Direct contact with natural killer cells reprograms monocyte-derived dendritic cells into a tolerogenic phenotype
Ammar J Alsheikh, Liang Chen, Daniel Korenfeld, Jason Tam, Suresh Patil, Hsi-Ju Wei, Jianing Li, Suju Zhong, Feng Dong, Terry Melim, Karin Orsi, Samuel Karsen, Yingli Yang, Susan Westmoreland, Aridaman Pandit, Kevin White, Timothy Radstake, Abel Suarez-Fueyo
TL;DR
Contact with natural killer cells changes monocyte-derived dendritic cells into a form that reduces inflammation and promotes immune tolerance.
Contribution
The study identifies a novel approach to reprogram dendritic cells into a tolerogenic phenotype using natural killer cell interactions and confirms key transcription factors involved.
Findings
Dendritic cells exposed to natural killer cells show reduced T-cell activation and increased regulatory mediators like IL-10 and IDO1.
CRISPR knockout experiments reveal that transcription factors IKZF1 and PU.1 are essential for the tolerogenic reprogramming of dendritic cells.
The study confirms that tolerogenic dendritic cells have decreased levels of co-stimulatory markers such as CD11c, CD40, and CD44.
Abstract
Tissue-infiltrating monocyte-derived dendritic cells (DCs) and macrophages are key antigen-presenting cells promoting inflammation and tissue damage by releasing proinflammatory cytokines and strongly activating T cells. These antigen-presenting cells are prominent in inflamed tissues, including in conditions such as inflammatory bowel disease. Modulating the number or function of these cells presents an opportunity to mitigate inflammation and promote immune tolerance in inflammatory disorders. In this study, we comprehensively evaluate a novel approach to modulate these cell populations. First, we employ an antibody-mediated approach to selectively deplete inflammatory DCs while simultaneously reprogramming surviving cells toward a tolerogenic phenotype. Second, we utilize a natural killer–DC co-culture system to dissect molecular mechanisms of DC tolerance induction through…
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Figure 6- —AbbVie10.13039/100006483
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Taxonomy
TopicsImmune Cell Function and Interaction · Immunotherapy and Immune Responses · Immune cells in cancer
Introduction
Myeloid antigen-presenting cells (APCs), including monocytes, dendritic cells (DCs), and macrophages, are key mediators of innate and adaptive immunity, especially within inflamed tissues during autoimmune and inflammatory disorders.1 Beyond their involvement in inflammation, myeloid APCs contribute to maintaining immune homeostasis and promoting tolerance by presenting self-antigens to tissue-specific CD4^+^ and CD8^+^ T cells.2 Recent advances in high-throughput single-cell profiling have highlighted the phenotypic and functional diversity of myeloid APCs, revealing their central contribution to tissue pathology as well as immune regulation in diseases like inflammatory bowel disease.3^,^4 This new knowledge highlights the relevance of selectively targeting pathogenic myeloid subsets for the treatment of various autoimmune diseases, while largely preserving the integrity of innate immune function.
Among the diverse subsets of myeloid APCs, monocytes are unique due to their plasticity, which enables them to differentiate into both macrophages and DCs. Circulating monocytes can rapidly migrate to inflamed tissues, where they differentiate into DCs and macrophages with an enhanced ability to produce proinflammatory cytokines and chemokines, thereby combating infections. Monocyte-derived DCs (MoDCs) and monocyte-derived macrophages (MoMACs), known for their potent capacity to amplify T-cell activation, are classified as inflammatory APCs.4^,^5 In contrast, tissue-resident conventional DCs (cDCs) and macrophages are critical for supporting immune homeostasis and maintaining self-tolerance.6–9 Notably, increased tissue infiltration of MoDCs and MoMACs is a hallmark of several autoimmune diseases, including inflammatory bowel disease (IBD),10–12 hidradenitis suppurativa (HS),13^,^14 rheumatoid arthritis,15^,^16 and atopic dermatitis.17^,^18 MoDCs and MoMACs further perpetuate inflammation by activating tissue-resident memory T cells and recruiting additional immune cells through the secretion chemokines such as CXCL10, CXCL12, CXCL16,19–25 and CCL2/MCP1.26^,^27 In some cases, MoMACs can directly induce tissue damage by producing matrix metalloproteinases (MMPs).28–30 Consequently, therapeutic strategies targeting monocyte migration or selectively depleting tissue-infiltrated monocyte-derived cell populations offer considerable promise for treating autoimmune diseases and inflammatory disorders.
In the present study, we comprehensively investigate the therapeutic potential of targeting inflammatory MoDCs via Dendritic Cell Immunoreceptor (DCIR)-directed monoclonal antibodies (mAbs) in vitro. While targeting DCIR with an mAb successfully depleted a fraction of inflammatory MoDCs in vitro, we observed that the remaining MoDCs exhibited a tolerogenic phenotype, an effect found to be independent of DCIR targeting. By characterizing a natural killer (NK)–DC co-culture system through transcriptomic and epigenomic profiling, as well as functional assays, we discovered that MoDCs undergo an extensive remodeling of their epigenetic landscape, rendering them unresponsive to external stimuli such as lipopolysaccharide (LPS) and suppressing their ability to activate T cells. Furthermore, integrated analysis of the epigenomic and transcriptomic data identified key transcriptional regulators responsible for this tolerogenic shift. CRISPR-based gene knockout experiments subsequently validated the function and relevance of these transcription factors. These data provide compelling support for the ability to modulate DC phenotype and offer avenues for future DC-targeting therapeutic approaches in inflammatory disorders.
Materials and methods
Study approval
All procedures involving human-derived samples were conducted in accordance with protocols approved by AbbVie Inc. Sample sizes were empirically determined to ensure sufficient statistical power, guided by previous studies employing similar analytical approaches.
Human sample collection and processing
Human tissue samples (Fig. 1) included full-thickness surgical resections and normal-adjacent tissue from patients with colon carcinoma. Ulcerative colitis and Crohn disease (CD) samples originated from chronic, treatment-refractory patients, age and sex-matched. Commercial providers (Discovery Life Sciences, Huntsville, AL, USA; BioIVT, Woodbury, NY, USA) shipped all specimens as formalin-fixed, paraffin embedded blocks. For CD mixed lymphocyte reaction (MLR) studies (Fig. 1I, J), cryopreserved lamina propria cells were dissociated from colonic biopsies of 3 treatment-naive, White female CD patients (age 20–40 y, Discovery Life Sciences). Healthy donor leukopaks and whole blood were sourced from Sanguine Biosciences (Waltham, MA, USA).
