The COX-2-PGE₂ pathway promotes tumor evasion in colorectal adenomas

Abstract

The mechanisms underlying the regulation of a checkpoint receptor, PD-1, in tumor-infiltrating immune cells during the development of colorectal cancer (CRC) are not fully understood. Here we demonstrate that COX-2-derived PGE₂, an inflammatory mediator and tumor promoter, induces PD-1 expression by enhancing NF-κB’s binding to the PD-1 promoter via an EP4-PI3K-Akt signaling pathway in both CD8⁺ T cells and macrophages. Moreover, PGE₂ suppresses CD8⁺ T cell proliferation and cytotoxicity against tumor cells and impairs macrophage phagocytosis of cancer cells via an EP4-PI3K-Akt-NF-κB-PD-1 signaling pathway. In contrast, inhibiting the COX-2-PGE₂-EP4 pathway increases intestinal CD8⁺ T cell activation and proliferation and enhances intestinal macrophage phagocytosis of carcinoma cells accompanied by reduction of PD-1 expression in intestinal CD8⁺ T cells and macrophages in Apc^Min/+ mice. PD-1 expression correlates well with COX-2 levels in human CRC specimens. Both elevated PD-1 and COX-2 are associated with poorer overall survival in colorectal cancer patients. Our results uncover a novel role of PGE₂ in tumor immune evasion. They may provide the rationale for developing new therapeutic approaches to subvert this process by targeting immune checkpoint pathways using EP4 antagonists. In addition, our findings reveal a novel mechanism explaining how NSAIDs reduce colorectal cancer risk by suppressing tumor immune evasion.

Introduction

Chronic inflammation is one of the known risk factors for colorectal cancer (CRC). Clinical and epidemiologic evidence suggests that long-term use of nonsteroidal anti-inflammatory drugs (NSAIDs) reduces the risk for developing colorectal adenomas and CRC and suppresses CRC development (1). Nonselective NSAIDs are known to exert certain anti-inflammatory effects by targeting cyclooxygenase enzymes (COXs) that include COX-1 and COX-2. COX-1 is constitutively expressed in most tissues. In contrast, COX-2 is an immediate-early response gene usually absent in healthy tissues and organs. However, it is highly inducible at sites of inflammation and in the tumor microenvironment of certain cancers, including CRC. For example, COX-2 expression is elevated in approximately 50% of colorectal adenomas and 85% of adenocarcinomas (2,3) and associated with poorer overall survival in patients with CRC (4). Both COX-1 and COX-2 convert arachidonic acid to prostanoids, including prostaglandins (PGs) such as PGE₂, PGD₂, PGF₂α, PGI_2, and thromboxane A₂ (TxA₂) by a two-step process. Among prostanoids, PGE₂ is the most abundant found in various types of human malignancies, including CRC, and its presence is often associated with a poor prognosis (5–8). More importantly, only PGE₂ and PGI₂ levels are elevated in CRC specimens compared to matched normal tissues (9).

Moreover, a urinary PGE₂ metabolite (PGE-M) has been evaluated as a biomarker for CRC (10,11). More importantly, a recent epidemiologic study revealed that regular use of NSAIDs, including aspirin, significantly reduced the risk of developing colorectal adenomas in women with high PGE-M levels. This effect was not observed in those with low PGE-M levels (12). PGE₂ exerts its cellular effects by binding to specific G-protein coupled receptors, EP1, EP2, EP3, and EP4. Among prostanoids, only PGE₂ has been demonstrated to promote intestinal adenoma formation and growth by induction of tumor epithelial cell proliferation and survival, as well as promoting angiogenesis in mouse models of intestinal adenomas (13). However, all the mechanisms that PGE₂ accelerates colorectal adenoma development are not fully understood. For example, relatively little is known about the impact of PGE₂ on host defenses against tumor cells within the tissue microenvironment.

Tumor immune evasion can occur through several mechanisms, including tumor cell resistance to immune attack, activation of immune checkpoint pathways in effector T cells, suppression of macrophage phagocytosis of tumor cells, and induction of massive infiltration of immunosuppressive cells. For example, tumor cells can evade immunosurveillance by directly impairing CD8⁺ T cell cytotoxic activity and proliferation through immune checkpoint receptors such as PD-1. Interaction of PD-1 with its ligands, PD-L1 and PD-L2, suppresses CD8⁺ T cell cytotoxicity and proliferation. It has been documented that a strong and persistent expression of PD-1 is observed in tumor-infiltrating effector T cells (14–16). Tumor-associated macrophages (TAMs) are a significant subpopulation of tumor-infiltrating immune cells (17). Increased infiltration of TAMs is recognized as a poor prognostic sign in CRC patients (18). A recent report revealed that PD-1 expression in macrophages inhibited their phagocytosis of tumor cells (19). However, the mechanisms by which PD-1 is regulated in CD8⁺ T cells and macrophages in the tumor microenvironment are still largely unknown.

Immunotherapies using checkpoint inhibitors offer great promise for treating some malignancies, but their effectiveness in many solid tumors has been disappointing. For example, the therapeutic effect of a PD-1 monoclonal antibody was mainly observed in patients with MSI CRC but not in patients with MSS CRC (20). However, MSI is only found in about 15% of sporadic CRC, whereas approximately 85% of sporadic CRC is classified as MSS. Interestingly, the elevation of COX-2 expression was observed in 79% of MSS CRC, but only 48% of MSI CRC (21). Other studies also revealed that MSI CRC has a low or absent COX-2 expression (22–24). One potential explanation for the poor efficacy of checkpoint inhibitors in MSS CRC is that the presence of COX-2-derived PGE₂ might attenuate their effect by reducing tumor-infiltrating CD8⁺ T cell abundance and cytotoxicity and/or by impairing TAM phagocytosis of tumor cells. In this study, we investigate whether activation of the COX-2-PGE₂ pathway induces tumor immune evasion by reducing tumor-infiltrating CD8⁺ T cell cytotoxicity and abundance via PD-1 and/or by impairing macrophage phagocytosis of tumor cells via PD-1.

