Dendritic cells form clusters in vivo, but the mechanism behind this has not been determined. In this article, we demonstrate that monocytes from mice deficient in the chemokine receptors CCR1, CCR2, CCR3, and CCR5 display reduced clustering in vitro, which is associated with impaired dendritic cell and macrophage differentiation. We further show that the differentiating cells themselves produce ligands for these receptors that function, in a redundant manner, to regulate cell clustering. Deletion of, or pharmacological blockade of, more than one of these receptors is required to impair clustering and differentiation. Our data show that chemokines and their receptors support clustering by increasing expression of, and activating, cell-surface integrins, which are associated with cell–cell interactions and, in the context of monocyte differentiation, with reduced expression of Foxp1, a known transcriptional suppressor of monocyte differentiation. Our data therefore provide a mechanism whereby chemokines and their receptors typically found in inflammatory environments can interact to promote murine monocyte differentiation to macrophages and dendritic cells.

An effective immune response requires coordinated and sequential interactions between leukocytes at the site of inflammation and in specialized anatomical compartments. This complex web of recruitment and interactions between different immune and inflammatory cell types throughout the body is largely controlled by chemokines, small, highly conserved cytokines capable of inducing migration of leukocytes to and from tissues (1, 2). The chemokine family is specifically defined by the presence of a conserved cysteine motif and is divided into CC, CXC, XC, and CX3C subfamilies on the basis of the specific orientation of this motif. Chemokines interact with their target cells by binding to members of the seven-transmembrane-spanning family of G protein–coupled receptors, which are classified as CCRs, CXCRs, XCR, and CX3CR according to the subfamily of chemokines that they bind (3). There is also a subfamily of receptors called atypical chemokine receptors that fine-tune chemokine responses, sculpt intratissue gradients, and contribute to the overall tissue-specific function of chemokines (4).

Based on expression pattern and function, chemokines and their receptors can be classified as being either “inflammatory” or “homeostatic” according to the biological contexts in which they function (5, 6). Inflammatory chemokines, such as CCL2, CCL5, or CXCL8, are rapidly induced in response to tissue damage or infection and recruit inflammatory leukocytes as part of the innate immune response. Inflammatory chemokines can be made by any cell, and in any tissue, experiencing physical or microbial insults. In contrast, homeostatic chemokines are made in a tissue-restricted fashion and are typically not inducible in response to damage or infection. Homeostatic chemokines, and their receptors, are involved in the precise navigation of cells to individual tissue compartments and contribute to the establishment of adaptive immune responses.

We have been interested in trying to delineate roles for the chemokine receptors CCR1, CCR2, CCR3, and CCR5 (henceforth referred to as inflammatory CC-chemokine receptors (iCCRs) in leukocyte recruitment to, and movement within, inflamed tissue sites. In essence, these receptors regulate all aspects of nonneutrophilic myeloid cell recruitment during inflammation and are therefore important potential therapeutic targets in inflammatory disease (3). Analysis of these receptors, and their involvement in the inflammatory response, has been confounded by the complicated and confusing interactions between these receptors and their ligands because the receptors are known to bind multiple ligands, and many of the ligands bind to multiple receptors (7). This, coupled with the fact that these four receptors occupy a tight chromosomal locus in the mammalian genome (8), means that it has been very difficult to precisely analyze their individual, and combinatorial, roles in the inflammatory response. Recently, we have generated bespoke mouse models that have allowed us to start to define the roles for these four receptors in the orchestration of the inflammatory response (9). In terms of monocytes, macrophages, and dendritic cells (DCs), our data highlight a prominent role for CCR2 in monocyte recruitment from the circulation and likely intratissue roles for CCR1 and CCR5. Notably, our analysis indicates that although, in inflamed CCR2−/− mice, intratissue DC and macrophage numbers are severely depleted, it is only in mice deficient for all the iCCRs that tissue DCs and macrophages become essentially undetectable. Importantly, no independent role for CCR1 or CCR5 was detectable in monocyte recruitment or DC or macrophage development. We have therefore hypothesized a role for the iCCRs in supporting inflammatory macrophage and DC maturation, which is complementary to that involved in recruitment and intratissue movement.

In this study, we have used our novel mouse models, alongside pharmacological blockers, to demonstrate a role for the iCCRs in macrophage and DC differentiation that is distinct from their role in cellular recruitment. Our data demonstrate that effective generation of GM-CSF–derived DCs in vitro is critically dependent on apparently redundant signaling through CCR1, CCR2, and CCR5. We further demonstrate that this relates to the ability of these receptors, and their ligands, to initiate LFA-1–dependent coupling between maturing monocytes, which is necessary for efficient DC differentiation.