*Tissue-infiltrated monocyte-derived DCs and macrophages represent the proinflammatory myeloid cells contributing to Crohn disease (CD) progression. Human colon IHC DC-SIGN: (A) control colon, (B) ulcerative colitis (UC) colon, (C) CD colon, (D) GraphPad Prism semi-quantitative IHC scoring (0 to 3). Histologic images captured at 10× magnification. (E) t-distributed stochastic neighbor embedding (TSNE) plot of single-cell RNA-seq analysis for ileum and colon biopsy collected from CD patients (data source: Broad Single Cell Portal, study ID: SCP1884). (F) CD14+ and CD14− cells in myeloid cells subcluster isolated from (E). (G) DGE analysis comparing the CD14+ vs CD14− myeloid cells as indicated in (F). (H) Significantly increased genes of TLRs, chemokines, MHCs, and MMPs in CD14+ myeloid cells identified in (C), compared to the CD14− myeloid cells. (I) Gating strategy for isolation of CD14+ and CD14− myeloid cells from CD colon. (J) Mixed lymphocyte reaction results from co-culturing allogenic T cells with CD14+ and CD14− myeloid cells isolated from CD colon as described in (E). Results from 2 donors with triplicate are shown. Mean ± SEM are shown, and statistical analysis is determined by a one-way ANOVA. *P < .05, *P < .01; ns, not significant.
Sex as a biological variable
For studies involving human tissues and PBMCs, male and female samples were pooled for analysis, and sex was not considered as a biological variable.
Immunohistochemical staining
For immunohistochemical (IHC) staining, tissue sections were stained with H&E and anti-DC-SIGN (R&D Systems, Cat#120507) antibody. DC-SIGN staining was scored in a semi-quantitative manner and summarized.
Flow cytometry and cell sorting
Cells were blocked with 5% human AB serum (Sigma, Cat#4522) or Human TruStain Fc receptor blocker solution (BioLegend, Cat#422302) for 30 min before mixed with antibodies and incubated on ice in the dark for 30 min. Blood or homogenized tissue cells were lysed and fixed before fluorescence-activated cell sorting (FACS) analysis using fix/lyse solution (Invitrogen, Cat#00-5333-54). Samples were acquired on a FACS Canto II, LSRFortessa cytometer (BD Biosciences, Franklin Lakes, NJ, USA), or Cytek Aurora (Cytek Biosciences, Fremont, CA, USA), and analysis was performed using FlowJo software (BD). Fluorescence minus one controls and isotype controls were included where appropriate. Antibodies used for FACS analysis (all from BioLegend unless otherwise indicated) include anti-human CD11b (clone M1/70), anti-human CD11c (BD, clone B-ly6), anti-human CD11c (clone S-HCL-3), anti-human CD14 (clone M5E2), anti-human IL-10R (clone 3F9), anti-human CD19 (clone HIB19), anti-human CD127 (clone A019D5), anti-human CD25 (clone BC96), anti-human CD3 (clone HIT3a), anti-human CD4 (clone OKT4), anti-human CD40 (clone 5C3), anti-human CD44 (clone IM7), anti-human CD45 (clone H130), anti-human CD56 (clone HCD56), anti-human CD69 (clone FN50), anti-human CD8 (clone HIT8a), anti-human HLA-DR (clone L243), anti-human IDO1 (clone 2E2/IDO1), and APC anti-human IgG (H + L) (Jackson ImmunoResearch, Cat#109-136-088). Intracellular staining was performed using the FOXP3 Fix/Perm Buffer Set (BioLegend, Cat#421403) after the surface staining. Snarf-1 dye, which also functions as a sensitive pH indicator, was used to monitor intracellular acidification, an early event in apoptosis, based on the increased 580/640 nm emission ratio.31^,^32
For RNA sequencing (RNA-seq) and assay for transposase-accessible chromatin (ATAC-seq) profiling, weakly NK-engaged DCs were isolated out of NK co-culture with or without LPS using FACS. Cells were collected and labeled with LIVE/DEAD Fixable Aqua Viability Dye (Thermo Fisher, Cat#L34966) and fluorescent surface marker for class II major histocompatibility complex (MHC) (HLA-DR Clone L243, BioLegend, Cat#307618) and NK surface markers (CD56 clone HCD56, BioLegend, Cat#318328; NKG2D clone 1D11, BioLegend, Cat#320826). A total of 100,000 live weak NK-engaged DCs expressing SNARF^+^CellTracker-Aqua^−^ were sorted into separate collection tubes using an AriaFusion cell sorter (BD Biosciences). Sorted cells were immediately used in downstream bulk RNA-seq and bulk ATAC-seq applications.
PBMC, monocyte, DC, and NK cell isolation and culture
PBMCs were isolated from leukopaks and whole blood using Ficoll density gradient centrifugation (Cytivia, Cat#17144002). Monocytes (CD14^+^) and NK cells were purified using microbeads (Miltenyi Biotec, Cat#130-050-201 and #130-050-401). Cells were cultured in complete RPMI1640 medium (Gibco, Cat#11875093) containing 10% FBS (Cytiva, Cat#SH30071.03), GlutaMAX (Gibco, Cat# 35050079), nonessential amino acids (Gibco, Cat#11140050), pyruvate (Gibco, Cat#11360070), penicillin + streptomycin (Gibco, Cat#15140122), 50 µM 2-mercaptoethanol (Gibco, Cat#21985023), and 25 mM HEPES (Gibco, Cat#15630080).
Anti-DCIR antibody generation
We utilized the anti-DCIR antibody generated previously.33 In brief, the mAb was generated using immunization of Sprague-Dawley JR14 rats with human DCIR (huDCIR)–containing vector followed by isolation of splenic B cells for hybridoma preparation. Hybridoma producing strong binders to huDCIR expressing HEK293 were selected and cloned into a human IgG1 backbone, expressed then fully humanized and produced from the Expi293 cells.
MoDC differentiation and stimulation and DC–NK cell co-culture
CD14^+^ monocytes were purified from PBMCs using microbeads (Miltenyi Biotec, Cat# 130-090-879). Human MoDCs were differentiated from monocytes supplemented with 40 ng/mL human IL-4 (Peprotech, Cat# 200-04) and 100 ng/mL human GM-CSF (Peprotech, Cat#300-03) for 6 to 7 days. For inhibiting the IDO1 activity, indoximod (MCE, Cat#HY-16724) was added together with GM-CSF and IL-4 during MoDC differentiation and MoDC maturation induced by 1 μg/mL CpG ODN2006 (Invivogen, Cat#tlrl-2006) and 500 ng/mL CD40L (PeproTech, Cat#310-02). Untouched human NK cells were isolated from fresh whole blood using microbeads (Miltenyi Biotec, Cat#130-092-657) and primed with 10 ng/mL human IL-15 (Peprotech, Cat#200-15) overnight. Purified MoDCs were labeled with Snarf-1 (Invitrogen, Cat#C1270), and NK cells were labeled with CellTracker Green (Invitrogen, Cat#C2925) per manufacturer’s protocol. Labeled NK cells and MoDCs were mixed at different NK-to-DC ratio (1:5, 1:1, 5:1, 10:1, 20:1) for overnight. Apoptotic MoDCs were determined by the increased 580/640 nm emission ratio of Snarf-1 after NK and DC co-culture. For LPS restimulation, weakly NK-engaged DCs were sorted from the live NKp46-CD11c^+^HLA-DR^+^Snarf-1^+^CellTracker Green^−^ cells from the NK cells and DCs co-culture and stimulated with 100 ng/mL ultrapure LPS (Invivogen, Cat#tlrl-3pelps) for 4 hours, and then cells were evaluated by flow cytometry or omic profiling for the different experiments.
