Blockade of DC-SIGN⁺ Tumor-Associated Macrophages Reactivates Antitumor Immunity and Improves Immunotherapy in Muscle-Invasive Bladder Cancer

Bladder cancer represents a most frequently diagnosed cancer worldwide with an estimated 549,000 new cases and 200,000 caner-related deaths annually (1). Bladder cancer is more prevalent in western countries, and ranks the fifth most common tumor in the United States (2). Patients with early-stage bladder cancer that have not invaded through the inner layer of the bladder wall have a 5-year survival rate of over 90% (3). However, approximately 25% of patients are diagnosed with advanced muscle-invasive bladder cancer (MIBC), indicating that tumors have already spread into the muscle wall of the bladder (4). In contrast to a favorable prognosis of early-stage bladder cancer, the 5-year overall survival rate of patients with MIBC is roughly 50%–60% (5, 6). Thus, elucidating the mechanisms leading to malignant development of MIBC is of key importance to improve the outcome of bladder cancer.

Immune evasion is a prerequisite for carcinogenesis (7). In this regard, aberrant function of macrophages plays an indispensable role in facilitating tumor cells to evade immune destruction. Studies have indicated that macrophages residing in tumor tissues, termed tumor-associated macrophages (TAM), may exert complicated roles in tumor immunity and progression. TAMs may exhibit proinflammatory (M1) or anti-inflammatory (M2) phenotypes, depending on the cytokine microenvironment in tumor tissues. While M1 TAMs may play an antitumor role and are associated with good prognosis, M2 TAMs may facilitate protumorigenic effects by virtue of inducing anti-inflammatory response and immune tolerance (8). In addition to traditional M1/M2 classification, recent studies revealed that assorted molecular signatures are critically implicated in the function of TAMs and tumor progression (9). For instance, it is recently reported that programmed cell death protein 1 (PD-1) is expressed both in M1 and M2 TAMs, and targeting PD-1⁺ TAMs may enhance macrophage phagocytosis and retard tumor growth (10). Coinciding with these findings, manipulation of TAMs, such as blockade of anti-inflammatory receptors on TAMs, has been shown to elicit considerable antitumor activities (11, 12).

Dendritic cell–specific C-type lectin (DC-SIGN), also termed CD209, was initially identified as a DC surface protein that binds to HIV-1 envelope glycoprotein gp120 (13). Later studies suggested that DC-SIGN is critically involved in the regulation of DC function, antigen presentation, T-cell immune response, and pathogen immune escape (14–16). DC-SIGN is the receptor of the glycan antigen Lewis x and may signal through IKKϵ- and CYLD-dependent Bcl3 pathway to initiate the expression of a variety of cytokines, including IL10, TGFβ, and IL27 (17). Notably, multiple studies uncovered an apparent distribution of DC-SIGN on macrophages (18, 19). DC-SIGN⁺ macrophages may exert an anti-inflammatory function following Mycobacterium tuberculosis infection (20).

In this study, we reported that DC-SIGN is predominately expressed in macrophages in MIBC. The level of DC-SIGN⁺ TAMs, but not total TAMs, is an independent prognostic factor in MIBC. In addition, we found that DC-SIGN⁺ TAM infiltration was positively associated with CD8⁺ T-cell tolerance. Of note, blockade of DC-SIGN suppresses the secretion of anti-inflammatory cytokines from macrophages, consequently leading to enhanced T-cell cytotoxic activities against tumor cells. Our findings provide a theoretical basis for targeting DC-SIGN⁺ TAMs as a promising therapeutic strategy for MIBC.