Wild-type (WT) and iCCR−/− mice were maintained, as described previously (9), in a specific-pathogen-free facility at the Beatson Institute in Glasgow under the auspices of a U.K. Home Office license. All procedures were approved by the local ethical review committee. Animals used were female and between 6 and 8 wk of age.

Tibia and femurs were collected from culled mice, and excess tissue was removed. Both ends of each bone were cut, and bone marrow (BM) was flushed out using 5 ml of RPMI-1640 (Sigma) (10% FBS) in a syringe with a 23G needle. The cell suspension was filtered through a 70-μm cell strainer and spun down for 5 min at 300 × g. The resulting pellet was incubated for 1 min with 1 ml of ACK lysis buffer (Life Technologies), spun down, and resuspended at an appropriate concentration for tissue culture.

  1. GM-CSF–derived DCs and macrophages: A total of 107 BM cells were resuspended in 10 ml of RPMI-1640 (Sigma), 10% FBS, l-glutamine, 50 μM 2-ME, primocin (henceforth cell media), and 20 ng/ml murine recombinant GM-CSF (PeproTech) and placed in tissue culture–treated petri dishes. At day 3, supernatant-containing cells in suspension were collected, spun down 300 × g for 5 min, and the pellet replated in a new petri dish with 10 ml fresh medium. Adherent clustered cells left stuck to the petri dish were collected and analyzed via quantitative PCR (qPCR). This was repeated at days 5 and 7. After collection at day 7, cells were counted and replated as required for phagocytosis and activation assays.

  2. FLT3 DCs: A total of 1.5 × 107 BM cells were resuspended in 10 ml of cell media/100 ng/ml murine recombinant FLT3L (PeproTech) and placed in tissue culture–treated petri dishes. At day 5, two thirds of the supernatant was collected and replaced with fresh FLT3-containing media. At day 8, nonadherent and loosely adherent cells were collected, replated and stimulated for 24/48 h, and analyzed via flow cytometry and ELISA.

  3. CSF-1 macrophages: A total of 5 × 106 BM cells were resuspended in 10 ml of Glasgow’s Minimal Essential Medium, 10% L929 conditioned media, 10% FBS, l-glutamine, 50 μM 2-ME, and primocin in a 90-mm petri dish. At day 4, cells were washed with PBS and 10 ml of fresh media was added to cultures. Differentiated macrophages were collected from day 5 to 7 after replating using TrypLe Select (A12177.01; Life Technologies) as detachment buffer. Macrophages were generally collected at day 5, stimulated for 24 h with cytokines/growth factors, and collected at days 6–7 for analysis.

Reagents and concentrations used to treat the in vitro cultures are listed in Table I.

A total of 107 BM cells were resuspended in 10 ml of cell media/20 ng/ml murine recombinant GM-CSF (PeproTech). Cells were placed in tissue culture–treated petri dishes. At days 3–4, medium was removed and replaced with fresh medium, taking care not to disrupt growing proliferation clusters. Clusters were then either examined under light contrast microscopy (Zeiss Primovert) and imaged by selecting consistent fields of view (AxioCam ERc 5s; Supplemental Fig. 1) and analyzing cluster number using Fiji Software’s particle analyzer (detection of particles > 6000 μm2) or collected by vigorous pipetting at days 4–5 and analyzed via flow cytometry.

Ag presentation and cross-presentation assay

GM-CSF–derived DCs and macrophages were collected, and CD11c+ cells were isolated by positive selection using CD11c-labeled magnetic beads (Miltenyi). The protocol was adapted from an established procedure described previously (10). In brief, 105 DCs were incubated with OVA in different forms for 5 h, stimulated with LPS overnight, fixed with 0.008% glutaraldehyde for 3 min, and then coincubated for 12 h with cognate T cells isolated from OT-1 and OT-2 mouse lymph nodes. For bead-bound OVA (bbOVA), 3-μm latex beads (Polysciences) were incubated with OVA solution in PBS (10 mg/ml) on an agitator overnight at 4°C and used at a ratio of 200:1 beads/DC. Latex beads incubated with BSA were used as control to achieve the following conditions: (1) OVA beads only (100% OVA), (2) 50% OVA beads:50% BSA beads, (3) 25% OVA beads:75% BSA, and (4) BSA beads only (OVA 0%). For soluble OVA (sOVA), the range was from 200 to 25 μg/ml. For OVA peptides, OVA 257–264 was 8 to 1 μg/ml, and OVA 323–339 was 2 to 0.25 μg/ml. T cells from OT-1 and OT-2 mice were coincubated with fixed DCs at a ratio of 2:1 T cells/DC for 12 h. The supernatant was then collected, and IL-2 levels were measured via ELISA.

