The presence of mast cells in some human colorectal cancers is a positive prognostic factor, but the basis for this association is incompletely understood. Here, we found that mice with a heterozygous mutation in the adenomatous polyposis coli gene (ApcMin/+) displayed reduced intestinal tumor burdens and increased survival in a chemokine decoy receptor, ACKR2-null background, which led to discovery of a critical role for mast cells in tumor defense. ACKR2–/–ApcMin/+ tumors showed increased infiltration of mast cells, their survival advantage was lost in mast cell–deficient ACKR2–/–SA–/–ApcMin/+ mice as the tumors grew rapidly, and adoptive transfer of mast cells restored control of tumor growth. Mast cells from ACKR2–/– mice showed elevated CCR2 and CCR5 expression and were also efficient in antigen presentation and activation of CD8+ T cells. Mast cell–derived leukotriene B4 (LTB4) was found to be required for CD8+ T lymphocyte recruitment, as mice lacking the LTB4 receptor (ACKR2–/–BLT1–/–ApcMin/+) were highly susceptible to intestinal tumor-induced mortality. Taken together, these data demonstrate that chemokine-mediated recruitment of mast cells is essential for initiating LTB4/BLT1-regulated CD8+ T-cell homing and generation of effective antitumor immunity against intestinal tumors. We speculate that the pathway reported here underlies the positive prognostic significance of mast cells in selected human tumors. Cancer Immunol Res; 6(3); 332–47. ©2018 AACR.
Mast cells are critical mediators of allergic responses, pathogen recognition, and modulation of immune responses at mucosal sites (1). The presence of mast cells in tumors has been correlated with either good or bad prognosis in several studies of rodents and humans (2), highlighting the complexity of effects mediated by mast cells. Tumor-associated mast cells promote a variety of cancers by enhancing inflammation and angiogenesis (3, 4). However, in human colon cancers, mast cell presence is correlated with better prognosis (5–7), but the mechanisms remain unknown. Although mast cells can exert their antitumorigenic functions by direct interactions with tumor cells, the majority of their effector functions rely on releasing a wide variety of mediators and modulating the immune response. Cytokines and chemokines produced by mast cells are known to have opposing roles in cancer progression by recruiting immune cells to tumor environment (8). Mast cells also rapidly synthesize a wide range of lipid mediators, such as leukotriene B4 (LTB4), leukotriene C4 (LTC4), and prostaglandin E2 (PGE2), that regulate a range of pathophysiological activities, including leukocyte recruitment and smooth muscle contraction (9).
Chronic inflammation promotes a variety of cancers, but inflammation can also play an important role in tumor surveillance (10, 11). Atypical chemokine receptor 2 (Ackr2, hereafter referred to as ACKR2, formerly known as D6) is a “decoy” chemokine receptor that binds and internalizes a large cadre of inflammatory chemokines targeting them for degradation, and, thus, limiting the inflammatory response (12). The importance of ACKR2-mediated chemokine scavenging in the resolution of inflammation became evident in different animal models such as chemical-induced cutaneous inflammation (13, 14), ovalbumin- or Mycobacterium-induced lung inflammation (15, 16), lipopolysaccharide (LPS)-induced placental inflammation (17), or methylcholanthrene/phorbol ester (TPA)-induced skin cancer (18), as well as azoxymethane (AOM)-induced dextran sodium sulfate (DSS)-promoted (AOM/DSS) colon tumor models (19).
In this study, ACKR2–/– mice were bred with ApcMin/+ mice, which develop spontaneous intestinal adenomas, in order to understand the link between chronic inflammation and tumor promotion. Contrary to the established role of ACKR2, the ACKR2–/– ApcMin/+ mice showed decreased inflammation coinciding with decreased tumor burden and improved survival. Using genetic, biochemical, and immunologic methods, we identified enhanced mast cell infiltration as the likely cause of reduction in tumor burden in the ACKR2–/– ApcMin/+mice. We demonstrate that LTB4 produced by mast cells mediates CD8+ T-cell recruitment and is essential for generating effective antitumor immunity against intestinal tumor development.
Materials and Methods
ACKR2–/– mice were obtained from Schering-Plough. To generate ACKR2–/–ApcMin/+, SA–/–ApcMin/+, BLT1–/–ApcMin/+, and Rag2–/–ApcMin/+ strains, male C57BL/6-ApcMin/+ mice (The Jackson Laboratory) were crossed individually with ACKR2–/–, SA–/– (Sash mice/C57BL/6J-KitW-sh/W-sh; The Jackson Laboratory), BLT1–/– (20), and Rag2–/– (Taconic Biosciences) mice, respectively. The following compound mouse strains were generated by breeding male ACKR2–/– ApcMin/+ mice individually with female KitW-sh/W-sh, BLT1–/–, or Rag2–/– mice, respectively: ACKR2–/– SA–/– ApcMin/+, ACKR2–/– BLT1–/– ApcMin/+, and ACKR2–/– Rag2–/– ApcMin/+. Animals were housed under specific pathogen-free conditions, and all mice carrying the ApcMin/+ allele were maintained on breeder chow. The genotyping PCR for ApcMin/+ and Rag2 was performed according to Jackson Laboratories protocols and for the detection of ACKR2 and BLT1 using previously described methods (refs. 13, 20; Supplementary Table S1). All the experimental protocols were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Louisville.
Survival of mice
The overall survival of the mice (ApcMin/+, ACKR2+/–ApcMin/+, ACKR2–/–ApcMin/+, ACKR2–/–SA–/–ApcMin/+, and ACKR2–/–BLT1–/–ApcMin/+) was followed from birth to the endpoint of moribund or death. Significance of differences in survival was determined by the Mantel–Haenszel/log-rank test.
Measurement of hematocrit
Animals were euthanized by CO2 asphyxiation, and 200 μL of blood was drawn from the heart and put into heparin-coated microvette (Sarstedt) tubes followed by analysis using the Hemavet-950 Hematology System (Drew Scientific).
Analysis of adenomas
Mice were sacrificed at 40 days or 110 days of age, and the entire intestinal tract was removed and flushed with 1× PBS using a blunt-end syringe to remove fecal material. Flushed small intestines were longitudinally opened and spread on filter paper using forceps, immobilized by placing another filter paper on the top, and fixed in 10% neutral formalin for 16 hours followed by transferring into 70% ethanol Polyps in proximal, middle, distal small intestines, and colon were counted by experienced lab personnel under a stereo microscope and classified by size: 0–1 mm, 1–2 mm, 2–3 mm, and >3 mm. The mean number of tumors/mouse ± SEM and the mean tumor diameter (mm) in the group ±SEM were calculated for the small intestine and colon, separately.