Two independent patient cohorts, comprising of 393 patients with bladder cancer who underwent radical cystectomy at Zhongshan Hospital of Fudan University (Shanghai, China) from 2002 to 2014 (n = 215) and Fudan University Shanghai Cancer Center from 2008 to 2012 (n = 178), were enrolled in this study. Patients meeting the following criteria were included: age range 18–80 years, pathologically diagnosed as MIBC according to the American Joint Committee on Cancer/Unio Internationale Contra Cancrum tumor–node–metastasis staging system 7th edition, complete follow-up data, and no autoimmune disease or multi-primary cancers. None of the patients enrolled in this study had participated in any clinical trials or received chemotherapy or radiotherapy before surgery. Two duplicated cases, 19 patients with non-urothelial carcinoma, 95 patients with non-MIBC, and 18 follow-up data unavailable patients were excluded. Two specimens were lost on the tissue microarray (TMA) when performing IHC. As a result, 257 eligible patients with MIBC were included [Zhongshan Hospital (ZH), n = 141; Fudan University Shanghai Cancer Center (FUSCC), n = 116]. This study was conducted according to the ethical principles stated in the Declaration of Helsinki, consent and approved by the Clinical Research Ethics Committee of Zhongshan Hospital and the Ethics Committee of Fudan University Shanghai Cancer Center, and written informed consent was obtained from every patient. Detailed flowchart of patient selection is illustrated in Supplementary Fig. S1.

After surgery, 119 patients (46.3%) received cisplatin-based combination chemotherapy and lasted at least one therapeutic cycle. Follow-up principle was based on the European Association of Urology guidelines for MIBC. Patients received follow-up visits at the hospital and subsequent examinations as follows: physical examination, urine cytology, chest imaging, and abdominal ultrasound or CT scan every 3–4 months for 1 year, then every 6 months for additional 2 years, and once per year afterwards. All follow-up data were collected from the date of surgery to July 2016. The overall survival (OS) was defined as the time period between surgery and death or last follow-up. The recurrence-free survival (RFS) was defined as the time period between surgery and the first recurrence, a new occurrence of MIBC, or death from any case. Detailed patients' characteristics are presented in Supplementary Table S1.

All 90 fresh human specimens were obtained from patients with MIBC who underwent radical cystectomy in four hospitals (Zhongshan Hospital, Shanghai, China; Ruijin Hospital, Shanghai, China; Fudan University Shanghai Cancer Center, Shanghai, China; and Shanghai General Hospital, Shanghai, China). Formal informed consent was received from every patient. All procedures of specimen collection, handling, and analysis were conducted in accordance with the ethical principles stated in the Declaration of Helsinki and approved by institutional review board and ethics committee of each hospital. Single-cell suspension was prepared as described previously (21). Briefly, fresh tumor tissues were washed with normal saline to remove the blood, immediately put into prechilled sterile tissue preservation solution (MACS Tissue Storage Solution, Miltenyi Biotec), and transported to the laboratory on ice. The necrotic and fatty tissues were removed from the tumor tissues. The tissue pieces were then disaggregated by gentle stirring in RPMI Medium 1640 containing 1 mg/mL Collagenase (Sigma Aldrich), 0.01% hyaluronidase (Sigma Aldrich), and 0.002% DNase I (Roche) for 2 hours at 37°C. Then, the tumor suspension was filtered through a 70-μm cell strainer (BD Falcon) and washed with Stain Buffer (BD Biosciences). Lysing Buffer (BD Biosciences) was used to remove red blood cells. Cells were not authenticated because of the fact that they are patient-derived cell samples. Cell suspension was Mycoplasma-free tested using PlasmoTest Mycoplasma Detection Kit (InvivoGen). Single-cell suspension was used for flow cytometry analysis, flow separation, and intervention experiments within 2 hours. In vitro intervention culture model, single-cell suspension was cultured for 12 hours in RPMI1640 medium containing 10% FBS and corresponding isotype and neutralizing antibodies.