Mixed lymphocyte reaction
For the MLR experiment in Fig. 1I and J, myeloid cell populations (EpCAM^−^CD45^+^CD11b^+^) were sorted into live CD14^+^ and CD14^−^ subsets using a BD FACSAria cell sorter. Allogenic pan-T cells were isolated from healthy donors with the Pan T Cell Isolation Kit (Miltenyi Biotec, Cat#130-096-535), following the manufacturer’s instructions, and stained with CellTracker Green (Invitrogen, Cat#C2925). Sorted CD14^+^ and CD14^−^ myeloid cells were co-cultured with pan-T cells at a 1:4 myeloid-to-T-cell ratio for 3 days to induce MLR, after which the number of viable T cells was quantified by flow cytometry.
For the MLR experiment in Fig. 3, PBMCs were isolated from fresh human whole blood and treated with 40 ng/mL human IL-4 and 100 ng/mL human GM-CSF for 3 days to induce MoDC differentiation, followed by CpG ODN2006 (Invivogen, Cat# tlrl-2006) and CD40L (PeproTech, Cat#310-02) to promote MoDC maturation in addition to anti-DCIR antibody to induce antibody-dependent cellular cytotoxicity (ADCC) for 2 to 3 days. For neutralization of IL-10, 20 μg/mL anti-human IL-10 antibody, clone JES3-19F1 (BioLegend, Cat#506802) was added along with anti-DCIR antibody during the ADCC assay. Pan T-cells were isolated by a human Pan T Cell Isolation Kit (Miltenyi Biotec Cat#130-096-535) and labeled with CellTrace CFSE Cell Proliferation Kit (Invitrogen, Cat#C34570) according to manufacturer’s instructions. PBMCs were washed and resuspend with allogenic pan-T cells to induce MLR at 1:2 ratio (PBMCs to allogenic T cells) by co-culture for 5 days. For MRL experiments in Fig. 6, CRISPR-modified MoDCs were co-cultured with allogenic pan-T cells to induce MLR at 1:3 ratio (MoDC to allogenic T cells) by co-culture for 5 days. For evaluating T-cell activation, cells from the MLR assay were collected and stained with fluorescent antibody panel. Cells were first washed with PBS and resuspended in LIVE/DEAD Fixable Blue Viability Dye (Invitrogen, Cat#L23105) and Human TruStain FcX block (BioLegend, Cat#422302) for 15 min at room temperature. Then cells were spun down and washed with staining buffer (BioLegend, Cat#420201) and stained with antibody mix including anti-CD3 BV570 (BioLegend, Cat#300436), anti-CD4 cFluor YG384 (Cytek Biosciences, Cat#R7-20041), anti-CD8 APC-Fire 810 (BioLegend, Cat#344764), and anti-CD25 Alexa Fluor 647 (Thermo Fisher Scientific, Cat#51-0259-42) for 20 min at 4 °C. Cells were then washed and analyzed on a Cytek Aurora full-spectrum cytometer (Cytek Biosciences).
RNA isolation and RNA-seq library preparation
Total RNA from MoDCs was isolated using a spin-column extraction kit according to the manufacturer’s instructions (Macherey-Nagel, Cat#740990). The isolated high-quality RNA was then further amplified and indexed using a Unique Molecular Identifier-based library amplification and preparation kit following the manufacturer’s instructions (Takara Bio, Cat#634354). RNA libraries were pooled and sequenced using an Illumina NextSeq2000 sequencer to obtain 150 paired-end base reads (Illumina). Pooled library was sequenced to achieve a coverage of 30 million reads per sample.
ATAC-seq and library preparation
Fifty thousand sorted MoDCs were prepared for ATAC using ATAC-seq kit (Active Motif, Cat#53150) according to the manufacturer’s instructions. In brief, cells were spun down and for 5 min at 4 °C 500 × g and resuspended and washed once with ice-cold PBS. Cells were then resuspended in 100 μL of lysis buffer and spun down for 10 min. Lysis buffer was removed, and nuclei were resuspended in tagmentation mix and incubated in a 37 °C shaker at 800 rpm for 50 min. This was followed by column DNA purification and PCR amplification using unique Nextera-compatible i7 and i5 indexed primers (Active Motif, Cat#53155). After lower-sided SPRI cleanup, libraries were quality controlled using automated electrophoresis Tapestation system (Agilent) on a D1000 ScreenTape (Agilent, Cat#5067-5582, Cat#5067-5583). Library concentration was quantified using Qubit dsDNA High Sensitivity kit (Thermo Fisher, Cat#Q32851). Libraries were diluted and pooled and multiplexed on a single NextSeq 2000 sequencer (Illumina) using P3 Kit 100-cycle run (Illumina, Cat#20040559).
RNA-seq and ATAC-seq data processing and analysis
RNA-seq libraries were demultiplexed using Bcl2fastq (Illumina). Reads were processed and aligned to the human genome (UCSC: hg38) by using CogentAP NGS V2.0 analysis pipeline (Takara). After filtering lowly expressed genes (count <10), differential expression analysis was performed using DEseq (DESeq2 R package) with adjusted P value cutoff of <0.05.
ATAC-seq data were demultiplexed using Bcl2fastq (Illumina), and reads are aligned to the human genome (UCSC: hg38) using R Subread and indexed and sorted using R Sam tools. After filtering non-autosomal reads, peak calling was run using Macs (format: BAMPE, q-value: 1e-2). Downstream and differential peak analysis was run using ChipQC (DESeq) using consensus peak counts in each group. Transcription factor motif enrichment was performed using Analysis of Motif Enrichment (part of MEME3 suite of tools). Bigwig files were created using R tracklayer for visualization in IGV browser.