Cleaned small intestines were divided into 3 equal length sections (proximal, middle, and distal) and fixed with 10% neutral formalin for 16 hours followed by storage in 70% ethanol until embedded in paraffin. The sections were cut at 5-μm thickness and used for hematoxylin and eosin staining (H&E) or immunohistochemical analysis. All images were scanned with an Aperio digital scanner (Leica Biosytems).
For mast cell IHC, formalin-fixed, paraffin-embedded (FFPE) sections were heated at 90°C in a 10 mmol/L Tris, 1 mmol/L EDTA (pH 9.0) solution followed by incubation for 1 hour at room temperature with rat monoclonal anti-mouse Mcpt-1 (clone RF6; eBioscience) 1/200 dilution (2.5 ng/μL final concentration) using standard histochemical protocols and Vectastain ABC-AP staining kit (Vector Laboratories). To measure tumor cell proliferation, mice were given 5′-bromo-2′-deoxyuridine (BrdUrd; 1 mg/mL) by intraperitoneal route and euthanized after 2 hours, followed by intestinal processing and paraffin embedding. IHC for BrdUrd was performed using a BrdUrd in situ detection kit from BD Pharmigen, according to the manufacturer's instructions. Cells containing BrdUrd-positive nuclei (brown nuclei) were manually counted by experienced lab personnel per 200× field. Apoptotic cells within tumors were determined by terminal deoxynucleotidyl transferase-mediated nick end-labeling (TUNEL) using the in situ Cell Death Detection Kit, Fluorescein (Roche). Briefly, FFPE tissue sections were dewaxed in xylene and alcohol. Epitope retrieval was performed by heating to 95°C to 100°C for 30 minutes in sodium citrate buffer (pH 6.0) and exposed to the TUNEL reaction mixture, according to the manufacturer's instructions. The TUNEL-positive, green-fluorescent cells were captured using a Nikon Eclipse TE300 fluorescence microscope and manually counted per field at 200× magnification.
Distal intestine tissue was filled with OCT compound, Swiss rolled, mounted with OCT, and rapidly frozen by immersing in 2-methylbutane (Sigma) chilled in dry ice. The OCT frozen tissue was cut into 6- to 7-μm size sections and fixed in cold acetone (−20°C) for 10 minutes. After air drying for one hour, sections were washed with 1× PBS and blocked for 30 minutes at room temperature with 5% goat serum, followed by incubation with primary rat anti-mouse CD8α (clone 53-6.7; BD Biosciences), anti-mouse Mcpt-1 (clone RF6, eBioscience), or isotype control antibodies (at a 1/50 dilution) for 60 minutes at room temperature in a humidified chamber. Slides were then washed with 1× Phosphate-buffered saline, followed by addition of goat anti-rat IgG–Alexa Fluor 488 at a 1/500 dilution and incubated for 30 minutes. Sections were counterstained with DAPI-1, mounted using Vectashield mounting medium (Vector Laboratories) and visualized on Nikon A1R confocal microscope using appropriate filters. The number of tumor-infiltrating CD8+ T cells from at least 6 tumors in each genotype was counted, calculated per 100 mm2 area, and plotted.
Bone marrow–derived mast cells (BMMC; 1 × 105) isolated, as described in a later section, were washed and spread on superfrost slides in 70 μL volume using a Shandon cytospin centrifuge. Cytospin preparations of BMMCs were stained indirectly using rat anti-CD117 (clone, 2B8; Santa Cruz Biotechnology) and biotin-labeled anti-FcϵR1 (clone MAR-1; eBioscience) antibodies and visualized using anti-rat Green Alexa-488–labeled antibody (Invitrogen) and Red Alexa-647 Streptavidin (Jackson Immuno Research Laboratories). The immunofluorescence images were captured using a Nikon A1R confocal microscope using appropriate filters.
RNA Extraction, real-time PCR, and microarrays
The distal small intestine tissue from WT (C57BL/6) and ACKR2–/– mice, as well as size-matched intestinal tumors of ACKR2–/–ApcMin/+ and ApcMin/+ mice (110 days age old) were dissected and frozen immediately in TRIzol reagent (Invitrogen) at −80°C. After quick thawing, tissue was homogenized using an Omni GLH general homogenizer, RNA was extracted using the RNA mini prep kit from Qiagen, followed by DNase treatment (Ambion Inc and stored at −80 °C until further usage.
RNA was extracted from BMMCs, macrophages, T cells, B cells, and dendritic cells isolated as described in later sections and frozen (−80°C) in TRIzol in a similar manner as above without homogenization. For BMMC microarray analysis, total RNA obtained from three cultures derived from independent mice for each genotype were analyzed on Bioanalyzer (Agillant-2100) and the integrity verified (RIN of 7.5 or above). The three samples for each genotype were pooled and used immediately for microarray analysis.
BMMCs microarray analysis was performed using the Affymetrix GeneChip Mouse Gene 1.0 ST Array, according to manufacturer's protocol at the microarray core facility, University of Louisville. The GeneChip-brand array is comprised of over 750,000 unique 25-mer oligonucleotide features constituting over 28,000 gene level probes with an average of 27 probes per gene. Briefly, total RNA was amplified and labeled following the Affymetrix standard protocol for whole transcript expression analysis and then hybridized to the Affymetrix Mouse Gene 1.0 ST arrays. The arrays were processed following the manufacturer-recommended wash and stain protocol on an Affymetrix FS-450 fluidics station and scanned on an Affymetrix GeneChip 7G scanner using the Command Console 3.1. The resulting CEL files were imported into Partek Genomics Suite 6.6, and transcripts were normalized on a gene level using RMA as the normalization and background correction method. One-way ANOVAs were set up to compare the genotypes (WT vs. ACKR2−/−) of interest. Pathway analysis was carried out by uploading data files into Ingenuity Pathway Analysis (IPA) software.
The microarray for tumors was carried out using the whole mouse genome chip (Mouse 430 2.0 array; Affymetrix). Total RNA obtained from three each of the following independent mice was pooled. WT small intestine tissue; small intestine tumors from ApcMin/+; and ACKR2−/−ApcMin/+ mice. Experimental and sample preparation variations were standardized by applying the global scaling procedure to all absolute analysis data using constant global target intensity. The data were analyzed using Affymetrix's MAS 5 algorithm for probe set summarization, followed by pairwise comparison.
For all semiquantitative real-time PCRs (qPCR), cDNA was synthesized from 1 μg of total RNA using random hexamer primers and Taqman reverse Transcription reagents (N8080234, Applied Biosystems), followed by qPCR reaction with SYBR-green master mix using PCR primers (Realtime Primers LLC; listed in Supplementary Table S2). Real-time qPCR was performed in a Bio-Rad CFX96TM real-time PCR thermocycler. Data were normalized to β-actin, and the relative expression of target genes was calculated using the 2−ΔΔCT method.