All tissues were formalin-fixed and paraffin-embedded before subjected to the preparation of MIBC TMAs. IHC protocol has been described previously (22). Briefly, TMAs were deparaffinized and hydrated, heated with an autoclave in sodium citrate buffer for antigen retrieval, and incubated in PBS buffer containing 1% BSA and 5% serum for antigen blocking. Incubation with the first primary antibodies was carried out at 37°C for 2 hours, followed by horseradish peroxidase–labeled secondary antibody incubation and visualization using DAB reagent. Subsequently, the TMAs were incubated with the second primary antibodies overnight at 4°C before proceeding to an alkaline phosphatase–labeled secondary antibody and Vector Blue staining.

For immunofluorescence staining, the sections were incubated with primary antibodies for 2 hours at 37°C, followed by incubation with FITC- and TRITC-conjugated secondary antibodies for 2 hours at 37°C. Finally, the slides were mounted with Antifade Mounting solution containing DAPI. All antibodies and reagents are summarized in Supplementary Table S2. All stained tissues were examined under Leica DM6000 B Microsystems independently by two pathologists who were blind to the clinical and follow-up data.

For CD68⁺ macrophage density, ≤103 per mm² was defined as low infiltration and >103 per mm² was defined as high infiltration. For DC-SIGN⁺ TAM density, ≤42 per mm² was defined as low infiltration and >42 per mm² was defined as high infiltration. For the CD8⁺ T-cell density, ≤96 per mm² was defined as low infiltration and >96 per mm² was defined as high infiltration.

Single-cell suspension was incubated with Human BD Fc Block (BD Biosciences) and then stained with the indicated mAbs for 30 minutes at 4°C in dark. When necessary, before staining with antibodies against intracellular proteins or transcription factors, cells were pretreated with Fixation/Permeabilization Solution Kit or Transcription Factor Fixation/Permeabilization Buffer (BD Biosciences), respectively, according to the manufacturer's instructions. Stained cells were washed and resuspended in cell staining buffer. For the analysis of TAMs, cells were pregated using CD45 and CD68 antibodies before subjected to the analysis of cell markers, flow cytometry was performed using a BD FACSCelesta flow cytometer, and data were analyzed using FlowJo Software (Tree Star). All flow cytometry antibodies and reagents are summarized in Supplementary Table S2.

Total RNA was extracted from fresh tissue samples using TRizol Reagent (Invitrogen) according to the manufacturer's protocol. cDNA was synthesized using a PrimeScript RT Reagent Kit (Takara, RR047A). qRT-PCR was performed on an Applied Biosystems StepOnePlus Real-Time PCR System using a SYBR Green Premix Ex Taq II Kit (Takara, RR820A). The qRT-PCR primers were as follow: DC-SIGN (5′-GCA GTC TTC CAG AAG TAA CCG C-3′; 5′-GCT CTC CTC TGT TCC AAT ACT GC-3′) and β-actin (5′-CAC CAT TGG CAA TGA GCG GTT C-3′; 5′-AGG TCT TTG CGG ATG TCC ACG T-3′). The relative expression of target genes was calculated using the 2^−ΔΔC_t method (using β-actin as a calibrator).

DC-SIGN⁺ TAMs (CD45⁺ CD14⁺ DC-SIGN⁺) and DC-SIGN⁻ TAMs (CD45⁺ CD14⁺ DC-SIGN⁻) were freshly isolated from four MIBC tumors using the MoFlo XDP (Beckman Coulter) and immediately lysed in lysis buffer per 500 cells. cDNA libraries were prepared using standard Smart-seq2 protocols (23). mRNA profiles were generated using Illumina NovaSeq 6000 platform.

Sequencing data were aligned to the human transcriptome using HISAT2 software. Thereafter, the differentially expressed genes between DC-SIGN⁻ TAMs and DC-SIGN⁺ TAMs were analyzed using R software (version 3.6.0) and the edgeR package (version 3.26.8; ref. 24). The biological process pathway annotation was classified using DAVID resources (https://david.ncifcrf.gov/tools.jsp; ref. 25). Gene set enrichment analysis was used for gene functional annotation. Heatmaps of expression levels were created using Morpheus tool (https://software.broadinstitute.org/morpheus/).

