regulatory role of IL-6 in this process.

pDCs have a unique role in the interplay between innate and adaptive immune responses, and accumulating evidence during the last years highlights that pDCs acquire either a regulatory or immune-activating phenotype in autoimmune diseases. Thus, it is important to understand the molecular mechanisms that underlie the pDC duplicitous phenotype in autoimmunity, which may assist in the design of rational therapies. In this study, to our knowledge, we describe a novel mechanism, consistent with the disease-protective function of pDCs in RA patients, that entails an IL-6–mediated inhibition of TNF-α secretion by pDCs. Specifically, pDCs from RA patients in remission upon treatment with TNF-α inhibitors showed an enhanced transcriptomic signature of IL-6 signaling, and ex vivo treatment of pDCs with IL-6 attenuated the functional activity of the sheddase ADAM17, which catalyzes the cleavage and release of TNF-α. Mechanistically, IL-6 signaling in pDCs deregulated the lysosomal function, which mediates ADAM17 degradation and recycling. Importantly, our findings place pDCs as an important TNF-α–producing cell subset in RA and provide insights into mechanisms that may contribute to disease remission. Whether the herein described changes on pDCs upon achieving clinical remission are mediated through a specific/direct effect of the TNF-α inhibitor on pDCs or through the systemic control of inflammation cannot be addressed by our data. Moreover, a differential molecular/cellular effect of the specific agents cannot be excluded. Although TNF-α inhibitors share many common characteristics and biologic activities, they have differences in biologic effects, clinical efficacy, and safety (46).

The anti-inflammatory properties of IL-6 have been characterized in other inflammatory conditions. IL-6 was shown to suppress IL-1β and TNF-α by PBMCs (47) or to induce IL-1Ra and a soluble TNF receptor (TNFsRp55) by macrophages (48). These in vitro findings were confirmed in animal models of endotoxic lung or endotoxemia by using IL-6 knockout mice (49). Moreover, ablation of IL-6R by hepatocytes resulted in massive hepatic inflammation and diminished insulin resistance (50), whereas IL-6Rα deletion from myeloid cells increased insulin-targeted tissue inflammation and rendered animals more susceptible to LPS-induced endotoxemia (51). Accordingly, our findings reveal that IL-6 signaling diminishes TNF-α secretion by pDCs, affecting thus the inflammatory potential of RASFs.

Interestingly, the clinical implication of our results may contrast the therapeutic benefit of IL-6R targeting seen in patients with rheumatic diseases. However, note that still a significant proportion of patients treated with IL-6R inhibitor fail to achieve remission (11), implying the existence of unrecognized mechanisms of therapy resistance. The molecular mechanisms responsible of clinical failure of IL-6R blocking in RA and other inflammatory diseases are generally unexplored. In this regard, it has been shown that tocilizumab may enhance IL-6Rα expression and soluble IL-6Rα secretion by DCs, which may acquire a proinflammatory profile following tocilizumab treatment (52). Our data raise the possible scenario that blocking IL-6R signaling may sustain TNF-α production by pDCs, representing a mechanism of unresolved inflammation, failure of achieving clinical remission, or promoting flares of the disease. Whether this mechanism is also involved in the induction of Th1 and Th17 immune responses that have been shown to mediate disease in RA patients (53) and respective mouse models (54) remains to be seen.

Our findings indicate that diminished release of TNF-α by IL-6–treated pDCs is facilitated through inhibition of ADAM17 activity. Of importance, decreased expression of ADAM17 was detected in pDCs ex vivo isolated from the peripheral blood of RA patients. The active form of ADAM17 has been demonstrated in cartilage and synovial tissue as well as in chondrocytes of RA and osteoarthritis patients (55), and exposure of synovial cells to low oxygen in vitro increased ADAM17 expression and enhanced TNF-α release (56). In support, inhibition of ADAM17 has been shown of great efficacy in mouse (57) and rat models of arthritis (58). ADAM17 expression levels followed by internalization and lysosomal degradation constitute steps of the life cycle of this protease, and distinct inflammatory stimuli were shown to regulate this life cycle differently (40). In line with this, our results demonstrate that IL-6 signaling impairs lysosomal function in pDCs, resulting in accumulation of ADAM17 due to defective degradation, and this is accompanied by increased accumulation of TNF-α. Whether lysosomal dysfunction is involved only in trafficking or also affects the functional activity of ADAM17 is an area of active investigation. The question raised by our findings is how IL-6 signaling could regulate lysosome functionality in pDCs. Cytokine signaling has been shown to control lysosomal features in diverse cellular settings (59), although STATs interfere with the lysosomal pH through association with H+-ATPase (60). Furthermore, IL-6 regulates autophagy operation and autophagolysosomal function (61, 62); however, blocking of initial steps in canonical autophagy pathway did not affect the lysosomal elements in pDCs. Therefore, delineating the mechanisms affecting the expression and function of ADAM17 and also understanding how inflammatory signaling regulates its activity could reduce the abundance of inflammatory mediators such TNF-α and diminish the associated inflammation. Interestingly, the reduced ADAM17 activity was associated with enhanced accumulation of intracellular TNF-α. It has been shown that in contrast to the pathogenic effects of soluble TNF-α, membrane TNF-α functions as an anti-inflammatory factor through reverse signaling in monocytes and macrophages (63), and it also inhibits NF-κB activation and decreases IL-6 and MCP-1 production by forward signaling in adipocytes (64). Notably, membrane TNF in monocytes from RA patients was involved in expansion of Tregs through binding to TNFRII (65). These results highlight that pDCs may orchestrate tolerance induction in RA at various levels ranging from inhibition of TNF-α secretion to induction of Tregs.