Western blot analysis of Mcpt-1
Size-matched intestinal tumors were dissected, snap-frozen in liquid nitrogen, and stored at −80°C. Tumors and normal distal intestine tissues were homogenized in Bio-Rad lysis buffer containing 1× protease inhibitor cocktail (Roche) using a hand-held homogenizer (Omni International). Homogenates were incubated for 1 hour at 4°C with constant agitation, followed by centrifugation. Total cell extracts (50 μg of total protein as determined by Pierce BCA protein assay kit, Thermo Fisher Scientific) were separated on 4% to 12% NuPAGE Bis-Tris precast gels and transferred onto nitrocellulose (Bio-Rad) membranes. Membrane blots were probed with rat anti-mouse Mcpt-1 (1 μg/mL; clone 285008; R&D Systems), followed by peroxidase-coupled secondary anti-rat IgG HRP and detected with Pierce ECL substrate. The same blot was stripped with stripping buffer (Thermo Scientific) and reprobed with β-actin-HRP antibody (Santa Cruz Biotechnology).
BMMCs were generated from 4- to 6-week old female mice. Bone marrow was flushed with RPMI from tibia and femur bones, gently mushed (pulped), filtered through 60-μm nylon mesh, and centrifuged at 450× g for 5 minutes. The pellet was resuspended in BMMC culture medium [DMEM containing 10% FCS, penicillin (100 units/mL), streptomycin (100 μg/mL), 2 mmol/L l-glutamine, and 50 μmol/L β-mercaptoethanol] supplemented with recombinant mouse SCF (12.5 ng/mL; R&D Systems, #455-MC) and recombinant mouse IL3 (10 ng/mL; R&D Systems, #403-ML). Cells were plated at a density of 1 × 106 cells/mL in a T-75 cm2 flask. Nonadherent cells were transferred after 48 hours into fresh flasks without disturbing the adherent (fibroblast) cells. Flasks were changed weekly or as needed to separate the nonadherent mast cells from the contaminating adherent cells with BMMC culture medium described above. Mast cells were visible after 4 weeks of culture and propagated further by dividing once or twice a week to a density of 0.5 × 106 to 1 × 106 cells/mL. For some experiments, mast cells were purified from non-homogenous cultures after 2 weeks on a MACS MS column (Miltenyi Biotec) using CD117 antibody positive selection, according to the manufacturer's protocol. BMMCs were confirmed by double-positive staining for mast cell–specific markers c-Kit and FcϵR1 (clones 2B8 and MAR-1, respectively; eBioscience) by FACS analysis and used at least at 95% purity.
Chemotaxis of BMMCs (from WT and ACKR2–/– mice) was evaluated using 5-μm pore size Transwell filters (Corning Costar). Briefly, cells were resuspended at a density of 5.0 × 106 cells/mL in chemotaxis buffer (RPMI 1640 and 0.5% BSA). The lower chamber was loaded with various concentrations of SCF (R&D Systems), CCL2, and/or CCL5 (PeproTech Inc.) in a volume of 600 μL, and 0.5 × 106 BMMCs were added to the upper chamber in 100 μL BMMC medium. After 3 hours at 37°C in 5% CO2, the upper chamber was removed, and the cells in the lower chamber were fixed in buffered 2% formaldehyde. The migrated cells were counted using a FACS Canto (BD Biosciences) by running for fixed time (fixed volume of cells) and analyzed using buffer only as a background control.
BMMCs (4 × 106/2 mL) were loaded with 1 μmol/L INDO-1 (Molecular Probes) and the calcium response after stimulation with different concentrations of CCL2, CCL5, and SCF as indicated and measured using spectrofluorimeter.
Preparation of immune cells
CD8+ T cells were isolated from the spleens and mesenteric lymph nodes of OT-1 mice (The Jackson Laboratory). Briefly, spleen and lymph nodes were crushed and passed through 70-μm strainers (Corning), and the red blood cells were lysed with RBC lysis buffer (BioLegend). After lysis, CD8+ T cells were selected using CD8α (Ly-2) microbeads (Miltenyi) on a MACS LS column, according to the manufacturer's protocol. The purified CD8+ T cells were >95% positive for CD8 as analyzed by FACS. These cells were used for mast cell and T-cell coculture studies.
For ACKR2–/–Rag2–/–ApcMin/+ adoptive transfer experiments (described in following sections), spleen cell suspensions from ACKR2–/–ApcMin/+ mice were positively enriched for CD8+ T cells using CD8α (Ly-2) microbeads, followed by purification on MACS LS columns. The CD8+ T cell–negative fraction was used subsequently to isolate CD4+ T cells using CD4 (L3T4) microbeads.
For CCR2 and CCR5 RNA expression analysis, T cells and B cells were isolated from spleens and mesenteric lymph nodes of WT and ACKR2–/– mice using a Pan T-cell isolation kit (#130-090-861) from Miltenyi and mouse B220 (clone RA3-6B2, BD Biosciences) positive selection on MACS MS columns.
Bone marrow–derived macrophages (BMDM) were generated by isolating cells from femurs and tibia of C57BL/6 and ACKR2–/– mice and culturing in DMEM containing 10% FCS, penicillin (100 units/mL), streptomycin (100 μg/mL), 2 mmol/L l-glutamine, and 50 μmol/L β-mercaptoethanol) supplemented with recombinant M-CSF (100 ng/mL; catalog #576404, BioLegend) for 7 to 8 days. Bone marrow–derived dendritic cells (BMDC) were generated by culturing cells from bone marrow in RPMI containing 10% FCS, penicillin (100 units/mL), streptomycin (100 μg/mL), 2 mmol/L l-glutamine, and 50 μmol/L β-mercaptoethanol in the presence of GM-CSF (40 ng/mL; catalog #415-ML, R&D Systems) and human Flt-3 (100 ng/mL; catalog #308-FKN, R&D Systems) for 8 to 9 days. The purity of the immune cell population was determined by flow cytometry using following antibodies: CD117 (clone 2B8, eBioscience), FcϵR1(clone MAR-1, eBioscience), MHCII (clone AMS-32.1, BD Biosciences), CCR7 (clone 4B12, BioLegend), CD86 (clone GL-1 BD Biosciences), CD11b (clone M1/70, BD Biosciences), CD11c (clone HL-3, BD Biosciences), F4/80 (clone BM8, eBioscience), CD4 (clone GK1.5, eBioscience) and CD8 (clone 53-6.7), CD19 (clone MB-19-1, eBioscience), CD3 (clone 145-2C11, BD Biosciences), GR-1 (clone RB6-8C5 eBioscience, and CD45 (clone 30-F11, eBioscience).