Materials and Methods

Animal experiments

All animal experiments were completed according to our animal protocols approved by the Institutional Animal Care and Use Committee at the Medical University of South Carolina (MUSC). Apc^Min/+ mice were purchased from Jackson Laboratory (Cat#002020). For experiments evaluating celecoxib or Ono-AE3–208 treatment, Apc^Min/+ male mice at the age of 6 weeks old were randomly divided into two groups fed with a control diet, a diet containing 500ppm of celecoxib, or provided with water containing Ono-AE3–208 (10 mg/kg) for ten weeks. No significant sex variable exists in the intestinal tumor burden of Apc^Min/+ mice.

Isolation of immunocytes from organs

All fat and Peyer’s patches were removed from excised intestines under a dissecting microscope for intestinal immune cell preparation. Mouse normal intestinal tissues and adenomas were minced and digested with RPMI 1640 medium containing 5% FBS, 1 mM MgCl₂, 1 mM CaCl₂, 2.5 mM HEPES, and 200 units/ml collagenase I (Gibco). The immune cells from intestinal tissues were enriched by using a discontinuous (44% and 67%) percoll (GE) separation method. Isolated immune cells were subjected to Flow Cytometry or in vitro culture. Excised spleens were smashed using a 40 μm cell strainer. The red blood cells (RBCs) in the spleen were lysed with RBC lysis buffer (eBioscience). Mouse CD8+ T cells were isolated from the spleen using a mouse CD8+ T Cell Isolation Kit (Miltenyi Biotec Inc., San Diego, CA) according to the manufacturer’s instructions; isolated CD8⁺ T cells with a purity of 98% were subjected to experiments listed below.

Prostaglandin measurement

Intestinal tissues were homogenized in PBS with 10% 2,6-di-tert-butyl-p-cresol as previously described (25,26). The levels of prostaglandins, including PGE_2, were measured using mass spectrometry and normalized with protein concentration as previously described (25,26).

Cell culture and reagents

Isolated mouse splenic CD8⁺ T cells were cultured in RPMI 1640 medium with 10% FBS (Gibco, Cat#10082147), 25 mM HEPES, 100 IU/ml IL-2 (Peprotech, Cat#200–02), 1:100 100x non-essential amino acids (Sigma-Aldrich) and 1mM sodium pyruvate, and 50 μM β-mercaptoethanol. Mouse CD8⁺ T cells were activated by adding anti-CD3/CD28 mAb-coated beads (Dynabeads Mouse T-Activator CD3/CD28, Gibco) according to the manufacturer’s instructions. Then cells were starved for 24 hrs. Human monocytic THP-1 (Cat#TIB-202) and HCT-116 (Cat#CCL-24&) cells were obtained from ATCC (Manassas, VA) and maintained in RPMI 1640 with 10% FBS. THP-1 monocytes were treated with 50 mg/ml phorbol 12-myristate 13-acetate (PMA) 24 hrs to stimulate their differentiation into macrophages, followed by incubation for 24 hrs in RPMI medium. Macrophages were treated with 20 ng/ml IL-4 (R&D Systems) and 20 ng/ml IL-13 (R&D Systems) for 24 hrs. THP-1-derived macrophages were starved for 48 hrs followed by treatment with the indicated dose of PGE₂ (Cayman Chemical, Ann Arbor, Michigan), 0.1 μM Ly294002 (Calbiochem-Novabiochem Corp., San Diego, CA), 1 μM MK2206 (Selleck Chemicals), and/or 1 μM BAY110785 (ENZO) for the times indicated. For bone marrow-derived macrophages (BMDMs), bone marrow cells were flushed aseptically from the femurs of Apc^Min/+ mice (Jackson Laboratory, Cat#000664) and cultured in Falcon™ Petri dishes (BD Biosciences, Cockeysville, MD) with DMEM medium supplemented with 10% FBS and 10 ng/ml of macrophage colony stimulating factor (M-CSF) for 4 days. After reseeding cells, cells were cultured in DMEM with 10% FBS, 10 ng/ml M-CSF (Life Technologies), and 20 ng/ml IL-4 (R&D Systems) for 2 days. BMDMs were cultured in serum-free DMEM medium for 2 days and treated with the indicated dose of PGE₂ and/or inhibitors for indicated time for 24 hr. All cell lines were tested using a MycoProbe Mycoplasma Detection Kit (R&D) and authenticated before each experiment using the ATCC STR database.

Quantitative PCR

Total RNA was isolated from indicated cells using an RNeasy Mini Kit (Qiagene), and 1 μg of RNA was reversely transcribed to cDNA using High-Capacity cDNA Reverse Transcription Kits (Bio-Rad). Real-time q-PCR was performed with a SYB® Gene Expression Assay Mix (Bio-Rad) using a QuantStudio 7 Flex Real-Time PCR System (Thermo Fisher Scientific, Waltham, MA). The sequences of the specific PCR primers were as follows (5’ to 3’): mPdcd1 (PD-1), forward: CCTCTGACACTGTGAGCCAG, reverse: GCAGGTACCCTGGTCATTCA; hPDCD1, forward: CTCCGATGTGTTGGAGAAGC, reverse: CGGCCAGGATGGTTCTTAG; β-actin, forward: AGAAAATCTGGCACCACACC; reverse: AGAGGCGTACAGGGATAGCA. The relative expression of each target gene represents an average of triplicates that are normalized against the transcription levels of β-actin.