The Cancer Genome Atlas (TCGA) BLCA mRNA and clinical data, including RNA-seq and clinicopathologic data for 412 tumors, were acquired from protein atlas website (https://www.proteinatlas.org/). The exclusion criteria were as follows: non-MIBC; preoperative chemotherapy; lack of follow-up data; and lack of RNA-seq data. Finally, 391 patients' data were enrolled in this investigation. The median value of DC-SIGN expression in MIBC data was set as the cut-off value.

Single-cell suspension dissociated from fresh MIBC tissues was randomly divided into four groups (isotype controls, anti-DC-SIGN, Pembrolizumab, and anti-DC-SIGN plus pembrolizumab) and was cultured for 12 hours in RPMI1640 medium containing 10% FBS and corresponding isotype and neutralizing antibodies. The antibodies included in this experiment were IgG2B isotype control (10 mg/mL, Clone 20116, R&D Systems), anti-DC-SIGN (20 mg/mL, Clone 120507, R&D Systems), and pembrolizumab (5 μg/mL, Selleck). After overnight culture, cells were subjected to flow cytometry analysis to examine the apoptosis of tumor cells by using FITC Annexin V Apoptosis Detection Kit I (BD Biosciences) or corresponding FCAS antibodies.

Patient baseline characteristics and disease factors were summarized using descriptive statistics. Results are expressed as mean ± SD. Two-sided Pearson χ² test or adjusted χ² test was used for categorical variables. Unpaired t test or paired t test or Mann–Whitney U test was used for continuous variables. Spearman correlation was employed to evaluate the correlation between different variables. Kaplan–Meier method was used to determine OS and RFS. Log-rank test was used to compare survival between two groups. Cox regression was used for multivariate analyses with hazard ratios and 95% confidence intervals. Interaction analysis was also conducted using Cox regression. A two-tailed P < 0.05 was considered statistically significant.

The cut-off values of DC-SIGN⁺ TAMs as prognostic biomarkers were defined according to the OS data of the ZH cohort and then applied in the FUSCC cohort. To obtain the best prognostic efficacy, X-Tile Software (Yale University, version 3.6.1) was used as described previously (26).

To decipher the involvement of DC-SIGN in MIBC development and prognosis, the colocalizations among DC-SIGN and different immune cell markers were first examined. Immunofluorescence assay revealed that DC-SIGN mostly colocalized with macrophage marker CD68 (Fig. 1A). The number of DC-SIGN⁺ TAMs varied distinctly in MIBC tissues and was apparently correlated with enhanced tumor stages in MIBC (Fig. 1B and C). We determined the mRNA level of DC-SIGN using RT-PCR analysis and found close correlations among mRNA expression of DC-SIGN, the infiltration of DC-SIGN⁺ cells, and DC-SIGN⁺ TAMs in these specimens (Supplementary Fig. S2A and S2B). We also found that neither macrophage infiltration nor CD45⁺ leucocyte infiltration correlated with tumor stages (Supplementary Fig. S2C and S2D). Furthermore, flow cytometry analysis also validated that MIBC specimens exhibited dramatically higher levels of DC-SIGN⁺ TAMs, compared with adjacent nontumor tissues (Fig. 1D and E).