The CFSE (Invitrogen) stock was freshly prepared at 10 μmol/L in DMEM. The purified OT-1 CD8+ T cells were washed with DMEM and labeled with 2.5 μmol/L CFSE for 10 minutes at 37°C in the dark. The reaction was stopped quickly with DMEM containing 2% FCS, and cells were then centrifuged, washed, and resuspended in 2 mL DMEM containing 10% FBS. The cells were counted and diluted to 1 × 106 cells/mL for plating.
BMMC and T-cell cocultures
The stock of OVA octapeptide SIINFEKL (ovalbumin 257–264; Sigma) was prepared in 1× PBS at a 0.5 mg/mL concentration and stored at −20°C. WT or ACKR2–/– BMMCs (2 × 106) were incubated with SIINFEKL for 3 hours in DMEM containing 10% FBS. Following incubation, BMMCs were washed 3 times and resuspended in DMEM containing 10% FBS. For coculture experiments, 1 × 106 CD8+ T cells or CFSE-labeled CD8+ T cells were mixed at different ratios with peptide-loaded BMMCs and cultured for 48 or 72 hours to measure activation or proliferation, respectively. CD8+ T cell activation was measured by increased surface expression of CD69 (clone H1-2F3; eBioscience), CD44 (clone IM-7), CD25 (clone PC61, BioLegend) by FACS. The CD8+ T cell proliferation was determined by FACS, following the CFSE dilution of CFSE-labeled CD8+ T cells.
Cells (1 to 5 × 105 cells) were resuspended in 50 μL FACS buffer (1× PBS containing 1% BSA), and FcR blocked (15 minutes at 4°C) using rat anti-mouse CD16/CD32 (clone 2.4G2; BD Biosciences) antibody. Cells were then incubated on ice in the dark with cell-specific fluorochrome-conjugated antibodies listed in other sections for 40-minute labeling. Cells were washed with cold 1× PBS, centrifuged at 400× g for 3 minutes, and fixed with 2% formaldehyde (Fisher Scientific). Cells were acquired using a FACS Caliber or FACS Canto (BD Biosciences) and analyzed using FlowJo software v7.6.5). The cell viability was measured using 0.5 μL of fixable viability Dye eFluor 660 (eBioscience) in 500 μL 1× PBS.
Intracellular IFNγ staining
T cells (1 × 106) were stimulated in 12-well plates for 5 hours with 1× cell stimulation cocktail containing phorbol 12-myristate 13-acetate (PMA) and ionomycin (eBioscience) and then washed, fixed in 1× buffer (# 00-8333, eBioscience), permeabilized (# 00-8222, eBioscience), and stained with rat anti-mouse IFNγ (clone XMG 1.2; BioLegend) according to eBioscience intracellular staining instructions.
Adoptive transfer of mast cells and CD8+ T cells
BMMCs (1 × 107 in PBS) derived from ACKR2–/– mice were given intravenously (i.v.) to 30-day-old ACKR2–/–SA–/–ApcMin/+ mice. Recipients and controls were analyzed for anemia and tumor development at 100 days age as described in other sections. ACKR2–/–Rag2–/–ApcMin/+ mice at 35 days of age were given 8.5 × 105 CD8+ T cells isolated from the spleens of ACKR2–/–ApcMin/+ mice or the same number of combined CD4+ and CD8+ T cells (1:1) in PBS.
Leukotriene B4 measurement
For measuring levels of LTB4, distal intestine tumors from ACKR2–/–ApcMin/+, ACKR2–/–SA–/–ApcMin/+, and ACKR2–/–BLT1–/–ApcMin/+ were resected into 500 μL of cold indomethacin (10 μmol/L) containing 1× PBS and homogenized immediately using a Omni GLH general homogenizer. The homogenates were centrifuged at 14,000× g for 10 minutes, and the LTB4 levels in the supernatant were quantified using a LTB4 EIA kit (Cayman Chemical). The LTB4 levels were normalized to the amount of protein in the homogenates (measured by BCA protein assay, Thermo Fisher Scientific).
Standard Mann–Whitney U nonparametric, two-tailed test and two-tailed Student t test were used for comparisons between two experimental groups, with a P value of <0.05 considered significant using Graph Pad Prism software (***, P < 0.001; **, P < 0.01; *, P < 0.05). The significance of survival of animals was calculated using Kaplan–Meier survival curves by the Mantel–Haenszel/log-rank test.
Decreased mortality and tumorigenesis in ACKR2–/–ApcMin+ mice
In AOM/DSS-induced colon tumor models, genetic deficiency of ACKR2 accelerates tumor growth by enhancing chemokine-mediated inflammation (19). In marked contrast, we found that ACKR2–/–ApcMin/+ mice displayed a significant decrease in morbidity and mortality relative to ApcMin/+ mice (Fig. 1A). The average life expectancy of ACKR2–/–ApcMin/+ at 208 days was a third more than the 154 days observed for ApcMin/+ mice. Heterozygous ACKR2+/–ApcMin/+ mice also showed improved survival with some mice approaching 300 days of age, whereas some of the ACKR2–/–ApcMin/+ mice were alive as long as 500 days (when the experiment was terminated). Intestinal adenomas cause severe anemia in ApcMin/+ mice, as seen here by their significant reduction in hematocrit values compared with WT mice at 110 days age (Fig. 1B). Consistent with their improved survival, ACKR2–/–ApcMin/+ displayed significantly increased hematocrit values compared with ApcMin/+ mice, indicating slower progression of the disease. No difference in hematocrit values was observed between the WT mice and the ACKR2–/– mice.
Tumor burden in the small intestine and colon was analyzed by measuring tumor number and size in longitudinally opened intestinal sections. In the small intestine, tumor number and size were significantly reduced in ACKR2–/–ApcMin/+ mice (Fig. 1C–F), indicating slower disease progression compared with ApcMin/+. The frequency of polyps was similarly reduced in proximal, middle and distal regions of ACKR2–/–ApcMin/+ small intestines (Fig. 1D). The gross appearance of longitudinally opened distal small intestines (Fig. 1F) provides an image that clearly illustrates the decrease in polyp numbers in ACKR2–/–ApcMin/+ mice. In contrast, the number of tumors and the size frequency of tumors found in the colons did not show significant differences between ApcMin/+ and ACKR2–/–ApcMin/+ mice (Supplementary Fig. S1). Analysis of 40-day-old mice demonstrated a significant decrease in hematocrits only in ApcMin/+ mice but not in the ACKR2–/–ApcMin/+ mice (Supplementary Fig. S2A). ACKR2–/–ApcMin/+ mice also showed significantly reduced intestinal polyps compared with ApcMin/+ mice (Supplementary Fig. S2B). Histopathologic analysis of small intestines showed significant microadenomas in ApcMin/+ mice at 40 days of age, whereas minimal disease was noted in ACKR2–/–ApcMin/+ mice (Supplementary Fig. S2C).