Western blotting

Whole-cell or nuclear extracts were prepared from the indicated cells in each experiment. Transfer membranes were blocked with 5% dry milk in TBS-T buffer for 1 hr and then incubated with anti-pAkt (1:1000, Cell Signaling, Cat#4058), anti-Akt (1:1000, Cell Signaling, Cat#4691), anti-NF-κB (1:1000, Santa Cruz, Cat#SC372), anti-GAPDH (1:5000, Cell Signaling, Cat#8884S), or anti-LaminA (1:1000, Santa Cruz, Cat#SC56137) antibody for overnight at 4 °C.

Flow cytometry analysis and sorting

For single color flow cytometry immunotypic analyses, each indicated single-cell suspension was incubated with anti-mPD-1-PE (1:100, BioLegend, Cat#135206), anti-hPD-1-PE (1:100), or the corresponding isotype controls in staining buffer (BD Biosciences). For multicolor flow cytometry immunotypic analyses, mouse intestinal immunocyte cells were incubated with the appropriate combination of the following antibodies in staining buffer at the following dilution for 30 min on ice: PD-1-PE (1:100), CD45-PE-Cy7 (1:250, BioLegend, Cat#103114), CD8-FITC (1:50, BioLegend, Cat#100706), CD4-AF700 (1:100, BioLegend, Cat#100536), CD3–Percep5.5 (1:100, Invitrogen, Cat#45–0031-82), and V450 (1:1500, Invitrogen, Cat#65–0863-14) for lymphocytes; PD-1-PE (1:100), CD45-APC (1:100, BioLegend, Cat#103112), CD11b-FITC (1:10, Miltenyi Biotech, Cat#130–081-201), Ly6G-Alexa700 (1:50, BD Biosciences, Cat#561236), CD11c-PE-Cy7 (1:80, BioLegend, Cat#117318), F4/80-Percep5.5 (1:100, Invitrogen, Cat#45–4801-32), V450 (1:1500) for granulocytes/monocytes. Dead cells were excluded using V450 staining. To analyze INFγ expression on CD8⁺ T cells, immunocyte cells were stained with cell surface markers for CD8⁺ T cells as described above. Then, the cells were fixed and permeabilized using a Cytofix/Cytoperm kit (BD Biosciences, Cat#554714) followed by intracellular cellular staining with anti-mouse INFγ-FITC antibody (1:50, BD Biosciences, Cat#554411) in permeabilization buffer for 30 min on ice. After incubation of antibodies, the cells were analyzed on a Fortessa X-20 cytometer (BD Biosciences). The flow cytometric profiles were analyzed by counting 30,000 events using FlowJo X software (Tree Star).

Chromatin Immunoprecipitation (ChIP) Assay

The Magna ChIP™ A/G Chromatin IP Kit (EMD Millipore, Cat#MAGNA0017) was used for ChIP assays according to the manufacturer’s instructions. Briefly, cells were treated with 1% formaldehyde for 10 min at room temperature to crosslink proteins to DNA, then quenched by adding glycine to 0.125 M for 5 min at room temperature. Chromatin was sonicated to an average length of 400–600bp. 5 μg of chromatin was immune-precipitated by 0.5 μg IgG or anti-NF-κB p65 antibody (Cell Signaling, Cat#8242). After being reverse crosslinked, the DNA was purified and eluted into 50 μl of elution buffer. Immunoprecipitated DNA was detected by PCR. The primers used for PCR were as follows (5’ to 3’): mPDCD1 (PD-1), forward: 5′-GTGAGACCCACACATCTCATTGC-3′ and reverse: 5′-CCTCACCTCCTGCTTGTCTCTC-3′ and hPDCD1 (PD-1), forward: 5’- AGA GAC ACA GAG GAG GAA GG −3’ and reverse: 5’-AGG GAC TGA GAG TGA AAG GT −3’. All ChIP assays were performed in three independent experiments.

Transient transfection and luciferase assays

The mouse PD-1 luciferase reporter plasmids with WT and mutant NF-κB binding sites (mNF-κB1 or mNF-κB2) were obtained from Dr. Jeremy Boss (27). Mutant NF-κB binding sites were generated by replacing NF-κB binding sites with the scrambled sequences. These scrambled sequences were predicted to have no NF-κB and other transcription factor binding potential (27). The human PD-1 promoter (−884 to +140) was cloned into a PGL3 vector (Promega, Cat#E1751). The NF-κB1 (GGGGATGGGCC) and NF-κB2 (GGGGAGACCCC) elements within the PD-1 promoter were mutated to mutant NF-κB1 (GTTTATGGGAA) and mutant NF-κB2 (GTTTAGACAAA) sites using the QuikChange II XL Site-Directed Mutagenesis Kit (Agilent Technologies). For dual-luciferase reporter assays, mouse CD8⁺ T cells or THP-1 cells were transfected with 5 μg of a mouse or human PD-1-Luc plasmid and 0.2 μg pRL-TK (Promega, Cat#E2231) DNA as a control using a Cell line Nucleofector Kit V (Lonza, Cat#VCA-1003) according to the manufacturer’s instructions. After transfection, CD8⁺ T cells were starved for 24 hrs and treated with 0.1μM PGE₂ for 24h. Transfected TPH-1 cells were treated with 50 ng/ml PMA for 24 hrs. TPH-1-derived macrophages were treated with 0.1 μM PGE₂ for 24 hrs after a 48 hr starvation period. Following treatment, cells were lysed using cell lysis buffer provided in the kit (Promega, Cat#E1960). Luciferase activity was measured using a Dual-luciferase reporter assay kit (Promega) with a Monolight 3010 luminometer (BD Biosciences/Pharmingen, San Diego, CA). The relative luciferase activity was determined and normalized to Renilla luciferase activity.