Adhesion assay

Recombinant mouse ICAM-1–Fc Chimera (553004; BioLegend) was resuspended to 10 μg/ml in PBS. A total of 50 μl of ICAM-1–Fc was then pipetted on top of glass coverslips and incubated overnight at 4°C. The ICAM solution was then removed, and the coverslips were washed with PBS and blocked with 50 μl of 1% BSA in PBS at room temperature for 30 min. Coverslips were then washed again, and 50 μl of GM-CSF cell suspension (200,000/ml) from WT or iCCR−/− cultures was added to the coverslips. The slides were then incubated at 37°C for 20 min and washed 3× before counting. Five fields of view were recorded for each coverslip with a light microscope (Zeiss Primovert, AxioCam ERc 5s) at ×100 magnification. Cell quantification was performed via Fiji Software Particle Analyzer Program (Supplemental Fig. 2). The setting for detection was particles > 50 μm.

Ag uptake assay

Day 8 DCs/macrophages were incubated at 37°C with OVA Alexa Fluor 488 Conjugate (Invitrogen) at various concentrations (60 to 15 μg/ml) for 30 min. Controls were performed at 4°C. Cells were washed in PBS before staining with Abs for flow cytometry. Fluorescence on the FITC channel was detected on the MACSQuant and analyzed via FlowJo.

qPCR

RNA was extracted using the RNeasy Minikit with on-column DNAse I digestion (Qiagen), following manufacturer’s instructions. RNA was converted into cDNA using the High-Capacity RNA-to-cDNA Kit (Applied Biosystems). iCCR expression was measured by qPCR (PerfeCTa SYBR Green FastMix; Quanta Biosciences) on a QuantStudioTM7 Flex Real-Time PCR system (Applied Biosystems) using standard curves specific for each gene. Results were normalized to the expression of the housekeeping gene GAPDH. Primers used are listed in Table II.

Surface Ag staining

Starting from single-cell suspensions, cells were first stained with a fixable viability dye (Zombie Aqua; BioLegend), and Fc receptors were blocked using FcR Blocking-Reagent (Miltenyi Biotec). Surface marker staining was performed with the appropriate Ab mix diluted in FACS Buffer (PBS/2 mM/EDTA/0.5% FBS) (Table III) on ice for 20 min. Cells were washed, fixed with Fixation Buffer (420801; BioLegend), and resuspended in FACS buffer before analysis. Data were acquired using either the MACSQuant (Miltenyi) or LSRII (BD Biosciences) flow cytometer and analyzed using MACSQuantify version 2.5 software (Miltenyi Biotec). Voltages and compensation were determined using UltraComp eBeads (01-2222-42; Bioscience) as single controls. Positive staining was determined using Fluorescence-Minus-One controls. The gating strategy is outlined in Supplemental Fig. 3.

In vivo models of inflammation

Granuloma formation

Anesthetized WT and iCCR−/− mice were injected s.c. with 50 μl of VacciGrade CFA (Invivogen) and culled 3 wk later. Granuloma and draining inguinal lymph nodes were collected for histology and flow cytometry.

Air pouch model

WT mice were anesthetized, and an air pouch was produced by s.c. injection of 3 ml of sterile air into the back. The process was repeated every 2 d for a total of three times (days 0, 2, and 4). On day 6, 106 sorted Ly6C++ monocytes were resuspended in 2 ml of GM-CSF–containing media and injected into the air pouch. Three days later, the mice were culled, and the air pouch fluid and membrane were collected for histology and flow cytometry.

Aldara skin inflammation

Imiquimod is a TLR-7 antagonist that induces the localized production of IL-1b, TNF-a, and other proinflammatory cytokines. One fourth of the content of an Aldara cream sachet (5% cream, ∼3.12 mg of imiquimod) was applied to the shaved back of a mouse every day for 4 d. On the fifth day, mice were culled, and the shaved skin was removed. The tissue was then cut coarsely and digested enzymatically to achieve a single-cell suspension that could be stained and analyzed via flow cytometry.

LPS lung challenge

A total of 30 μl of a 250 ng/ml solution of LPS was dropped onto the nostril of anesthetized mice. Mice were culled 48 h later, a catheter was surgically placed in the trachea through the neck, and 1 ml of PBS with 2 mM EDTA was injected into the lung. The fluid was removed with the same syringe, and the lungs were washed three times overall with a final bronchoalveolar lavage volume of around 3 ml. Cells were counted, spun down, resuspended, and stained for flow cytometry.

Studies on monocyte differentiation have identified two main cytokines responsible for driving macrophage proliferation and survival: GM-CSF and CSF-1 (11). CSF-1 appears to regulate “steady-state” macrophage survival, differentiation, and proliferation, while GM-CSF is responsible for shaping macrophage function during inflammatory responses (12). Similarly, exposing BM monocytes in vitro to either GM-CSF or FLT3L will produce DCs with very different phenotypes (13). Because GM-CSF secretion is increased during inflammation (14), we hypothesized that chemokine receptors prominent during inflammatory responses might be involved in the differentiation of GM-CSF–derived macrophages and DCs. To test this, we examined in vitro GM-CSF–dependent monocyte and macrophage differentiation from WT monocytes and monocytes deficient in CCRs 1, 2, 3, and 5 (iCCR−/−) (9).