In the ApcMin/+ model, inflammation is known to promote intestinal tumor development (21, 22). Consistent with decreased tumor burden, analysis of mRNA from the ACKR–/–ApcMin/+ tumors revealed that the expression of inflammatory mediators COX-2, CXCL1, IL1β, and TNFα decreased in ACKR2–/– ApcMin/+ mice compared with ApcMin/+ mice (Supplementary Fig. S3). Tumor size reduction coincided with lower cell proliferation, as demonstrated by reduced BrdUrd incorporation in ACKR2–/–ApcMin/+ tumors compared with ApcMin/+ tumors (Fig. 1G). However, tumors from ACKR2–/–ApcMin/+ mice contained more apoptotic cells compared with ApcMin/+ mice indicating that immune surveillance might play a role in controlling tumor growth (Fig. 1H).
Increased mast cell infiltration in ACKR2–/–ApcMin/+ tumors
In contrast to the proinflammatory phenotype observed with several ACKR2-null mouse models (13, 17, 23, 24), we found that ACKR2–/–ApcMin/+ mice displayed decreased inflammation and tumor burden. To examine the molecular mechanisims underlying this phenotype, gene expression profiles of ACKR2–/–ApcMin/+ and ApcMin/+ intestinal tumors were compared by microarray analysis. Tumors from ACKR2–/–ApcMin/+ were enriched in RNA transcripts for mast cell proteases (Mcpt) 1 and 2 relative to ApcMin/+ tumors (Fig. 2A). This expression pattern was confirmed by qPCR (Fig. 2B) and Western blot analysis (Fig. 2C), and IHC analysis revealed it was due to a significant increase in mast cell infiltration of tumors in ACKR2–/–ApcMin/+ mice relative to ApcMin/+ mice (Fig. 2D). The pattern appeared to be established early in disease, as small intestinal microadenomas analyzed from 40-day-old ACKR2–/–ApcMin/+ mice were also enriched in mast cells relative to ApcMin/+ mice (Supplementary Fig. S4).
Mast cells attenuate tumor burden in ACKR2–/–ApcMin/+ mice
To determine whether or not mast cells were responsible for protection from intestinal tumorigenesis, ACKR2–/–ApcMin/+ mice were crossed with the mast cell deficient Kitw-sh/w-sh (SA) mice and tested for tumor burden. These ACKR2–/–SA–/–ApcMin/+ mice died with average lifespans of 180 days, close to those of ApcMin/+ mice (Fig. 3A), indicating mast cells were required for the survival advantage of ACKR2–/–ApcMin/+ mice. The hematocrit values of ACKR2–/–SA–/–ApcMin/+ mice were significantly lower than ACKR2–/–ApcMin/+ mice, consistent with faster disease progression in the absence of mast cells (Fig. 3B).
Although the total number of polyps in the small intestines of ACKR2–/–SA–/–ApcMin/+ mice was increased slightly, the increase was not significantly different from ACKR2–/–ApcMin/+ mice (Fig. 3C). However, analysis of the size distribution of polyps in the small intestine revealed a significant increase that was restricted to larger tumors (1–2 mm, 2–3 mm, and 3 mm and above) in ACKR2–/–SA–/–ApcMin/+ relative to ACKR2–/–ApcMin/+ mice, despite comparable total number of tumors in both groups (Fig. 3C and D). H&E staining (Fig. 3E) of small intestinal sections revealed the presence of large tumors in the distal intestine of ACKR2–/–SA–/–ApcMin/+ compared with mast cell sufficient mice. Tumors in the range of 0 to 1 mm were significantly less frequent in ACKR2–/–SA–/–ApcMin/+ mice, which suggests that tumor growth, and not tumor initiation, was affected most by the absence of mast cells. To further test for mast cell contributions to immune surveillance, BMMCs from ACKR2–/– mice were infused intravenously into 4-week-old ACKR2–/–SA–/–ApcMin/+ mice and analyzed for tumor burden at 110 days of age. The hematocrit values of mast cell–transferred ACKR2–/–SA–/–ApcMin/+ mice were significantly higher (Fig. 3F), with concomitant reductions in tumor size and numbers (Fig. 3G and H). Hence, our genetic and cell-transfer experiments indicate mast cells are responsible for slower tumor progression in ACKR2–/– ApcMin/+ mice.
Sustained expression of CCR2 and CCR5 in ACKR2–/– mast cells
To explore the mechanisms behind the increased mast cell abundance in ACKR2–/–ApcMin/+ polyps, we generated mast cells from the bone marrow of WT and ACKR2–/– mice to compare their properties. First, BMMCs positive for mast cell markers c-Kit and FcϵR1 (Supplementary Fig. S5A and S5B) were analyzed for global changes in gene expression patterns using RNA microarray analyses. The expression of chemokine receptors CCR2 and CCR5 was upregulated in ACKR2–/– mast cells (Fig. 4A and B). No significant differences in expression of any other CC or CXC chemokine receptor were observed between the WT and ACKR2–/– mast cells (Supplementary Fig. S6). Because CCR2, CCR5, and other chemokine receptors, such as CXCR2 and BLT1, have been implicated in mast cell homing to tissues (25–27), these results suggest that sustained elevated expression of CCR2 and/or CCR5 on mast cells might contribute to their increased presence in ACKR2–/–ApcMin/+ tumors.
To test for functional differences in migration, ACKR2–/– mast cells were tested in chemotaxis assays with increasing concentrations of CCL2 or CCL5, the ligands for CCR2 and CCR5, respectively. ACKR2–/– mast cells migrated efficiently, showing the typical bell shape response curve (Fig. 4C and D). WT mast cells did not show chemotactic responses to either CCL2 or CCL5, but they migrated efficiently when tested with the well-known mast cell chemotactic ligand, stem cell factor (SCF; Fig. 4E). The signaling functions of CCR2 and CCR5 receptors on ACKR2–/– mast cells were further tested in intracellular calcium flux assays. ACKR2–/– mast cells released intracellular calcium in a dose-dependent response to CCL2 (Fig. 4F) and CCL5 (Fig. 4G). WT mast cells did not respond to either CCL2 or CCL5, even at the highest concentrations tested, 60 nmol/L or 10 nmol/L, respectively. However, both WT and ACKR2–/– mast cells mobilized intracellular calcium equally well in response to 2,4-dinitrophenyl-human serum albumin (DNP-HSA)–mediated IgE-R stimulation, ensuring no inherent defects in the activation of WT BMMCs (Supplementary Fig. S7A and S7B).