Recent advances in high-dimensional single-cell approaches established a functional heterogeneity in the pDC compartment. To this end, mass cytometry and single-cell transcriptomic analysis revealed new cell subsets that express pDC markers (such as CD123, BDCA2, and BDCA4), but they also express markers of classical DCs such as CD11c and CD33 (6668). Our results also showed that BDCA2+BDCA4+ pDCs express a high intensity of CD11c in the peripheral blood of healthy individuals, whereas CD11c expression is reduced in pDCs in the periphery of RA patients in remission. Further experiments are required to shed light in the phenotypic and functional diversity of the pDC compartment in the context of RA but also in the autoimmunity field.

Overall, the findings in the present study place pDCs as an important TNF-α–producing cell in RA, and they unravel an anti-inflammatory role of IL-6 in limiting TNF-α production by this unique cell subset. In addition, we provide mechanistic insights in TNF-α production by pDCs, using CAL-1 cell line as a tool by characterizing a lysosomal-dependent regulation of ADAM17 activity. A thorough understanding of the interplay between inflammatory and regulatory pathways may facilitate the design of effective treatments not only in RA but also in other chronic inflammatory diseases and may shed light into resistance mechanisms of immunotherapy.

We thank the Rheumatology Clinic and Blood Donation Department at the University Hospital of Heraklion for sample collection, Xara Vlata for flow cytometry acquisition (IMBB-Forth), Peter Künzler for excellent technical assistance (RA, fibroblast isolation and culture), and all of the patients and healthy volunteers for participating in the study.

This work was supported by the Hellenic Society for Rheumatology, the Pancretan Health Association, the Celgene Award for Investigative Rheumatology, and European Union Project Innovative Medicine Initiative 6 (BeThe Cure Contract 115142-2 to P.V. and D.B.), and was cofinanced by Greece and the European Union (European Social Fund) through the Operational Programme Human Resources Development, Education and Lifelong Learning in the context of the Reinforcement of Postdoctoral Researchers—2nd Cycle (State Scholarships Foundation Grant MIS-5033021; administered to G.P.) and Strengthening Human Resources Research Potential via Doctorate Research projects (State Scholarships Foundation Grant MIS-5000432; administered to P.G.).

The online version of this article contains supplemental material.

G.P. designed and performed experiments, analyzed data, generated figures, and wrote the manuscript; P.G. performed experiments, analyzed data, generated figures, and wrote the manuscript; J.R.G. and A.G. performed the microarray analysis; G.A.P. and I.I. analyzed the transcriptomic data; B.R. and I.T. provided the CAL-1 cell line; C.O. provided RASFs; G.B. and D.B. participated in the interpretation of data and editing of the manuscript; P.S. performed clinical evaluation of the patients, provided human specimens, supervised the study, and wrote the manuscript; and P.V. designed and supervised the study, performed data analysis, and wrote the manuscript. All authors provided critical reviews of the manuscript.

The microarray data presented in this article have been submitted to ArrayExpress under accession number E-MTAB-11933.

Abbreviations used in this article:

ADAM17

a disintegrin and metalloproteinase 17

CQ

chloroquine

Lamp-1

lysosomal-associated membrane protein 1

3-MA

3-methyladenine

pDC

plasmacytoid dendritic cell

RA

rheumatoid arthritis

RASF

RA synovial fibroblast

TACE

TNF-α–converting enzyme

TFEB

transcription factor EB

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The authors have no financial conflicts of interest.

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