Bioinformatic analysis of tissue-infiltrating monocytes, DCs, and macrophages
Single cell RNA-seq data of HS were extracted from the National Center for Biotechnology Information Gene Expression Omnibus database (study ID: GSE220116)34 and raw data of CD were downloaded from Broad Single Cell Portal (study ID: SCP1884).35 BioTuring (https://bioturing.com/) was used to perform the differential gene expression (DGE) analysis36 with Venice method37 comparing CD14^+^ and CD14^−^ myeloid cells in the disease lesional tissues.
CRISPR-Cas9 knockout of genes in MoDCs
CD14^+^ monocytes were cultured in media supplemented with 40 ng/mL human IL-4 (Peprotech, Cat#200-04) and 100 ng/mL human GM-CSF (Peprotech, Cat#300-03). At day 3, 10^6^ cells per sample were harvested, washed with PBS, and resuspended in the buffer provided in the P3 Primary Cell 4D-Nucleofector X Kit (Cat#V4XP-3032, Lonza). For RNP complex preparation, 61pM Sp HiFi Cas9 Nuclease V3 (IDT) was preincubated with 100 pM sgRNA mix (Genscript) of the indicated gene or nontargeted control (sgRNA sequences in Table S1) for 10 min at room temperature. This RNP complex was added to the cell suspension prior to electroporation using program CA-137 in the 4D-Nucleofector System (Lonza). Cells were recovered in media supplemented with 40 ng/mL human IL-4 (Peprotech, Cat#200-04) and 100 ng/mL human GM-CSF (Peprotech, Cat#300-03), cultured for 3 days. To determine the indel rate, the targeted loci were amplified by PCR (primer sequences in Table S1) from DNA extracted from the edited MoDCs followed by Sanger sequencing (Genscript) and analysis using the Inference of CRISPR Edits (ICE) tool (Synthego).
Following MoDC differentiation, cells were stimulated with 100 ng/mL LPS (Invitrogen, Cat#tlrl-3pelps) for 4 hours and secreted cytokines were measured from supernatants from all knockout conditions using a 48 Plex Milliplex MAP Human Cytokine/Chemokine/Growth Factor Panel (Millipore Sigma, Cat#HCYTA-60K-PXBK48) based on the manufacturer’s instructions and was run on a Luminex FLEXMAP 3D instrument (Invitrogen, Cat#APX1342). Results reported are calculated cytokine concentrations based on standard curves. MLR T-cell activation was assessed by flow cytometry as indicated above.
Statistical analysis
Statistical analysis was performed using GraphPad Prism software. Significance between 2 groups was determined by unpaired, 2-tailed Student t test, and significance between multiple groups was determined using one-way analysis of variance (ANOVA) with Dunnett posttest. For all statistical comparisons, *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001. All composited and representative data are generated from at least 2 independent experiments.
Results
Tissue-infiltrated monocyte-derived DCs and macrophages are prominent in IBD
In contrast to DCs and macrophages that differentiate from infiltrating monocytes in response to immune stimuli, tissue-resident myeloid cells play a critical role in maintaining immune homeostasis under steady-state conditions. For instance, colon-resident macrophages and DCs exhibit resistance to immune activation and produce anti-inflammatory cytokines, such as IL-10, to induce tolerance in both innate and adaptive immune responses.8^,^38^,^39 However, during inflammation, monocytes are recruited to affected tissues and differentiate into DC-SIGN/CD209^+^ macrophages and DCs,40^,^41 adopting a more inflammatory phenotype.4 In inflamed colonic mucosa from patients with IBD, we observed a significant infiltration of DC-SIGN^+^ monocyte-derived macrophages and DCs (Fig. 1A–D).
Macrophages and DCs derived from tissue-infiltrated monocytes can serve as significant sources of proinflammatory cytokines, such as TNF-α, IL-6, and IL-23,42 and chemotactic agents such as CCL2/MCP1, CXCL12/SDF1, CXCL13, and CCL22/MDC,43^,^44 which prime and activate both innate and adaptive immune responses. In addition to cytokine production, these cells contribute directly to tissue destruction through the secretion of MMPs.45 Single-cell RNA-seq of lesional tissues from IBD revealed elevated expression of DC-SIGN/CD209 and TREM2 in CD14^+^ monocytes, indicating their differentiation into proinflammatory DCs and macrophages. These cells are characterized by increased mRNAs of co-stimulatory markers (eg CD11c/ITGAX, CD40, CD44, CD83, and CD86) and proinflammatory cytokines (eg IL-1B, IL-18, and TNF) (Fig. 1E–G).35 The pathogenic role of these cells is further supported by elevated mRNA levels of Toll-like receptors (TLRs), chemokines, MHCs, and MMPs, compared to CD14^−^ myeloid cells (Fig. 1H). Additionally, we found that CD14^+^ myeloid cells isolated from the colons of Crohn’s disease patients induced stronger T-cell proliferation in an MLR assay (Fig. 1I, J). These findings suggest that colonic CD14^+^ myeloid cells are key APCs that sense commensal bacteria and drive excessive T-cell activation in the gut. Consequently, selectively depleting DCs and macrophages derived from CD14^+^ monocytes presents a potential strategy to eliminate pathogenic myeloid cells and restore immune tolerance in disease lesions, while preserving noninflammatory, tissue-resident macrophages and DCs for the treatment of IBD.
In addition to IBD, we observed that CD14^+^ myeloid cells also represent proinflammatory APC subsets in the lesional skin of HS,34 exhibiting elevated mRNA levels of various proinflammatory cytokines such as IL-6 and TNF, along with TLRs, chemokines, and MMPs (Fig. S1A–F). Notably, these cells showed significant upregulation of key inflammasome components, including NLRP3, PYCARD/ASC, CASP1, CASP4, IL1A, IL-1B, and IL-18, indicating increased inflammasome/IL-1 pathway activation (Fig. S1G). Although abnormal inflammasome/IL-1 signaling activation has been documented in HS patients, it is not effectively addressed by anti-TNF-α therapy.46 These observations suggest that targeting tissue-infiltrated CD14^+^ myeloid cells could provide additional therapeutic benefits that complement anti-TNF-α treatment. These data together highlight the relevance of manipulation of these inflammatory cells for multiple inflammatory indications.
Our analysis in samples from IBD and HS showed CLEC4 (DCIR) as a marker expressed in dendritic cells derived from monocytes (Fig. 1G and Fig. S1C), confirming previous reports.47 We have previously developed an anti-DCIR antibody, clone 5E11, enabling us to investigate the mechanisms of modulation for future therapeutic potential.33
Depleting and modulating the phenotype of MoDCs through an anti-DCIR mAb
We initially validated that the anti-DCIR antibody selectively binds to human monocytes, MoDCs, and MoMACs (Fig. 2A, B). To study the effect of this antibody on MoDCs, we utilized primary human PBMCs cultured in human GM-CSF/IL-4 to differentiate MoDCs followed by CD40L/CpG maturation in the presence of anti-DCIR antibody or isotype control. We observed depletion of 70% of CD11c^+^CD14^+^ myeloid APCs in vitro, demonstrating the potential of this tool antibody to eliminate these cells (Fig. 2C, D).