In addition, the prognostic merit of DC-SIGN⁺ TAMs was also assessed. Two independent cohorts of MIBC cases were enrolled in this study. Notably, both cohorts indicated that high levels of DC-SIGN⁺ TAMs, but not total TAMs, predicted unfavorable prognosis in patients with MIBC (Fig. 2A–D). As predicted, multivariate regression analyses indicated that DC-SIGN⁺ TAMs may serve as an independent prognostic factor to predict unfavorable survival of patients with MIBC (Fig. 2E and F). Because our aforementioned data showed that the number of DC-SIGN⁺ TAMs was associated with MIBC stages, we performed Kaplan–Meier survival analysis separately in patients with pT2 stage tumors and pT3+4 stage tumors. As shown in Supplementary Fig. S3A–S3D, patients with high-DC-SIGN⁺ TAM infiltration displayed significantly worsened prognosis both in pT2 and pT3+4 MIBC groups, compare with those with low-DC-SIGN⁺ TAM infiltration. We also performed interaction test between DC-SIGN macrophages and pN stage, and found that DC-SIGN⁺ TAM infiltration was independent from lymph node metastases (Supplementary Fig. S3E). Moreover, when DC-SIGN⁺ TAMs were considered as a continuous variable, multivariable cox regression analysis validated that DC-SIGN⁺ TAM infiltration was an independent prognostic parameter in predicting the survival of patients with MIBC (Supplementary Table S3). These findings imply that DC-SIGN⁺ TAMs may contribute to the progression and poor prognosis of MIBC.

Adjuvant chemotherapy has been widely utilized in the treatment for patients with MIBC, despite limited evidence in favor of adjuvant chemotherapy (27). Thus, we investigated the relationship between DC-SIGN⁺ TAM infiltration and adjuvant chemotherapy benefit in patients with MIBC. As shown in Fig. 3A and B, adjuvant chemotherapy did not provide survival benefit in patients with pT2 stage MIBC. However, after dividing patients into high- and low-DC-SIGN⁺ TAM–infiltrating groups, we found that adjuvant chemotherapy led to a favorable outcome in patients with low-DC-SIGN⁺ TAM infiltration, whereas no overall beneficial effect was revealed in patients with high-DC-SIGN⁺ TAM infiltration (Fig. 3C–F). Next, we performed Cox proportional hazards regression analysis to assess the relationship between DC-SIGN⁺ TAM levels and adjuvant chemotherapy benefit in patients with MIBC. As shown in Supplementary Table S4, high-DC-SIGN⁺ TAM infiltration was strongly associated with unresponsiveness to adjuvant chemotherapy in MIBC.

To further determine the role of DC-SIGN⁺ TAMs in MIBC prognosis and immunity, we sought to characterize DC-SIGN⁺ TAMs. Using flow cytometry analysis, we first found that DC-SIGN⁺ TAMs largely had high expression of CD206, a M2 macrophage marker (Fig. 4A and B). In contrast, only a minor proportion of DC-SIGN⁻ TAMs had apparent expression of CD206 (Fig. 4B). In addition, DC-SIGN⁺ TAMs express low levels of proinflammatory cytokines, including IL12 and TNFα compared with DC-SIGN⁻ TAMs (Fig. 4C and D). Of note, the levels of immunosuppressive cytokines, including CCL22, IL10, TGFβ, and PD-L1, were strongly enriched in DC-SIGN⁺ TAMs (Fig. 4E and F). These findings suggest that DC-SIGN⁺ TAMs may elicit immunosuppressive roles during MIBC progression.

We further characterized the transcriptomic profiling of DC-SIGN⁺ TAMs through RNA-seq technique. DC-SIGN⁻ TAMs and DC-SIGN⁺ TAMs were freshly isolated from MIBC tumors and subjected to single-read deep sequencing (Supplementary Table S5). Compared with DC-SIGN⁻ TAMs, DC-SIGN⁺ TAMs exhibited apparently enriched expression signature patterns involved in immune signaling pathway, including chemokine, C-type lectin, adoptive immune response, and inflammatory response (Fig. 5A and B). However, we did not see apparent enrichments in DC-SIGN⁻ TAMs, probably because these TAMs are heterogeneous and may contain distinct subtypes (Fig. 5A and B). In addition, GESA analysis also confirmed that multiple M2-polarized macrophage-associated signaling pathways, including STAT6, cholesterol metabolism, PI3K–AKT–mTOR, and hypoxia signaling pathways, were hyperactivated in DC-SIGN⁺ TAMs (Fig. 5C; refs. 28, 29). Moreover, we found that compared with DC-SIGN⁻ TAMs, the expression profiles of transmembrane receptors, cytokines, and their intracellular signal transducers were altered largely toward M2-like phenotype (Fig. 5D and E). These findings suggested that DC-SIGN⁺ TAMs may mainly exhibit an M2-like anti-inflammatory phenotype.