CFSE cell proliferation

Mouse splenic activated CD8⁺ T cells were labeled with 5 μM carboxyfluorescein succinimidyl ester (CFSE) (Life Technologies, Cat#C34554) according to the manufacturer’s instructions. CFSE-labeled CD8⁺ T cells were starved for 24 hrs and treated with 0.1 μM PGE₂, 20 μg/ml IgG, 20 μg/ml anti-PD-1 (Bio X Cell, Cat#BE0273), 0.1 μM Ly294002, 1 μM MK2206, and/or 1 μM BAY110785 for 48 hrs. For intestinal CD8⁺ T cells, immune cells isolated from intestinal normal tissues or tumors were labeled with 0.5 μM CFSE. CFSE-labeled immune cells were cultured in RPMI1640 medium with 10%FBS for 24 hrs. Then the cells were stained with the anti-CD8, anti-CD3 antibodies, and V450. CD8⁺ T cells proliferation was measured by a Fortessa X-20 cytometer.

Macrophage phagocytosis

THP-1-derived macrophages were starved for 4h and then treated with 0.1 μM PGE₂, 20 μg/ml IgG, 20 μg/ml anti-PD-1 (Nivolumab, BioVision, Cat#A137–100), 0.1 μM Ly294002, 1 μM MK2206, and/or 1 μM BAY110785 for 24 hrs. After treatment, 0.5 ×10⁶ macrophages were co-cultured with 1 ×10⁶ GFP-labeled HCT-116 cells for 3 hrs. For mouse intestinal macrophages, 5 × 10⁵ mouse intestinal immune cells were co-cultured with 2 × 10⁵ GFP-labeled MC26 cells for 3 hrs. Human cells were incubated with the antibodies indicated, including hCD45-PE-Cy7 and hCD11b-APC, as well as V450 in flow cytometry buffer. Mouse cells were incubated with antibodies, including mCD45-PE-Cy7, mCD11b-APC, and mF4/80-PerCP/Cy5.5 as well as V450 in flow cytometry buffer. Human macrophages that phagocytose GFP-labeled tumor cells were referred to as CD45⁺CD11b⁺GFP⁺ cells, whereas mouse macrophages that phagocytose GFP-labeled tumor cells were indicated as CD45⁺CD11b⁺F4/80⁺GFP⁺ cells by flow cytometry analysis. The phagocytic index was calculated as the percentage of CD45⁺CD11b⁺GFP⁺ cells in total CD45⁺CD11b⁺ cells for human cells and the percentage of CD45⁺CD11b⁺F4/80⁺GFP⁺ cells in total CD45⁺CD11b⁺F4/80⁺ cells for mouse cells.

Assays for CD8⁺ T cell cytotoxicity against tumor cells

Mouse splenic activated CD8⁺ T cells were treated with 30 U/ml IL-2, 0.1 μM PGE₂, 20 μg/ml IgG, 20 μg/ml anti-PD-1 (Bio X Cell), 0.1 μM Ly294002, 1 μM BAY110785, and/or 1 μM MK2206 for 48 hrs. After treatment, CD8⁺ T cells were co-cultured with 2.5 × 10⁵ MC26 in 6-well plate at ratios (E:T=2:1) for 16 hrs. Then, the cells were incubated with anti-mEpCAM-PE-Cy7 (1:1000, BioLegend, Cat#118216) and anti-mCD3-APC (1:100, BioLegend, Cat#103112) antibodies in staining buffer (Biolegend) for 30 min on ice. The cells were then washed twice with 1 ml of staining buffer and stained with propidium iodide (PI) using a TACS™ Annexin V-FITC Apoptosis Detection Kit according to the manufacturer’s instructions (R&D System). The EpCAM⁺CD3^-PI⁺ cells were dead epithelial cells and the percentage of dead epithelial cells in total epithelial cells (EpCAM⁺CD3^-) was presented.

Statistical analysis

All in vitro experiments were performed at least 3 times. All animal experiments were repeated at least twice. Results were expressed as mean + SEM. In general, we used the Bonferroni test by factorial analysis of variance to compare outcomes among multiple groups of mice. Student’s t-test or Mann-Whitney U test were used to compare two groups. For all tests, p<0.05 was considered statistically significant.

Data availability statement

The data generated in this study are available within the article and its supplementary data file. Raw data generated in this study are available from the corresponding author upon request. The data analyzed in this study were obtained from the TCGA Colorectal Adenocarcinoma Provisional dataset and in Gene Expression Omnibus (GEO) at GSE17537 and GSE17538.

RESULTS

The COX-2-PGE₂-EP4 pathway is involved in the induction of PD-1 expression in intestinal CD8⁺ T cells and macrophages

To determine whether the COX-2-PGE₂ pathway regulates immune checkpoint receptors, we first examined the effect of a COX-2 selective inhibitor, celecoxib, on the expression of immune checkpoint receptors in mouse intestinal immune cells. Treatment of Apc^Min/+ mice with celecoxib reduced PD-1 expression in CD8+ T cells, which are resident in both small and large intestinal adenomas and matched normal tissues (Fig. 1A). Interestingly, CD8⁺ T cell abundance in adenomas is much less than that found in normal tissues; however, celecoxib treatment restored CD8⁺ T cell abundance in intestinal adenomas as well as matched normal tissues (Fig. 1B). Celecoxib treatment also significantly reduced intestinal prostaglandin levels, including PGE₂ (Fig. S1A), providing evidence that PGE₂ levels correlated with CD8⁺ T cell abundance by induction of PD-1 in intestinal tumors and matched normal tissues. Similarly, celecoxib treatment also reduced PD-1 expression in macrophages, resident in both small and large intestinal adenomas, and matched normal tissues (Fig. 1C). In contrast, celecoxib treatment does not appear to affect PD-1 expression in other intestinal immune cells (Fig. S1B). CTLA4 expression in intestinal CD8⁺ T and CD4⁺ T cells was not significantly affected following celecoxib treatment as well (Fig. S1C).