*Surviving monocyte-derived DCs following ADCC exhibit tolerogenic phenotypes. (A) Flow cytometry analysis of house-made anti-DCIR (clone 5E11) antibody binding to the CD14+ monocytes, B cells, T cells, and NK cells from human PBMCs (n = 4 donors). Means ± SEM are shown, and statistical analysis is determined by one-way ANOVA test with Dunnett correction compared to the isotype staining control. *P < .01. (B) Flow cytometry analysis of our anti-DCIR (5E11) antibody’s binding to the CD14+ monocytes, monocyte-derived DCs and macrophages (n = 4 donors). (C) Study design of ADCC assay using anti-DCIR antibody to deplete monocyte-derived DCs differentiated and matured in vitro. (D) Cell count, (E–G) Mean Fluorescence Intensity (MFI) of surface co-stimulatory markers (CD11c, CD40, and CD44), and (H) intracellular IDO1 expression of surviving CD14+CD11c+ monocyte-derived DCs. (I) IL-10 cytokine production, and (J) %Treg (CD25+CD127−CD4+ T cells) induction in the ADCC assay as described in (C). Representative data from 1 of 3 donors are shown with error bars indicating mean ± SEM.
While our antibody can effectively induce the depletion of MoDCs and MoMACs, both in vivo and in vitro, a significant portion (∼30%) of targeted cells survived ADCC in our in vitro experiments. We decided to study these cells and discover that these surviving APCs displayed markedly reduced expression of DC maturation and co-stimulatory markers, such as CD11c, CD40, and CD44 (Fig. 2E–G), along with increased expression of tolerogenic factors, including IL-10 and IDO1 (Fig. 2H, I). Additionally, one of the key functions of tolerogenic DCs is their ability to stimulate regulatory T-cell (Treg) differentiation. We also observed a shift toward enhanced differentiation of Tregs (Fig. 2J).
Furthermore, when anti-DCIR antibody was added to PBMCs in an MLR assay (Fig. 3A, B), we observed suppression of allogeneic T-cell activation, as evidenced by reduced proliferation and CD69 expression in both CD4^+^ and CD8^+^ T cells (Fig. 3C–F). This was accompanied by a decrease in Th1- and Th2-associated cytokine production, including IL-2, IFN-γ, IL-13, and CCL22/MDC (Fig. 3G–J). Additionally, production of CCL2/MCP1, a chemokine abundantly secreted by both activated myeloid cells and T cells during antigen presentation, was also inhibited (Fig. 3K). These findings provide proof of concept that targeting myeloid APCs with anti-DCIR antibodies can induce bidirectional tolerance in both innate and adaptive immune responses.
Depletion of DCs and induction of tolerogenic DC by ADCC suppress T-cell activation. (A) Study design of mixed lymphocyte reaction (MLR) assay after using anti-DCIR antibody to deplete monocyte-derived DCs differentiated and matured in vitro. (B) Gating strategy of flow cytometry analysis on the activation of allogenic T cells. (C–F) T-cell proliferation and activation indicated by the increased T-cell count and CD69 expression on CD4+ (C, D) and CD8+ T cells (E, F). (G–K) Cytokine production from the MLR assay as described in (A). Representative data from 1 of 2 donors are shown, with error bars indicating mean ± SEM.
NK cell–induced DC tolerance is independent of anti-DCIR antibody-mediated receptor signaling
Previous studies have shown that DCIR amplifies immune-tolerizing signals and prevents myeloid cell activation.48^,^49 We previously reported that our anti-DCIR antibody can induce immune-inhibitory through agonistic action on the DCIR.33 To distinguish the effects of DCIR-mediated suppressive signaling from NK cell–induced DC reprogramming, we employed a co-culture system to enforce the lysis of mature MoDCs by IL-15–activated NK cells. This was achieved by increasing the NK-to-DC ratio (Fig. 4A) without relying on ADCC effects.50 In this system, Snarf-1–labeled MoDCs were co-cultured with an increased number of CellTracker Green–labeled NK cells primed with human IL-15. CellTracker Green, cleaved by intracellular esterases, is retained in viable cells with intact membranes but can be transferred to target cells during NK-mediated killing through cytotoxic content release and trogocytosis-mediated membrane exchange.50 Following co-culture, we identified 2 distinct populations of DCs: double-positive cells for both CellTracker Green and Snarf-1, indicating strong NK engagement (referred to as strong engagers), and single-positive cells for Snarf-1 alone, indicating weak NK engagement (referred to as weak engagers) (Fig. 4B). Snarf-1 dye, which also functions as a sensitive pH indicator, was used to monitor intracellular acidification, an early event in apoptosis, based on the increased 580/640 nm emission ratio.31^,^32 Weakly NK-engaged DCs showed no significant change in intracellular pH, indicating an absence of apoptotic activity. In contrast, strongly NK-engaged DCs displayed increased intracellular acidification, indicating an apoptotic phenotype, which correlated with higher NK-to-DC ratios (Fig. 4C). In line with previous studies,50 we observed minimal DC apoptosis at low NK-to-DC ratios (1:1 and 1:5), but saturated and incomplete killing at higher ratios (10:1 and 20:1) (Fig. 4D). This parallels the incomplete killing observed in the aforementioned anti-DCIR antibody-mediated ADCC, further suggesting that the survival of MoDCs from the NK-mediated cytolysis occurs independently of the DCIR receptor’s inhibitory signaling.
*DC tolerance induced by NK cells is not dependent on the receptor signaling targeted by the antibodies during ADCC. (A) Study design of monocyte-derived DC (MoDCs)–NK cell co-culture. (B) Gating strategy for identifying the strong and weakly NK-engaged DC population in (A). (C, D) Flow cytometry analysis of the MoDCs’ apoptosis (indicated by increased 580/640 nm emission ratio of Snarf-1) from the isolated strong and weakly NK-engaged MoDCs with increased proportion of NK cells (NK:DC ratio: 1:5, 1:1, 5:1, 10:1, 20:1) in co-culture (n = 4 biological replicates). (E) Study design for investigating DC anergy of the surviving weakly NK-engaged MoDCs in response to LPS stimulation, compared with the MoDCs cultured alone. (F) MFI of CD40 expression on MoDCs as treated in (E) (n = 3 biological replicates). (G, H) MFI of CD69 expression on CD4+ and CD8+ T cells, and (I) IFN-γ cytokine production from the MLR assay by co-culturing allogenic T cells with normal MoDCs and sorted surviving weakly NK-engaged MoDCs (n = 3 biological replicates). Means ± SEM are shown, and statistical analysis is determined by a one-way ANOVA, *P < .05, **P < 0.01, **P < 0.001; ns, not significant.