It is known that the expression of CCR2 and BLT1 changes during maturation of BMMCs, with high expression in mast cell progenitors that is subsequently downregulated in mature mast cells (26). To examine if the ACKR2–/– mast cells follow a similar pattern, we measured CCR2 and CCR5 mRNA at various time points during the development of mast cells in culture (Fig. 4H and I). Transcripts for CCR2 and CCR5 were abundant early during differentiation in both cell populations and decreased dramatically in WT cells thereafter, as expected. In the ACKR2–/– cell populations, the levels of CCR2 and CCR5 remained high (>500-fold) relative to WT cells after 6 weeks of differentiation in culture. We next asked whether elevated expression of CCR2 and CCR5 in ACKR2–/– mast cells is common to other leukocytes. Analysis of total RNA from purified T cells, B cells, bone marrow–derived macrophages (BMDM), and dendritic cells (DC) from ACKR2–/– mice all showed comparable levels of CCR2 and CCR5 expression (Supplementary Fig. S8). There was a slight increase in CCR2 and CCR5 expression in ACKR2–/– BMDCs and in CCR5 expression in ACKR2–/– B cells compared with WT. However, these changes were relatively weak when compared with changes in mast cells.
Mast cells are critical for CD8+ T cell recruitment in ACKR2–/– ApcMin/+ tumors
Tumor antigen–specific CD8+ T cells play a crucial role in antitumor immunity (28, 29). We hypothesized that the reduced tumor burden in ACKR2–/–ApcMin/+ is due to mast cell–mediated recruitment of CD8+ T cells. Therefore, CD8+ T cells were visualized in the distal intestine tumors from 110-day-old ApcMin/+, ACKR2–/–ApcMin/+, and ACKR2–/–SA–/–ApcMin/+ mice by immunofluorescence. The CD8+ T cell numbers were significantly increased in ACKR2–/–ApcMin/+ tumors compared with ApcMin/+ (Fig. 5A). We further analyzed sections from the same mice for the presence of mast cells by immunofluorescence. The ACKR2–/–ApcMin/+ tumors displayed significantly more mast cellscompared with ApcMin/+, and, as expected, the ACKR2–/–SA–/–ApcMin/+ tumors were devoid of mast cells (Fig. 5B). This absence of mast cells in ACKR2–/–SA–/–ApcMin/+ tumors was associated with significantly reduced tumor infiltration by CD8+ T cells (Fig. 5A), consistent with a critical role for mast cells in their recruitment.
Mast cells are reported to facilitate immune surveillance by presenting tumor antigens to CD8+ T cells and inducing their proliferation and cytotoxic potential (30–32). To test the efficacy of ACKR2–/– mast cells in mediating T-cell activation, coculture experiments were performed using BMMCs and ovalbumin-specific CD8+ T cells from OT-1 TCR transgenic mice. Both WT and ACKR2–/– mast cells were equally efficient in the uptake and presentation of ova peptide (Supplementary Fig. S9), as well as upregulation of the CD8+ T-cell activation markers CD69, CD44, and CD25 (Fig. 5C). ACKR2–/– mast cells induced CD8+ T-cell proliferation and IFNγ production to the same extent as CD8+ T cells activated by WT mast cells (Fig. 5D and E).
CD8+ T cells control tumor progression in ACKR2–/–ApcMin/+ mice
To investigate the role of the adaptive immune system in intestinal tumor progression, the ACKR2–/–ApcMin/+ mice were crossed onto the Rag2–/– background and analyzed for anemia and tumor burden. Hematocrit values in ACKR2–/–Rag2–/–ApcMin/+ mice showed a significant decrease compared with ACKR2–/–ApcMin/+ mice, indicating severe disease (Fig. 6A). Measurement of tumor numbers and size at 110 days of age revealed that although tumor frequencies were similar in ACKR2–/–ApcMin/+ and ACKR2–/–Rag2–/–ApcMin/+ mice, those tumors grew to larger sizes in the absence of a functional adaptive immune system (Fig. 6B and C). To further examine the role of T cells, T cells from ACKR2–/–ApcMin/+ mice were adoptively transferred into ACKR2–/–Rag2–/–ApcMin/+ mice. As shown in Fig. 6D, the hematocrit values of ACKR2–/–Rag2–/–ApcMin/+ mice increased significantly with CD8+ T-cell transfer compared with the mice with sham transfer. There was also a significant decrease in tumor number and size in ACKR2–/–Rag2–/–ApcMin/+ mice upon transfer compared with the sham mice (Fig. 6E and F). The tumor protective effect of transferred CD8+ T cells was clearly visible in the representative images of distal ileum sections (Fig. 6G). Thus, genetic and adoptive transfer experiments suggest that CD8+ T cells are both necessary and sufficient for tumor protection in this model.
To determine if T cells contribute to the enhanced presence of mast cells in ACKR2–/–ApcMin/+ adenomas, cell lysates prepared from the distal intestine tumors of ACKR2–/–Rag2–/–ApcMin/+ mice were analyzed for Mcpt-1 (Supplementary Fig. S10). ACKR2–/–Rag2–/–ApcMin/+ mice tumors showed similar expression of Mcpt-1 as ACKR2–/–ApcMin/+ tumors, indicating that the absence of T cells did not alter mast cell recruitment in ACKR2–/–ApcMin/+ tumors.
LTB4–BLT1 axis mediates CD8+ T-cell recruitment into ACKR2–/–ApcMin/+ tumors
Our findings from an implantable cervical cancer model show that BLT1 expression on CD8+ T cells is critical for their migration to tumors (33). To examine whether mast cell–derived LTB4 is responsible for recruitment of BLT1-expressing CD8+ T cells to ACKR2–/–ApcMin/+ tumors, the ACKR2–/–ApcMin/+ mice were crossed onto the BLT1–/– background. The ACKR2–/–BLT1–/–ApcMin/+ mice displayed accelerated disease and a significant decrease in survival (Fig. 7A and B). The tumor numbers, as well as the size of the tumors, in the higher range increased significantly in ACKR2–/–BLT1–/–ApcMin/+ as compared with ACKR2–/–ApcMin/+ (Fig. 7C and D). Frozen sections of distal intestine tumors from 110-day-old ACKR2–/–BLT1–/–ApcMin/+ mice were analyzed for mast cell and CD8+ T cell recruitment by immunofluorescence. The ACKR2–/–BLT1–/–ApcMin/+ tumors, despite elevated numbers of mast cells, showed significantly reduced CD8+ T cell infiltration compared with ACKR2–/–ApcMin/+ tumors (Fig. 7E and F). Analysis of LTB4 levels in the adenomas showed negligible LTB4 in mast cell–deficient ACKR2–/–SA–/–ApcMin/+ tumors compared with mast cell–sufficient tumors (Fig. 7G). These data together with reduced CD8+ T cell migration to mast cell–deficient ACKR2–/–SA–/–ApcMin/+ tumors strongly suggest that mast cells produced LTB4 to promote effector T-cell recruitment into intestinal adenomas.