Fig. 1. Inhibition of the COX-2-PGE2 pathway reduces PD-1 expression in intestinal CD8⁺ T cells and macrophages and decreases intestinal CD8⁺ T cell abundance.

Apc^Min/+ mice were treated with vehicle, celecoxib, or Ono-AE3–208 as described in the Methods. (A-B) The percentage of PD-1-positive CD8⁺ T cells in total CD8⁺ T cells (A) and the percentage of CD8⁺ T cells in total T lymphocytes (B) in both small intestinal (SI) and large intestinal (LI) tumors (T) and matched normal tissues (N) were analyzed by Flow Cytometry. (C) The percentage of PD-1-positive macrophages in total macrophages, which are resident in both SI and LI tumors and matched normal tissues, were analyzed by Flow Cytometry. (D-E) The percentage of PD-1-positive CD8⁺ T cells in total CD8⁺ T cells (D) and the percentage of CD8⁺ T cells in total T lymphocytes (E) in both small intestinal and large intestinal tumors and matched normal tissues were analyzed by Flow Cytometry. (F) The percentage of PD-1-positive macrophages in total macrophages, which are resident in both SI and LI tumors and matched normal tissues, were analyzed by Flow Cytometry. The error bar indicates + SEM. * indicates p<0.05.

Expression of EP2 and EP4 receptors is higher in human CRC specimens than in normal colon tissues (28–30). In contrast, the expression of EP3 receptors is reduced in human CRC specimens compared to adjacent normal colon tissues (30). Multiple lines of in vivo evidence further demonstrated that PGE₂ promotes intestinal adenoma formation and growth via EP2 and EP4 receptors, but not EP3 (31). The involvement of EP1 in intestinal tumorigenesis remains unclear because different groups have reported conflicting results (31). Importantly, immune cells mainly express EP2 and EP4 receptors. For example, macrophages express EP2 and EP4, but not EP1 or EP3 (32). Moreover, the EP4 receptor has a higher affinity for PGE₂ than EP1 or EP2 (33). Indeed, EP2 and EP4 mRNA levels were higher than EP1 and EP3 in splenic CD8⁺ T cells and bone marrow-derived macrophages (BMDM) isolated from Apc^Min/+ mice as well as in THP-1-derived macrophages (Fig. S2A). Notably, only EP4, but not EP2, is expressed on the cell surface of splenic CD8⁺ T cells and bone marrow-derived macrophages (BMDM) isolated from Apc^Min/+ mice (Fig. S2B). In human THP-1-derived macrophages, EP2 is marginally detectable, whereas EP4 is highly expressed (Fig. S2B). Therefore, we examined whether inhibition of PGE₂ signaling pathways by targeting its EP4 receptor has a similar effect as that seen following treatment with COX-2 inhibitors. Like celecoxib, treatment of Apc^Min/+ mice with an EP4 antagonist (Ono-AE3–208) led to a reduction of PD-1 expression in CD8⁺ T cells and macrophages, which are resident in both small and large intestinal adenomas and matched normal tissues accompanied by increased intestinal CD8⁺ T cell abundance (Fig. 1D–F). As expected, Ono-AE3–208 treatment did not affect PD-1 expression in other intestinal immune cells or CTLA-4 expression in intestinal CD8⁺ T and CD4⁺ T cells (Fig. S2C–D).

PGE₂ directly induces PD-1 expression via an EP4-PI3K-AKT-NF-κB pathway in CD8⁺ T cells and macrophages

To further evaluate whether PGE₂ directly induces PD-1 expression in CD8⁺ T cells and macrophages, in vitro experiments were performed. PGE₂ induces PD-1 mRNA levels in mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages (Fig. 2A–B) as well as mouse BMDMs (Fig. S3A–B). Significantly, PGE₂ induces PD-1 expression levels on the cell surface (Figs. 2C and S3C–D). Moreover, time- and dose-dependent effects of PGE₂ on PD-1 expression were observed in these cells (Figs. 2A–C and S3A–D).

Fig. 2. PGE₂ induces PD-1 expression via a PI3K-NF-κB pathway in CD8⁺ T cells and macrophages.

(A-B) PGE₂ induces PD-1 expression at mRNA in a dose- and time-dependent manner in mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages. (C) After PGE₂ treatment, the percentage of PD-1-positive cells in total mouse splenic activated CD8⁺ T cells or total THP-1-derived macrophages were determined by Flow Cytometry. (D) After PGE₂ treatment, the levels of phosphor-AKT and the nuclear NF-κB p65 in mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages were measured by Western blotting. The images are representative of three independent experiments. The band density of pAkt in CD8⁺ T cells was normalized by the band density of Gapdh. The results were reported as a mean of fold-induction from three independent experiments (middel panel). (E). The effect of a PI3K inhibitor (Ly294002), an AKT inhibitor (MK2209), and an NF-κB inhibitor (BAY110785) on PGE₂ induction of PD-1 expression on the cellular surface. The error bar indicates ± SEM. *p<0.05.