To determine whether the reprogramming of DCs into a tolerogenic phenotype is directly induced by engagement with NK cells, we isolated weakly NK-engaged DCs from the co-culture and restimulated them with LPS (Fig. 4E). We found that these NK-engaged DCs exhibited attenuated maturation, as indicated by reduced expression of co-stimulatory and maturation markers such as CD40, compared to DCs cultured alone (Fig. 4F). Moreover, when co-cultured with allogeneic T cells, the weakly NK-engaged DCs induced significantly lower T-cell activation, as shown by decreased CD69 expression (Fig. 4G, H) and reduced IFN-γ production (Fig. 4I). This demonstrates that the diminished T-cell activation observed in our previous MLR experiment with anti-DCIR treatment (Fig. 3) is not solely due to the reduction of MoDCs through ADCC but is also a result of NK-induced tolerogenic DC reprogramming.
The maturation and antigen-presenting function of DCs might be affected by sensing the secretory factors from the activated NK cells. To test whether direct NK-DC contact is required for this reprogramming, we cultured MoDCs with IL-15–activated NK cells separated by a Transwell membrane (Fig. S2A). We observed that soluble factors from NK cells were insufficient to induce MoDC apoptosis and failed to attenuate MoDC maturation following LPS stimulation (Fig. S2B–D). This confirms that direct contact between NK cells and DCs, and the subsequent apoptosis of MoDCs, is essential for inducing tolerogenic DCs.
In addition to the myeloid-specific anti-DCIR antibodies used in our study, other antibodies have been shown to induce DC tolerance in different models. For example, anti-CD83 antibodies in a murine collagen-induced arthritis model51 and daratumumab (an anti-CD38 antibody) in a phase 2 clinical trial for systemic lupus erythematosus52 both demonstrated tolerance-inducing effects by targeting activated DCs, and led to an increased IL-10 production. Based on our results, we conclude that NK cell–induced tolerance in DCs is independent of receptor signaling triggered by specific antibodies.
Epigenetic remodeling of chromatin accessibility drives the transcriptional regulation of tolerogenic phenotypes in NK-programmed DCs
Tolerogenic DCs are characterized by the downregulation of co-stimulatory markers (eg CD11c, CD40, CD44, CD86, and MHC-II)53 and the upregulation of tolerizing factors such as IL-10 and PD-L1,54 across various procedures identified for inducing tolerogenic DCs, including treatment with IL-10, vitamin D3, and dexamethasone. It has been reported that both NK and T cells can modulate DC function posttranslationally by removing co-stimulatory markers like CD86 and MHC-II through trogocytosis.55 However, maintaining tolerogenic phenotypes via posttranslational mechanisms can be challenging in the highly inflamed microenvironments of disease lesions. It is important to explore whether transcriptional regulation, through epigenetic changes, also contributes to the induction and maintenance of DC tolerance.
Therefore, we analyzed the transcriptomic and epigenetic profiles of weakly NK-engaged DCs isolated from NK-DC co-cultures compared to DCs cultured alone, both before and after LPS stimulation (Fig. 5A). Bulk RNA-seq revealed significant alterations in gene expression patterns in weakly NK-engaged DCs compared to DCs cultured alone, and these changes persisted even after LPS stimulation, unlike in the control DCs alone condition that responded robustly to LPS (Fig. 5B). Notably, while LPS-stimulated DCs alone showed elevated expression of proinflammatory cytokines and chemokines such as IL-1B, IL-6, and CXCL10, weakly NK-engaged DCs displayed anergy to LPS treatment in general (Fig. 5C). DGE analysis further highlighted a significant downregulation of genes involved in LPS signaling pathways, including TLR4, CD14, MYD88, and TRAF6 in the isolated NK-engaged DCs, indicating a state of LPS anergy (Fig. 5D). Moreover, we observed an increase in the expression of tolerogenic factors (eg IL-10, IDO1, and AIRE) and a reduction in co-stimulatory markers (eg CD40, CD44) in weakly NK-engaged DCs (Fig. 5D), aligning with findings from previous anti-DCIR treatment (Fig. 2). This transcriptional shift was supported by ATAC-seq analysis, which showed extensive chromatin remodeling in weakly NK-engaged DCs (Fig. 5E). Promoter regions of proinflammatory cytokines (eg TNF-α and IL-6) and co-stimulatory markers (eg CD40, CD11c/ITGAX) exhibited reduced accessibility (Fig. 5F). Additionally, based on the differential chromatin accessibility indicated by ATAC-seq peaks, transcription factor motif enrichment analysis predicted that activities of multiple transcription factors (eg PU.1/Spi1, IRF1, and IKZF1) were downregulated in the weakly NK-engaged DCs, potentially contributing to their tolerogenic phenotype (Fig. 5G).
Epigenetic remodeling of chromatin accessibility induces the transcriptional regulation of tolerogenic phenotypes in NK-programmed DCs. (A) Study design of bulk RNA-seq and ATAC-seq analysis on the surviving MoDCs from the NK–DC co-culture (weak NK engager) and MoDC cultured alone collected before and after LPS stimulation. (B) Principal component analysis (PCA) plot of the RNA-seq results using edgeR showing the 4 groups as defined in (A). (C) Gene expression changes identified by DGE analysis comparing the effects of LPS stimulation in DCs cultured alone or co-cultured with NK cells (weak NK engager). (D) Gene expression changes identified by DGE analysis comparing DC (weak NK-engager) vs DC alone. (E) Correlation of changes in chromatin accessibility determined by ATAC-seq and changes in gene expression determined by bulk RNA-seq in DCs alone compared to DCs co-cultured with NK cells (weak NK-engager). (F) Genome browser plots showing chromatin accessibility indicated by ATAC-seq peaks in proinflammatory cytokines genes (eg IL-1B, IL-6) and DC co-stimulatory marker genes (eg CD40 and CD11c) loci. Highlighted peaks indicate promoters that have lower accessibility in DC (weak NK-engager) as compared to DCs alone (DEseq, FDR <0.05). (G) Predicted top 20 most enriched transcription factors motifs based on the differentially chromatin accessible peaks identified by the ATAC-seq results.
In summary, our results suggest that NK-mediated reprogramming of DCs is achieved through enhanced transcription of anti-inflammatory factors and selective closure of proinflammatory gene loci, driven and maintained by epigenetic modifications, even after perturbation of strong immune stimuli (for instance, LPS used in our study). The epigenetically mediated reprogramming suggests that NK-induced DC tolerance may be a stable and long-lasting effect, even within the inflammatory conditions present in disease tissues.