The results presented here using ACKR2–/–ApcMin/+ mice in combination with the deletion of mast cells (SA–/–), lymphocytes (Rag2–/–), and LTB4 receptors (BLT1–/–) demonstrate that chemokine-mediated mast cell recruitment initiates LTB4/BLT1-regulated CD8+ T-cell homing. This, in turn, leads to the generation of effective antitumor immunity against intestinal tumor growth that slows progression of fatal disease in this spontaneous tumor model.
In colorectal cancers, an absence of TLR signaling due to MyD88 deletion results in the reduction of adenomas in ApcMin/+ mice, highlighting the critical role of inflammation in tumor development in this model (34). The current study shows that deletion of inflammatory chemokine scavenging receptor ACKR2 in the context of ApcMin/+ mice was protective, as the adenomas in ACKR2–/–ApcMin/+ mice were established more slowly and displayed reduced inflammation and growth in tumor mass. This is in contrast to the well-established role of ACKR2 in limiting inflammation demonstrated in several other models involving ACKR2–/– mice, including colitis-associated colon cancers (18, 19). Thus, the pathway by which oncogenesis is initiated appears to be a critical determinant of the consequences of elevated inflammatory responses in ACKR2–/– mice.
Among the many models that outline the protective role of ACKR2, the psoriasiform inflammation of the skin appears relevant to the current observations (13, 35). In the TPA-induced psoriasis model, the characterization of the immune cell infiltrate in skin sections reveals that ACKR2–/– mice recruit increasing numbers of mast cells during the time course of disease progression, whereas WT mice do not have any such increase in mast cell accumulation (13). The infiltration of macrophages and neutrophils into inflamed skin sections does not differ between WT and ACKR2–/– mice. Interestingly, more CD3-positive T cells were recruited into the treated ACKR2–/– mice skin. Inhibiting either mast cells or T cells reduced the TPA-induced inflammatory pathology in ACKR2–/– mice. As detailed in the current study, the consequence of the similarly elevated mast cell and T-cell recruitment to the site of intestinal tumors in ACKR2−/−ApcMin/+ mice appear to be in generating effective antitumor immunity.
Clearly, the basis for the unexpected tumor protective phenotype appears to be dependent on mast cells because they are significantly increased in ACKR2–/–ApcMin/+ adenomas. As apparent with the Mcpt-1 staining, mucosal mast cell numbers increased significantly at the periphery, as well as in intratumoral regions, of ACKR2–/–ApcMin/+ adenomas. Several studies have used mast cell–deficient strains, such as WBB6F1-Kitw/w-v and C57BL/6-Kitw-sh/w-sh mice, to determine their role in cancer. However, the results obtained are variable, depending on the tumor model used (2). Gounaris and colleagues reported infiltration of mast cells into intestinal polyps in murine models of conditional β-catenin overexpression and truncation of the APC (APCΔ468) gene (36). In lethally irradiated APCΔ468 mice, the polyp number is reduced following the transfer of bone marrow from mast cell–deficient mice compared with WT mice, suggesting a protumor role for mast cells in polyp development. These results are in contrast to the observations of Sinnamon and colleagues in ApcMin/+ mice, where mast cell infiltration in intestinal polyps was shown to negatively correlate with tumor growth (37). A similar protective phenotype for mast cells has been observed in ApcMin/+ tumors lacking the receptor for glycation end products (RAGE), with significant increases in mast cell number correlating with 58% reduced polyp growth (38). The loss of survival advantage of the ACKR2–/–ApcMin/+ mice and the development of large-sized tumors in mast cell–deficient ACKR2–/–SA–/–ApcMin/+ mice reported here suggests a protective role for mast cells. The ability of adoptively transferred ACKR2–/– mast cells to reduce tumor growth further supports the notion that mast cells are protective in intestinal adenoma development. Although the basis for the protumor and antitumor activities observed in these studies remains to be established, a few differences need consideration. In APCΔ468 mice, chymase-producing mucosal-type mast cells are detected in the intraepithelial regions of benign adenomas, and tryptase-producing connective tissue-type mast cells are preferentially enriched in the stroma and invasive regions of adenocarcinomas, indicating the plasticity of mast cell subtypes in a changing tumor environment (39). Although APCΔ468 mice develop invasive intestinal tumors, only benign adenomas develop in ApcMin/+ mice, and they die around the age of 150 days before developing adenocarcinomas. It is possible that mast cells display plasticity and switch between immune enhancing phenotypes and immune suppressive phenotypes, depending on the tumor microenvironment (40–42). Therefore, in this study, we sought to examine the basic mechanisms underlying the protective nature of mucosal mast cells by focusing on their interactions with effector CD8+ T cells.
Mast cells are present in limited numbers in the mouse gut, but their numbers increase during infection and inflammatory conditions (43). Under reduced inflammatory conditions, the intestinal tumors from ACKR2–/–ApcMin/+ mice recruited more mast cells than ACKR2-sufficient ApcMin/+ mice. Phenotypic and functional characterization of BMMCs revealed that ACKR2–/– mast cells have high expression of functional CCR2 and CCR5, whereas WT mast cells did not, providing a likely explanation of how mast cells are selectively recruited into ACKR2–/– tumors. Both CCR2 and CCR5, along with other chemoattractant receptors like CXCR2 and BLT1, are implicated in mast cell homing to the tissues (25–27). However, unlike CCR2 and CCR5, no change in the expression of CXCR2 or BLT1 was detected. There are several potential explanations for why CCR2 and CCR5 are overexpressed or sustained in ACKR2–/– mast cells. Downregulation of CCR2 mRNA has been demonstrated during development of B cells (44), DCs (45), and monocytes (46). CCR2 mRNA is downregulated in BMMCs after 2 weeks of culture, which appears to be a mechanism to confine them into tissues preventing them from emigrating (47). We also detected CCR2 and CCR5 mRNA expression in 2-week-old WT mast cell cultures and observed dramatic downregulation beyond 4 weeks of growth in culture. In contrast, the expression of these transcripts persisted in ACKR2–/– mast cells, suggesting that posttranscriptional processing might be altered in the ACKR2–/– mast cells.
The dramatic upregulation of functional expression of CCR2 and CCR5 observed in cultured mast cells suggests that the enhanced mast cell infiltration observed in ACKR2–/–ApcMin/+ tumors is likely mediated through the activity of either or both of these chemokine receptors. Reciprocal control of ACKR2 and CCR2 in the recruitment of macrophages to developing lymphatic vessel surfaces has been demonstrated (48). It is suggested that in the absence ACKR2-scavenging function on lymphatic endothelium, CCL2/CCR2-mediated macrophage recruitment is enhanced. Likewise, Ly6Chigh monocytes increase in the spleen and blood of ACKR2–/– mice in a CCR2-dependent manner and exhibit enhanced immune suppressive activity (49). During LPS-induced inflammation, increased CCL2 accumulation in the lymph nodes of ACKR2–/– mice recruit high CCR2-expressing CD11b+ Gr1high cells (50). Blocking CCR2 inhibits the migration of other cellular flow and restores the migration of antigen presenting DCs. Graham and colleagues suggest that the draining lymph node area becomes like a clogged sink in the absence ACKR2, preventing the normal flow of leukocytes (51). However, our results showed a significantly different outcome, i.e., a more efficient antitumor immune response in the absence of ACKR2. In the context of generating immunity to the altered self-antigens, this clogged sink is what might be needed for unconventional antigen presentation by antigen presenting cells such as mast cells. Clearly, further work is needed to explore this possibility.