The PD-1 promoter contains two potential NF-κB binding sites located at NF-κB1 (−560 to −551) and NF-κB2 (−240 to −231) for human and NF-κB1 (−1265 to −1256) and NF-κB2 (−974 to −965) for mouse. LPS has been shown to induce PD-1 expression via binding of NF-κB to the NF-κB2 binding site within the PD-1 promoter in mouse macrophages (27). In addition, our previous results demonstrated that PGE₂ could activate NF-κB via an EP4-dependent PI3K pathway in colorectal carcinoma cells (34,35). These results prompted us to postulate that PGE₂ induces PD-1 expression by activating NF-κB via an EP4-PI3K pathway. To test this hypothesis, we first examined whether PGE₂ induces AKT and NF-κB activation. As shown in Fig. 2D, treatment of mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages with PGE₂ led to increased phospho-AKT levels, and nuclear translocation of NF-κB. PGE₂ induction of NF-κB nuclear translocation was also observed in mouse BMDMs (Fig. S3E). Moreover, inhibition of PI3K, AKT, or NF-κB by each respective inhibitor attenuated the effect of PGE₂ on induction of PD-1 expression in mouse splenic activated CD8⁺ T cells, THP-1-derived macrophages, and mouse BMDMs (Figs. 2E and S3F). Interestingly, Ly294002 treatment at 0.1 μM inhibited PGE₂ induction of PD-1 mRNA expression in these cells (Fig. S3G).

To further examine whether PGE₂ induces PD-1 transcription by increasing binding of NF-κB to cis-elements in the PD-1 promoter, ChIP and PD-1 luciferase reporter assays were performed. As shown in Fig. 3A–B, PGE₂ induced binding of NF-κB to the PD-1 promoter in both mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages. Moreover, PGE₂ induces PD-1 transcription in both mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages (Fig. S3H). As expected, site-directed mutations of the NF-κB2 binding element within the PD-1 promoter abolished PGE₂ induction of transcription in these cells, whereas mutations of the NF-κB1 binding site within the PD-1 promoter did not affect PGE₂ induction of transcription (Fig. 3C–D). These results demonstrate that only the NF-κB2 binding site is required for PGE₂ induction of PD-1 transcription. Since EP4, but not EP2, is expressed on the surface of these cells, our results reveal that PD-1 induction is regulated mainly via the EP4-PI3K-AKT-NF-κB pathway.

Fig. 3. PGE₂ induces PD-1 transcription by inducing the binding of NF-κB to the PD-1 promoter.

(A-B) Representative image of three independent ChIP assays for NF-κB binding to the PD-1 promoter in mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages after PGE₂ treatment. (C-D) The luciferase activity of WT and mutant PD-1 promoters in mouse splenic activated CD8⁺ T cells and THP-1-derived macrophages was measured after PGE₂ treatment. The error bar indicates ± SEM. *p<0.05.

PGE₂ suppresses CD8⁺ T cell proliferation and cytotoxicity and impairs macrophage phagocytosis of tumor cells via an EP4-PI3K-AKT-NF-κB-PD-1 pathway in vitro

We first evaluated the impact of PGE₂ on CD8⁺ T cell proliferation and cytotoxicity against tumor cells and found that PGE₂ inhibited proliferation of mouse splenic activated CD8⁺ T cells in a time-dependent manner (Figs. 4A). In addition, PGE₂ suppressed mouse splenic activated CD8⁺ T cell cytotoxicity against MC26 cells in a dose-dependent manner (Fig. S4A). Blockage of PD-1 signaling by using a neutralizing antibody attenuated the effect of PGE₂ on suppression of CD8⁺ T cell proliferation and cytotoxicity against MC26 cells (Fig. 4B–C). Moreover, inhibition of PI3K, Akt, or NF-κB by each respective inhibitor also blocked the effect of PGE₂ on suppression of CD8⁺ T cell proliferation and cytotoxicity against MC26 cells (Fig. 4B–C). Like CD8⁺ T cells, PGE₂ reduced THP-1-derived macrophage phagocytosis of HCT-116 cells (Fig. 4D) and mouse BMDM phagocytosis of MC26 cells (Fig. S4B). Treatment with a PD-1 monoclonal antibody, a PI3K inhibitor, an Akt inhibitor, or an NF-κB inhibitor attenuated the effect of PGE₂ on inhibition of THP-1-derived macrophage phagocytosis of HCT-116 cells (Fig. 4D) and BMDM phagocytosis of MC26 cells (Fig. S4B). These results demonstrate that PGE₂ suppresses CD8⁺ T cell proliferation and cytotoxicity and impairs macrophage phagocytosis of tumor cells via an EP4-PI3K-AKT-NF-κB-PD-1 pathway in vitro.

Fig. 4. PGE₂ suppresses CD8⁺ T cell proliferation and cytotoxicity and inhibits macrophage phagocytosis via a PI3K-Akt-NF-κB-PD-1 pathway.

(A) After PGE₂ treatment, the percentage of proliferating CD8⁺ T cells was measured by CFSE-based proliferation assays. (B) The effect of a PD-1 neutralizing antibody and an indicated inhibitors on PGE₂ reduction of CD8⁺ T cell proliferation. (C) Mouse splenic activated CD8⁺ T cells were treated with PGE₂, a PD-1 neutralizing antibody, and/or an indicated inhibitor. After treatment, CD8⁺ T cells were co-cultured with GFP-labeled MC26 cells. The percentage of dead MC26 cells in total MC26 cells was determined by Flow Cytometry. (D) THP-1-derived macrophages were treated with PGE₂, a PD-1 neutralizing antibody, and/or an indicated inhibitor. After treatment, THP-1-derived macrophages were co-cultured with GFP-labeled HCT-116 cells. The phagocytic index is the percentage of macrophages that phagocytose GFP-labeled tumor cells in total macrophages. The error bar indicates ± SEM. *p<0.05.