PU.1-IKZF1-IL-10 signaling is critical for NK-programmed DC tolerogenesis
To further assess which of the identified transcription factors contribute directly to the tolerogenic phenotype observed in DCs after co-culture with NK cells, we utilized CRISPR-Cas9 technology to knock out these transcription factors in human MoDCs (Fig. 6A). MoDCs with knockout of PU.1 exhibited a decrease trend in proinflammatory cytokine production, such as TNF-α and IL-6, while PU.1 and IKZF1-knockout led to an increase in IL-10 secretion in response to LPS stimulation (Fig. 6B). Consistently, when these PU.1 or IKZF1-knockout MoDCs were mixed with allogeneic T cells in MLR, there was a significant reduction in T-cell activation, as evidenced by decreased expression of IL2Rα/CD25 expression on the T cell (Fig. 6C). Finally, we observed a reduction of proinflammatory cytokines TNF-α and IFN-γ and an increase in IL-10 in the MLR media (Fig. 6D). These results indicate that impaired activity of PU.1 and IKZF1 plays a direct role in the induction of DC tolerogenesis including modulating T-cell activation and cytokine production and enhances IL-10 production.
*IL-10 signaling is critical for the tolerogenic effect from the NK-programmed DC. (A) Study design of knocking-out transcription factors and evaluating the resultant DC tolerance effects. (B) IL-10, TNF-α, and IL-6 cytokine production was assayed from LPS-stimulated MoDCs after CRISPR-Cas9 editing. CRISPR-KO MoDC effect on T-cell activation in MLR evaluated by flow cytometry showing (C) CD25 expression on CD4 T cells and CD8 T cells (n = 3 donors) and (D) evaluated MLR secreted cytokines IL-10, TNF-α, and IFN-γ (n = 3 donors). Means ± SEM are shown, and statistical analysis is determined by one-way ANOVA Fisher least significant difference test compared to the scramble sgRNA control. *P < .05, **P < .01, ***P < .001, ***P < .0001. (E) Study design of neutralizing IL-10 during the depletion of MoDCs by anti-DCIR–induced ADCC effect. (F) Cell count and (G, H) MFI of DC co-stimulatory/maturation markers (CD40 and CD44) on surviving CD14+CD11c+ MoDCs in the ADCC assay induced by anti-DCIR antibody in combination with or without anti-IL-10 neutralizing antibody (n = 3 donors). (I) IL-10R expression and (J) cell viability of the MoDCs differentiated by GM-CSF and IL-4 treatment and matured by CpG and CD40L stimulation, along with indoximod, an IDO1 inhibitor (n = 4 donors). (K) Schematic summary of the tolerogenesis of DCs from the NK engagement induced by DC-targeting antibody (eg anti-DCIR).
Furthermore, the role of IDO1 and IL-10 in the immunosuppressive function of tolerogenic DCs, induced by tolerizing agents like dexamethasone,56 prompted us to explore their function in our system. Interestingly, neutralizing IL-10 with an anti-human IL-10 antibody (clone JES3-19F1) during the ADCC assay (Fig. 6G) not only reduced DC depletion but also increased DC maturation marker expression, compared to anti-DCIR treatment alone (Fig. 6H–J). Contrary to suppressing NK cells, IL-10 has been shown to enhance NK cell cytotoxicity,57 suggesting that IL-10 signaling is instrumental not only in inducing DC tolerance through NK cell interactions but also in promoting the clearance of inflammatory DCs. Meanwhile, IDO1, critical in tryptophan metabolism, can activate the aryl hydrocarbon receptor (AhR), promoting IL-10 receptor (IL-10R) expression.58 To this end, we treated monocytes with indoximod, a validated IDO1 inhibitor,59 alongside GM-CSF and IL-4 treatment during MoDC differentiation and maturation with CpG and CD40L stimulation. Inhibition of IDO1 led to reduced IL-10R/CD210 expression on the MoDCs but did not impact cell viability (Fig. 6I, J). In summary, our data demonstrate that direct NK-DC engagement could downregulate PU.1 and IKZF1 activities, enhancing IL-10 expression in DCs. Also, the resultant increase in IDO1 expression elevates IL-10R on DCs, thereby augmenting the impact of IL-10 signaling and converting the surviving DCs into a tolerogenic phenotype (Fig. 6K).
Discussion
Myeloid APCs can exhibit both proinflammatory and tolerogenic roles, influencing outcomes in autoimmune and inflammatory disease. Evidence from CD11c-DTR transgenic mice suggests that depletion of myeloid APCs can protect against excessive inflammation in various autoimmune disease models.60–64 Our study identifies tissue-infiltrated, monocyte-derived DCs and macrophages as central drivers of inflammation and immune dysregulation in both IBD and HS. By delineating the distinct roles of tissue-resident versus monocyte-derived myeloid cells, we show that inflammatory lesions in IBD and HS are characterized by prominent infiltration of CD14^+^ monocyte-derived DCs and macrophages. These populations display heightened proinflammatory, antigen-presenting, and tissue-destructive functionalities, evidenced by upregulated surface costimulatory markers, increased production of inflammatory cytokines (eg TNF-α, IL-6, IL-23), chemokines, and MMPs, as well as the capacity to drive excessive T-cell activation. Our findings underscore that tissue-infiltrated CD14^+^ myeloid cells function as key APCs that sense commensal bacteria and serve as initiators and amplifiers of the local inflammatory response.
Importantly, we show that selective targeting of these pathogenic myeloid subsets—specifically through an anti-DCIR antibody—results in effective depletion of inflammatory monocyte-derived DCs and macrophages. Notably, about 30% of these cells survive ADCC; these survivors are not mere bystanders but exhibit reduced maturation and co-stimulatory marker expression, upregulated tolerogenic factors such as IL-10 and IDO1, and an increased propensity to induce Tregs. Functionally, the presence of the anti-DCIR antibody in MLRs led to suppressed T-cell activation, accompanied by broad inhibition of both Th1/Th2 cytokine secretion and chemokine production.