Next, the involvement of DC-SIGN⁺ TAMs in MIBC immune microenvironment was investigated. Through IHC analysis, we found that MIBC with high-DC-SIGN⁺ TAMs exhibited higher levels of CD8⁺ T, CD4⁺ T, regulatory T (Treg) leucocytes, as well as neutrophils, as compared with tumors with low-DC-SIGN⁺ TAM infiltration (Fig. 6A). Flow cytometry analysis also validated that the levels of CD8⁺ cells and CD4⁺ FoxP3⁺ cells were significantly higher in DC-SIGN⁺ TAM–infiltrating MIBC specimens, as compared with low-DC-SIGN⁻ TAM–infiltrating samples (Fig. 6B). This indicated an association between DC-SIGN⁺ TAM infiltration and T-cell immunity in MIBC. Next, we analyzed whether DC-SIGN⁺ TAM infiltration may influence the prognostic merit of CD8⁺ T killer cells in MIBC. As shown in Fig. 6C, when analyzing low- and high-DC-SIGN⁺ TAM–infiltrating groups separately, it is noteworthy that CD8⁺ cells predicted a highly favorable prognosis in patients with low-DC-SIGN⁺ TAM infiltration, whereas high-DC-SIGN⁺ TAM infiltration seemed to abrogate the beneficial effect of CD8⁺ cells. These findings implicate that DC-SIGN⁺ TAMs may potentially facilitate CD8⁺ cells immune tolerance in MIBC. Subsequently, the associations among the infiltrations of DC-SIGN⁺ TAMs and different subtypes of CD8⁺cytotoxic T lymphocytes (CTL) were investigated. As shown in Fig. 6D, MIBC specimens with high-DC-SIGN⁺ TAM infiltration exhibited reduced levels of PRF1⁺, GZMB⁺, and IFNγ⁺ CD8⁺ lymphocytes, but increased proportions of TIGIT⁺ and LAG3⁺ CD8⁺ lymphocytes, compared with samples with low-DC-SIGN⁺ TAM infiltration. Overall, these results explicitly imply that DC-SIGN⁺ TAMs may suppress the proportion of CTLs, whereas promote the accumulation of suppressor T lymphocytes.

We then investigated the effects of abrogating DC-SIGN⁺ TAMs on immune microenvironment in MIBC using a DC-SIGN–neutralizing antibody. After incubating with DC-SIGN–neutralizing antibody for 12 hours, single-MIBC cell suspensions were subjected to flow cytometry analysis to determine the phenotypes of DC-SIGN⁺ TAMs. As shown in Fig. 7A, the expression of proinflammatory cytokines, including IL12 and TNFα, were remarkably upregulated after blocking DC-SIGN, whereas the levels of anti-inflammatory molecules were significantly downregulated. In addition, we uncovered that the total number of CD8⁺ CTLs was increased, while Tregs declined following neutralizing DC-SIGN (Fig. 7B).