Blocking the COX-2-PGE₂-EP4 pathway inhibits tumor immune evasion by increasing intestinal CD8⁺ T cell activation and inducing macrophage phagocytosis

Since inhibition of the COX-2-PGE₂-EP4 pathway reduces PD-1 expression in intestinal CD8⁺ T cells and macrophages, it was conceivable that inhibition of this pathway could induce intestinal CD8⁺ T cell activation and macrophage phagocytosis of tumor cells. As shown in Fig. 5A–D, treatment of Apc^Min/+ mice with celecoxib or Ono-AE3–208 resulted in increased proliferation and IFNγ production in CD8⁺ T cells in intestinal adenomas and matched normal tissues, demonstrating that inhibition of the COX-2-PGE₂-EP4 pathway increases CD8⁺ T cell activation in intestinal adenomas and matched normal tissues. Treatment of Apc^Min/+ mice with celecoxib or Ono-AE3–208 also enhanced the ability of macrophages isolated from intestinal adenomas and matched normal tissues to phagocytize mouse MC26 cells (Fig. 5E–F). As expected, treatment of Apc^Min/+ mice with celecoxib or Ono-AE3–208 reduced both small and large adenoma burden (Fig. 5G–H). These results suggest that PGE₂ accelerates colorectal adenoma formation by suppressing CD8⁺ T cell cytotoxicity and macrophage phagocytosis against transformed epithelial cells via PD-1 in normal tissues and promotes colorectal adenoma growth by suppressing tumor-infiltrating CD8⁺ T cell cytotoxicity and TAM phagocytosis of tumor cells via PD-1.

Fig. 5. Inhibition of the COX-2-PGE2 pathway promotes intestinal CD8⁺ T cell activation and macrophage phagocytosis.

Apc^Min/+ mice were treated with vehicle, celecoxib, or Ono-AE3–208 as described in the Methods. (A) The percentage of proliferated CD8⁺ T cells in small intestinal (SI) tumors (T) and matched normal tissues (N) were analyzed by Flow Cytometry. (B) The percentage of INFγ-positive CD8⁺ T cells in total CD8⁺ T cells in SI tumors and matched normal tissues were analyzed by Flow Cytometry. (C) The percentage of proliferated CD8⁺ T cells in small intestinal tumors and matched normal tissues were analyzed by Flow Cytometry. (D) The percentage of INFγ-positive CD8⁺ T cells in total CD8⁺ T cells in SI tumors and matched normal tissues were analyzed by Flow Cytometry. (E-F) After macrophages isolated from SI tumors and matched normal tissues were co-cultured with GFP-labeled MC26 cells, the phagocytic index is determined as described in Fig. 4. (G and H). Tumor numbers were counted, and size was measured. The error bar indicates ± SEM. *p<0.05.

PD-1 expression correlates well with COX-2 levels, and high levels of PD-1 and COX-2 are associated with poor survival in CRC patients

To further assess whether our preclinical results may have clinical relevance in humans, we first examined the correlation between mRNA levels of PDCD1 (PD-1) and PTGS2 (COX-2) in a human CRC database. We found a positive correlation between COX-2 and PD-1 mRNA levels in two TCGA and GEO databases (Fig. 6A–B). An evaluation of the GEO database reveals that CRC patients with high levels of both COX-2 and PD-1 in tumor tissues experienced a much poorer overall survival than those with low levels of both COX-2 and PD-1 (Fig. 6C). In the TCGA database, CRC patients with high levels of both COX-2 and PD-1 in tumor tissues tended to have poorer survival compared to patients with low levels of both COX-2 and PD-1. However, the difference does not reach statistical significance (p=0.068) (Fig. 6D).

Fig. 6. Correlations and survival between COX-2 and PD-1 in CRC patients.

(A-B) Pearson correlations between COX-2 and PD-1 mRNA expression in the TCGA Colorectal Adenocarcinoma Provisional dataset (n=381) (A) and the GSE17537 and GSE17538 datasets of the GEO database (n=434) (B). (C-D) Kaplan-Meier survival curves of the GSE17537 and GSE17538 datasets of GEO database (C) and the TCGA Colorectal Adenocarcinoma Provisional dataset (D).

DISCUSSION

Some of the roles of PGE₂ in regulating immunity and host defense against viral, fungal, and bacterial pathogens have previously been reported (36). For example, one study showed that PGE₂ suppressed CD8⁺ T cell survival and function during a chronic lymphocytic choriomeningitis virus infection (37). It is not clear whether PGE₂ induces tumor immune invasion by inhibiting CD8⁺ T cell proliferation and cytotoxicity in vivo. This report provides the first in vivo evidence demonstrating that inhibition of the COX-2-PGE₂-EP4 pathway increases intestinal CD8⁺ T cell abundance and activation accompanied by a reduction of PD-1 and tumor burden. Moreover, two in vivo studies suggest that PGE₂ might promote intestinal adenoma formation and growth via its effects on macrophages. One study revealed that treatment with celecoxib resulted in a reduction of polyp burden accompanied by conversion of TAMs from the M2 to M1 in the Apc^Min/+ mice (38). Another study reported that deletion of EP4 in myeloid cells resulted in reducing adenoma burden in the Apc^Min/+ mice (39). However, the mechanisms by which PGE₂ promotes colorectal adenoma formation and growth via macrophages were not fully understood or examined. Here our results demonstrate for the first time that inhibition of the COX-2-PGE₂-EP4 pathway increases intestinal macrophage phagocytosis of tumor cells accompanied by reduced levels of PD-1 in intestinal macrophages as well as decreased tumor burden. The most striking of our findings is that inhibition of the COX-2-PGE₂-Ep4 pathway promotes intestinal CD8⁺ T cell cytotoxicity and macrophage phagocytosis of epithelial cells via PD-1, which occurs well before tumor formation. This finding provides a potential explanation for the chemopreventive effect of NSAIDs on CRC that occurs very early during a precancer stage.