To distinguish DCIR-dependent suppression from NK cell–induced DC modulation, we employed an NK–DC co-culture system. Our mechanistic data demonstrate that this tolerogenic reprogramming of DCs can be induced via direct engagement with NK cells, independent of antibody-mediated DCIR signaling. Using the NK–DC co-culture system, we reveal that DCs surviving strong NK engagement display anergy to proinflammatory stimuli such as LPS, with persistent transcriptional downregulation of costimulatory and maturation genes, and stable upregulation of tolerogenic mediators including IL-10 and IDO1. ATAC-seq analysis reveals extensive chromatin remodeling in these tolerized DCs, with decreased promoter accessibility at inflammatory gene loci and downregulation of transcription factors PU.1 and IKZF1. Functional experiments using CRISPR-mediated knockouts of PU.1 and IKZF1 further confirm their role in promoting tolerogenic gene expression and suppressing DC-driven T-cell activation. Interactions between NK cells and DCs are complex and context-dependent resulting in either mutual activation65^,^66 or inhibition.55 Previous work has shown that DC-derived cytokines like IL-12, IL-15, and IL-18 can activate NK cells and induce IFN-γ, which in turn can enhance DC maturation and inflammatory cytokine secretion.66–69 Conversely, regulatory roles of NK cells have also been observed, such as in acute infection models with Listeria monocytogenes and Yersinia pestis, where IFN-γ–producing NK cells suppress DC IL-12 production via IL-10, thereby curbing further NK cell activation and tissue damage.70 Moreover, IFN-γ from NK cells can induce expression of checkpoint ligands such as PD-L1 and HLA-E, which engage inhibitory receptors, such as PD1 and CD94/NKG2A, and mitigate excessive NK cell activity.71 The outcome of NK–DC interactions may also be influenced by the ratio of NK to DCs, with lower NK-to-DC ratios promoting DC maturation, while higher ratios can induce DC anergy and tolerance.50 Thus, enhancing local NK–DC interactions in the disease lesion through antibody mediation might offer a potent tolerogenic effect. For instance, in models of graft-versus-host disease and collagen-induced arthritis, anti-CD83 antibodies not only depleted mature DCs but also reprogrammed surviving DCs to express higher levels of IDO1, PD-L2, and IL-10, facilitating Treg induction and reducing T-cell activation.51^,^72 Similarly, in a phase 2 trial for daratumumab (anti-CD38) in systemic lupus erythematosus, depletion of plasma B cells and DCs led to a reduction in autoantibodies and type I interferon production, alongside a significant increase in IL-10 production.52 Although activated B cells can also be targeted by CD83 and CD38 antibodies, using the anti-DCIR antibody and DC–NK coculture system described in this study, we further clarify that the IL-10–producing DCs can be directly induced by the NK engagement. These results together suggest that NK-driven reprogramming acts via stable, epigenetic silencing of inflammatory pathways and transcriptional activation of tolerance-related programs.
Although the precise mechanisms remain to be fully elucidated, our data and the literature suggest that increased cell-to-cell contact at higher NK:DC ratios promote stronger downregulation of costimulatory markers (eg CD86, HLA-DR) and upregulation of tolerogenic mediators such as IL-10 and IDO1. This may be due to an enhanced capacity for direct signaling or paracrine effects, which lead to the activation of key transcription factors like IKZF1 and PU.1 that drive the tolerogenic program. The mechanism underlying tolerogenic DC generation following NK cell engagement remains to be further studied and clarified. While we hypothesize that IDO1 and IL-10 are critical for DC phenotype conversion, it is uncertain whether the increases in IDO1 and IL-10 result from direct NK cell contact and NK-derived secretory factors, or from DCs phagocytosing apoptotic cells—a process known as efferocytosis, which is reported to promote immune tolerance marked by increase IL-10.73^,^74 Nevertheless, those proposed mechanisms are not mutually exclusive, and both support that NK-induced DC reprograming is contact-dependent and correlates with the killing efficacy of NK cells, suggesting that the depletion of proinflammatory DCs and induction of tolerogenic DCs are inseparable events. Given these complexities, the induction of tolerogenic DCs appears pivotal during NK-mediated killing in DC-depletion therapies, potentially mitigating safety concerns about off-target effects from activated NK cells.
PU.1 is a critical transcription factor necessary for the differentiation and maturation of both cDCs75 and MoDCs.76 Enforced overexpression of PU.1 in monocytes promotes their differentiation toward DCs, as indicated by an upregulation of DC maturation markers, such as CD11c,77 CD40,76 and proinflammatory cytokines including IL-1276 and TNF-α.78 Although the mechanisms of how PU.1 activity is suppressed in NK-programmed DCs remain to be further elucidated, previous reports suggest that IL-10 enhances the expression of the transcription factor c-Maf and suppresses PU.1 expression in human monocytes.76 Given our observation that IL-10 is crucial for both NK-mediated DC depletion and reprogramming, we hypothesize that enhanced IL-10 signaling in the NK-engaged DCs may contribute to the silencing of PU.1 activity, which deserves further study. Additionally, our data show that knockout of IKZF1 in MoDCs significantly increases IL-10 production. Interestingly, IKZF1 has been reported as a key transcription factor that, through binding to Foxp3, supports Treg function for suppressing IFN-γ production and promoting IL-10 secretion in CD4^+^ T cells.79 However, our findings suggest that IKZF1 plays an opposing role in MoDCs, where it promotes proinflammatory cytokine production and suppresses DC-derived IL-10. Indeed, heterozygous loss-of-function mutations of IKZF1 in humans are associated with reduced numbers of plasmacytoid DCs and cDC1s.80 Lenalidomide, a drug that induces proteasomal degradation of IKZF1, has also been shown to reduce the expression of proinflammatory cytokines, including TNF-α, IFN-α, and IL-12, while promoting IL-10 production in human DCs and monocytes.80 Similarly, murine IKZF1-deficient macrophages produce more IL-10 but less IL-6 and TNF-α following bacterial infection, suggesting IKZF1 functions as a negative regulator in myeloid APC tolerance.81 In summary, we propose that sustained suppression of PU.1 and IKZF1 activities is a critical mechanism for maintaining DC tolerance induced by NK cell engagement under inflammatory conditions.
External transplantation of tolerogenic DCs has been explored in various autoimmune diseases, though challenges in maintaining their tolerogenic phenotype in inflammatory environments hinder their widespread application.82 To address these obstacles, various DC-targeted approaches have been proposed for constant in situ generation of tolerogenic DCs with different payloads, such as mRNA for PD-L1 (eg Moderna’s mRNA-6981) and IL-10,83 and siRNA for IRF5.84 Our findings highlight the potential of using cell-depleting antibodies to not only eliminate proinflammatory DCs but also to instigate immune tolerance through NK-mediated reprogramming. This insight complements the mechanism of actions for developing DC-targeted antibody therapies and might also serve as a valuable biomarker in clinical trials assessing multiple myeloid APC-depleting antibodies.
The observation of increased pathogenic myeloid cells in autoimmune conditions is supported by multiple methodologies and across several indications, which lends consistency and credibility to the central finding. However, a limitation of our study is the limited number of human samples, and further work is needed to determine how generalizable these findings are in various patient populations and disease stages.
Supplementary Material
vlag007_Supplementary_Data
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