Furthermore, the characteristics of CD8⁺ CTLs were investigated. Compared with the control group, blockade of DC-SIGN led to significant upregulation of GZMB⁺, INFγ⁺, and PRF1⁺ CD8⁺ CTLs, whereas the numbers of TIGIT⁺ and LAG3⁺ CD8⁺ cells declined markedly (Fig. 7C). This suggests that blockade of DC-SIGN may promote the antitumoral activity of CD8⁺ CTLs. Moreover, we found that in TCGA cohort, the mRNA level of DC-SIGN is highly correlated with that of PD1, suggesting an involvement of DC-SIGN⁺ TAMs in PD-L1–PD-1 pathway–induced CTL inactivation (Fig. 7D). Indeed, this piece of data were validated by our results showing that the numbers of DC-SIGN⁺ TAMs and PD1⁺ cells were strongly correlated in MIBC specimens. As such, we speculated that blockade of DC-SIGN may exert a synergistic role with PD-1 inhibitor pembrolizumab in mediating CTL activation and tumor cell elimination. As predicted, we found that treatment with DC-SIGN–neutralizing antibody and pembrolizumab alone can promote the proliferation of CD8⁺ CTLs (Fig. 7E). Of great intrigue, the combination of DC-SIGN–blocking antibody and pembrolizumab led to significantly enhanced proliferation of CD8⁺ CTLs, as compared with single-treatment groups. Likewise, we uncovered that DC-SIGN–neutralizing antibody and pembrolizumab had a synergistic effect in triggering the apoptosis of tumor cells (Fig. 7F). Taken together, these findings indicate that blockade of DC-SIGN may elicit a synergistic effect with pembrolizumab to facilitate the activation of CD8⁺ CTLs and the clearance of tumor cells.

The prognosis of MIBC remains largely unchanged in the past several decades, partly because of insufficient therapeutic options (30). Recent studies indicated that MIBC can be classified into multiple molecular subtypes, suggesting patients with MIBC may require personalized therapeutic strategies based on their molecular signatures (31). In this study, we uncovered that DC-SIGN⁺ TAMs were abundantly detected in a significant proportion of MIBC specimens. High level of DC-SIGN⁺ TAMs may contribute to adjuvant chemotherapy unresponsiveness and immune tolerance in MIBC. Targeting DC-SIGN using neutralizing antibody caused markedly impaired secretion of anti-inflammatory cytokines, resultantly leading to tumor-reactive T-cell responses in MIBC. These findings highlight the importance of targeting DC-SIGN⁺ TAMs in the immunotherapy of MIBC.

Since its initial characterization, the role of DC-SIGN in immune regulation has attracted significant research attention. DC-SIGN is predominately expressed in DCs and macrophages (18). Intriguingly, DCs were rarely found within bladder tumor specimens (32). This suggests that intratumoral DC-SIGN is mainly expressed on macrophage to participate in the regulation of MIBC progression, coinciding with our data showing that DC-SIGN was primarily colocalized with macrophage marker CD68 in MIBC tissues. Of note, studies on DCs revealed a pivotal role of DC-SIGN in regulating the expression and secretion of multiple cytokines, including IL10, IL12, IL27, and TGFβ (33). Thus, we speculate that macrophagal DC-SIGN may also contribute to the expression of the cytokines. Indeed, our results revealed that DC-SIGN⁺ TAMs expressed higher levels of these cytokines than DC-SIGN⁻ TAMs, and neutralizing DC-SIGN led to impaired expression of cytokines. These findings implicate that DC-SIGN expression may facilitate the expression and secretion of macrophagal anti-inflammatory cytokines in MIBC.