It is clear from the immunofluorescence staining of tumors that mast cell infiltration was correlated with CD8+ T cell recruitment in ACKR2–/– tumors. Although T-cell cytokine-mediated mast cell hyperplasia is known to occur in the gut (52), the abundance of mucosal mast cells in ACKR2–/–Rag2–/–ApcMin/+ tumors clearly suggests that mast cell recruitment in this model is independent of T-cell function. In contrast, we observed enhanced recruitment of CD8+ T cells in mast cell–enriched ACKR2–/–ApcMin/+ tumors and their absence in mast cell–deficient tumors, indicating a critical role for mast cells in T-cell recruitment. Because ACKR2–/–Rag2–/–ApcMin/+ mice displayed large tumors irrespective of mast cell abundance, direct cytotoxicity, or phagocytic capabilities of mast cells were likely not involved in controlling ACKR2–/–ApcMin/+ tumors.
Although the tumor-infiltrating mast cells might mediate positive or negative outcomes for cancer patients, the presence of cytotoxic CD8+ T cells within the core of the tumor or at the invasive margin is usually correlated with good prognosis (53, 54). Analysis by Galon and colleagues shows a clear association between high densities of CTLs and memory CD8+ T cells (CD8+/CD45RO) and longer disease-free survival (28). The development of large-size tumors in ACKR2–/–Rag2–/–ApcMin/+ without a change in the number of tumors clearly shows that T cell–mediated immune responses are critical in controlling tumor growth in these mice. The ability of adoptively transferred CD8+ T cells or CD4+/CD8+ T cells to reinstate tumor protection demonstrates that CD8+ T cell–mediated immune response is essential in ACKR2–/–ApcMin/+ mice. Although further studies are clearly needed to dissect the role and contribution of different CD4+ subsets, including Tregs, the use of SA–/– and Rag2–/– compound mice outlines the importance of mast cell–mediated T-cell responses in ACKR2–/–ApcMin/+ mice.
Mast cell–derived LTB4 is known to mediate the recruitment of effector CD8+ T cells to inflammatory sites via BLT1 (25, 55). Eicosanoids are one of the early mediators produced in the tumor milieu by inflammatory cells such as mast cells, neutrophils, and macrophages. Significantly, higher LTB4 is produced by human colon adenocarcinoma tissue (56), relative to the corresponding normal mucosal tissues. The data on ACKR2–/–BLT1–/–ApcMin/+ mice presented here highlight the importance of the LTB4/BLT1 axis in mediating effector T-cell recruitment into intestinal adenomas in ACKR2–/–ApcMin/+ mice. The ACKR2–/–ApcMin/+ mice devoid of either mast cells or BLT1 develop large intestinal tumors with reduced CD8+ T-cell recruitment. Not only the survival advantage of the ACKR2–/–ApcMin/+ mice was lost in ACKR2–/–BLT1–/–ApcMin/+ mice, but the compound mice retained the accelerated tumor development observed in BLT1–/–ApcMin/+ mice (57). The data showed that mast cells are the cellular origin of LTB4 because elevated production was detected in ACKR2–/–ApcMin/+ adenomas, whereas mast cell–deficient tumors did not produce detectable LTB4. This is consistent with the role of mast cell–produced LTB4 in CD8+ T-cell migration reported in an airway hyperresponsiveness model, where adoptive transfer of WT BMMCs into LTB4-deficient leukotriene A4 hydrolase (LTA4H–/–) mice triggered LTB4 release in BAL fluid and restored AHR response (58).
A model summarizing our findings on the role of mast cell mediated immune surveillance in the ACKR2–/–ApcMin/+ mice is shown in Fig. 8. Elevated recruitment of mast cells resulted in reduced tumor burden and enhanced survival of the ACKR2–/–ApcMin/+ mice relative to ApcMin/+ mice. Accelerated tumor growth in mast cell–deficient ACKR2–/–ApcMin/+ mice points to the importance of these cells in regulating adenoma progression. The lack of immune surveillance in either ACKR2–/–Rag2–/–ApcMin/+ or BLT1–/–ACKR2–/–ApcMin/+ mice, despite mast cell abundance, points to BLT1-mediated CD8+ T cell recruitment as a critical controller of adenoma growth in ApcMin/+ mice. In selected human tumors, colorectal cancers in particular, high mast cell infiltration has been associated with better prognosis. The mechanisms responsible for a tumor-protective function of mast cells and links to the known antitumor function of infiltrating cells have remained elusive (5–7, 53, 54). Based on the results reported here, we surmise that interaction between mast cells and CD8+ T cells with a pivotal role of chemokines and their receptors underlies these clinical observations.
Disclosure of Potential Conflicts of Interest
A. Mantovani is a consultant/advisory board member for Verily, Efranat, AbbVie, Compugen, Novartis, Pierre Fabre, and Roche. No potential conflicts of interest were disclosed by the other authors.
Conception and design: S.R. Bodduluri, E. Krishnan, T.C. Mitchell, V.R. Jala, B. Haribabu
Development of methodology: S.R. Bodduluri, E. Krishnan, V.R. Jala, B. Haribabu
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): S.R. Bodduluri, S. Mathis, P. Maturu, S.R. Satpathy, P.M. Chilton, S. Lira, V.R. Jala, B. Haribabu
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): S.R. Bodduluri, S.R. Satpathy, P.M. Chilton, M. Locati, V.R. Jala, B. Haribabu
Writing, review, and/or revision of the manuscript: S.R. Bodduluri, P.M. Chilton, M. Locati, A. Mantovani, V.R. Jala, B. Haribabu
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): V.R. Jala, B. Haribabu
Study supervision: V.R. Jala, B. Haribabu
Other (performed research experiments): P. Maturu
This work was supported by NIH grants CA-138623 (B. Haribabu) KLCRP (V.R. Jala) and James Graham Brown Cancer Center at U of L. Part of this work was performed with assistance of the U of L Microarray Facility, which is supported by NCRR COBRE P20RR018733, KY-INBRE NCRR P20RR016481, and the J.G. Brown Cancer Center at U of L.
The authors thank Dr. Nejat Egilmez for critical reading of the manuscript. We thank Michelle Smith and Becca Baby for expert technical assistance.
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.