Following immune activation or infection, PD-1 expression is elevated in effector T cells, macrophages, DCs, B cells, and NKT cells. PD-1 expression is induced in CD8⁺ T cells following TCR stimulation via NF-AT (40). However, TCR-dependent PD-1 induction is weak and transient. It has been well documented that a strong and persistent expression of PD-1 is observed during chronic viral infection (41). Several cytokines such as INFα, IL-6, and IL-12 have enhanced TCR-induced PD-1 expression in CD8⁺ T cells (42,43). In addition to CD8⁺ T cells, PD-1 expression can be regulated by cytokines and TLR ligands such as LPS in macrophages. For example, LPS, TNFα, INFα, and IL-1β induced PD-1 expression in macrophages in vitro (27,44,45). However, how PD-1 is regulated in CD8⁺ T cells and macrophages in the tumor microenvironment is still largely unknown. Previous studies have shown that PD-1 expression in CD8⁺ T cells can be upregulated by different transcription factors such as FoxO1, IRF9, Notch, and c-Fos (42,46–48), and we found that activation of NF-κB by PGE₂ via an EP4-PI3K-AKT pathway initiates PD-1 transcription via binding to the PD-1 promoter in CD8⁺ T cells. Our finding extends the scope of our understanding of the underlying mechanisms for PD-1 regulation in CD8⁺ T cells and macrophages. Moreover, our results also reveal that PGE₂ leads to AKT activation in both CD8⁺ T cells and macrophages. Although AKT has been shown to activate NF-κB by phosphorylation of IKKα (49), further research is needed to determine the molecular mechanisms by which PGE₂-induced AKT activation induces NF-κB activation in CD8⁺ T cells and macrophages. In addition, our results also reveal that PGE₂ only induces PD-1, but not CTLA4, in CD8⁺ T cells. This finding indicates that combined inhibition of PGE₂ signaling and CTLA4 may have more anti-tumor efficacy than combined inhibition of PGE₂ signaling and PD-1.

Previous in vitro studies showed that PGE₂ regulates CD8⁺ T cell proliferation and function. For example, treatment of CD8⁺ T cells with PGE₂ resulted in suppression of cell proliferation by inducing replicative senescence (50). PGE₂ treatment also inhibited CD8⁺ T cell cytotoxicity by inducing CD94 and the NKG2A C-type lectin receptor complex (51) or by inhibiting IFNγ release induced by TCR (52). Moreover, PGE₂ produced by carcinoma cells also blocked initial priming of naïve CD8⁺ T cells to cytotoxic T cells by tumor cells (53). Here we reveal a novel mechanism by which PGE₂ inhibits CD8⁺ T cell proliferation and cytotoxicity against tumor cells by inducing PD-1 via an EP4-PI3K-AKT-NF-κB pathway. In addition, in vitro studies have shown that PGE₂ promotes M2 macrophage polarization via a CREB/CRTC pathway in bone marrow-derived macrophages (54) and the COX-2-PGE₂ pathway mediated the effect of bladder tumor cells on induction of PD-L1 in bone marrow-derived macrophages and MDSCs in a co-culture system (55). These PD-L1 positive cells were immunosuppressive (55). Moreover, PGE₂ also inhibited alveolar macrophage phagocytosis against bacteria (56). However, the mechanisms underlying PGE₂ regulation of macrophage phagocytosis have not been determined. This study found that PGE₂ suppresses macrophage phagocytosis of tumor cells by induction of PD-1 via an EP4-PI3K-Akt-NF-κB pathway.

PD-1⁺ tumor-infiltrating lymphocytes are associated with poor prognosis in human breast cancer (57) and soft tissue sarcomas (58) and correlate with poor survival in patients with high-grade upper tract urothelial carcinomas (59) and gastric cancer (60). PD-1⁺ tumor-infiltrating immune cells are also associated with advanced tumor-node-metastasis stage and poor survival in patients with renal cell carcinoma (RCC) (61) and operable breast cancer (62). Consistent with these reports, our results showed that COX-2 levels are positively associated with PD-1 in human CRC specimens. A recent study revealed that PGE₂ levels are also associated with PD-1 levels in tumor-infiltrating CD8⁺ T cells in human lung cancer specimens (63), consistent with our results. Moreover, our results reveal that high levels of both COX-2 and PD-1 are associated with poorer overall survival in CRC patients. Recent studies showed that combined inhibition of PGE₂ signaling and PD-1 increased CTL proliferation in vitro (64), and EP4 antagonists enhanced antitumor efficacy of PD-1 in a syngeneic mouse model of CRC (65) and in a mouse model of colitis-associated tumorigenesis (66).

In summary, we report that PGE₂ promotes tumor immune evasion by suppressing CD8⁺ T cell cytotoxicity and macrophage phagocytosis against tumor cells by induction of PD-1 via an EP4-PI3K-AKT-NF-κB pathway. In contrast, inhibition of the COX-2-PGE₂-EP4 pathway suppresses tumor immune evasion by increasing CD8⁺ T cell abundance as well as cytotoxicity and enhances macrophage phagocytosis of tumor cells. These findings uncover a novel role of PGE₂ in tumor immune evasion and may provide a rationale for the development of new therapeutic approaches to subvert tumor immune evasion by targeting immune checkpoint pathways using EP4 antagonists. Given that EP4 antagonists have similar efficacy as celecoxib in the prevention and treatment of colorectal adenomas and CRC, the use of these agents will likely avoid the serious cardiovascular side effects associated with NSAIDs and COXIB use.

Prevention Relevance Statement.

These findings provide a potential explanation underlying the chemopreventive effect of NSAIDs on reducing CRC incidence during premalignancy and provide a rationale for developing EP4 antagonists for CRC prevention and treatment. Simply targeting PGE₂ signaling alone may be efficacious in CRC prevention and treatment, avoiding side effects associated with NSAIDs.