The role of TAMs in tumor immunity and progression remains controversial. Studies showed that TAMs displayed distinct prognostic values in different tumor types (34). Although M1/M2 activation could partially clarify the diverse roles of TAMs in tumor progression, its clinical significance remains to be fully elucidated. In addition, recent studies suggested that TAMs have complex molecular signatures and these molecular signatures may serve as prognostic indicators and therapeutic targets. For instance, a recent study showed that blockade of CSF1R could selectively eliminate immunosuppressive TAMs and promoted the antitumor activity of CD8⁺ CTLs (35). Recent investigations indicated that DC-SIGN expression on macrophages may promote immune tolerance (20, 36). However, although DC-SIGN has been frequently regarded as an alternative marker of M2 macrophages, DC-SIGN is detectable in both M1 and M2 macrophages (37, 38). More importantly, DC-SIGN is a functional receptor directly involved in the expression and secretion of multiple anti-inflammatory cytokines. In-line with these reports, our RNA-seq data confirmed that multiple signaling pathways involved in macrophage M2 polarization were specifically activated in DC-SIGN⁺ TAMs. However, we found that few pathway and expression signatures were enriched in DC-SIGN⁻ TAMs, suggesting that DC-SIGN⁻ TAMs may contain distinct TAM populations. These findings indicate that the characteristics of DC-SIGN⁻ and DC-SIGN⁺ TAMs may be different from classical M1/M2 phenotypes. Combined with these findings, we speculate that targeting DC-SIGN⁺ TAMs may serve as a promising approach to facilitate tumor immunity.

In summary, we for the first time report that DC-SIGN⁺ TAMs may serve as a prognostic indicator and therapeutic target in MIBC. DC-SIGN⁺ TAMs may play a pivotal role in facilitating tumor immune evasion via the secretion of anti-inflammatory cytokine and the induction of CD8⁺ T-cell tolerance. These findings expand our understanding of DC-SIGN⁺ TAMs in the immune modulation of invasive tumors and provide a rational approach to eliminate protumor immune environment.

No potential conflicts of interest were disclosed.

Conception and design: Y. Xia, L. Xu, W. Zhang, J. Xu

Development of methodology: Y. Zhu

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): B. Hu, Z. Wang, H. Zeng, Y. Chen, T. Wang, J. Wang, L. Liu, Y. Zhu

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): B. Hu, Z. Wang, H. Zeng, L. Xu, W. Zhang, J. Xu

Writing, review, and/or revision of the manuscript: B. Hu, Z. Wang, L. Xu, W. Zhang, J. Xu

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): Y. Qi, T. Wang, J. Wang, Y. Chang, Q. Bai, Y. Wang, B. Dai, J. Guo

Study supervision: Y. Zhu, W. Zhang, J. Xu

We thank Dr. Lingli Chen (Department of Pathology, Zhongshan Hospital, Fudan University, Shanghai, China) and Dr. Peipei Zhang (Department of Pathology, Ruijin Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, China) for their excellent pathologic technology help. This study was funded by grants from National Natural Science Foundation of China (grant no. 81671628 to W. Zhang; grant no. 31770851 to J. Xu; grant no. 81702496 to L. Liu; grant no. 81702497 to Q. Bai; grant no. 81702805 to Y. Xia; grant no. 81772696 to J. Guo; grant no. 81871306 to W. Zhang; grant no. 81872082 to Y. Zhu; grant no. 81902556 to Y. Wang; grant no. 81902898 to J. Wang; grant no. 81974393 to J. Guo), National Key R&D Program of China (grant no. 2017YFC0114303 to B. Dai), Shanghai Municipal Natural Science Foundation (grant no. 16ZR1406500 to B. Dai and grant no. 19ZR1431800 to L. Xu), Guide Project of Science and Technology Commission of Shanghai Municipality (grant no. 17411963100 to B. Dai), Shanghai Sailing Program (grant no. 18YF1404500 to Y. Chang; grant no. 19YF1407900 to J. Wang; grant no. 19YF1427200 to Y. Wang), Shanghai Municipal Commission of Health and Family Planning Program (grant no. 20174Y0042 to L. Liu; grant no. 201840168 to Y. Zhu; grant no. 20184Y0151 to Y. Wang), Fudan University Shanghai Cancer Center for Outstanding Youth Scholars Foundation (grant no. YJYQ201802 to Y. Zhu), Nantong Science and Technology Project (grant no. MS 12018095 to B. Hu), and Shanghai Cancer Research Charity Center (to B. Dai). All these study sponsors have no roles in the study design, in the collection, analysis, and interpretation of data.

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