After culturing BM monocytes for 5 d in GM-CSF, the differences between WT and iCCR−/− cultures were striking (Tables I, II, III). Although the WT cultures showed multiple clusters of differentiating macrophages and DCs, these clusters were almost completely absent in iCCR−/− cultures (Fig. 1A). Although clusters did eventually appear by day 7 in iCCR−/− cultures, these were significantly smaller and fewer (Fig. 1B) compared with those seen in WT cultures.

FIGURE 1.

iCCR−/− deficiency impairs in vitro generation of GM-CSF–derived macrophages and DCs.

(A) Representative bright-field images at ×40 and ×100 original magnifications, showing proliferation clusters in GM-CSF WT and iCCR−/− monocyte-derived DC cultures. (B) Quantification of cluster numbers (at day 5) and cluster size (day 7) in GM-CSF cultures grown in 12-well plates from WT and ICCR−/− bone marrow cells. Cultures were grown in triplicate and five fixed field-of-view images were taken for each, for a total of 15 data points. Cluster quantification and size determination were performed with Fiji Software’s particle analysis by selecting clusters >6000 μm2. Unpaired t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (C) Representative FACS plot showing the proportions of the four main subpopulations identified in WT and iCCR−/− GM-CSF cultures (mature DCs: CD11c+MHCII++, immature DCs: CD11c+MHCII+, macrophages: CD11c+MHCIIint, monocytes: CD11cMHCII). (Di) Cell counts of both the adherent and suspended fractions, in WT and iCCR−/− GM-CSF cultures, showing the total yields for each of the four subpopulations. Results elaborated from an average of at least three independent GM-CSF cultures for both WT and iCCR−/− samples. Unpaired t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Dii) Pie charts showing the average prevalence of each subpopulation in WT and iCCR−/− GM-CSF cultures. (E) Expression of nuclear protein Ki67 in WT and iCCR−/− GM-CSF cultures, measured as MFI, with associated representative histograms (WT: red, iCCR−/−: blue). *p < 0.05, **p < 0.01, ****p < 0.0001.

FIGURE 1.

iCCR−/− deficiency impairs in vitro generation of GM-CSF–derived macrophages and DCs.

(A) Representative bright-field images at ×40 and ×100 original magnifications, showing proliferation clusters in GM-CSF WT and iCCR−/− monocyte-derived DC cultures. (B) Quantification of cluster numbers (at day 5) and cluster size (day 7) in GM-CSF cultures grown in 12-well plates from WT and ICCR−/− bone marrow cells. Cultures were grown in triplicate and five fixed field-of-view images were taken for each, for a total of 15 data points. Cluster quantification and size determination were performed with Fiji Software’s particle analysis by selecting clusters >6000 μm2. Unpaired t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (C) Representative FACS plot showing the proportions of the four main subpopulations identified in WT and iCCR−/− GM-CSF cultures (mature DCs: CD11c+MHCII++, immature DCs: CD11c+MHCII+, macrophages: CD11c+MHCIIint, monocytes: CD11cMHCII). (Di) Cell counts of both the adherent and suspended fractions, in WT and iCCR−/− GM-CSF cultures, showing the total yields for each of the four subpopulations. Results elaborated from an average of at least three independent GM-CSF cultures for both WT and iCCR−/− samples. Unpaired t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Dii) Pie charts showing the average prevalence of each subpopulation in WT and iCCR−/− GM-CSF cultures. (E) Expression of nuclear protein Ki67 in WT and iCCR−/− GM-CSF cultures, measured as MFI, with associated representative histograms (WT: red, iCCR−/−: blue). *p < 0.05, **p < 0.01, ****p < 0.0001.

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Table I.

Concentration of reagents used to treat in vitro cultures

ReagentsConcentrationSupplierCode
LPS 100 ng/ml Invivogen tlr-eblps 
GM-CSF 20 ng/ml Miltenyi 130-095-739 
FLT3l 100 ng/ml PeproTech 250-31L 
UCB 35625 20 µM Tocris Biosciences 2757 
Maraviroc 20 µM Tocris Biosciences 3756 
BMS CCR2 22 20 µM Tocris Biosciences 3129 
rNIF 1.2 µg/ml R&D 5845-NF 
BIRT377 5 µg/ml Tocris Biosciences 4776/10 
ICAM 10 µg/ml BioLegend 553004 
ReagentsConcentrationSupplierCode
LPS 100 ng/ml Invivogen tlr-eblps 
GM-CSF 20 ng/ml Miltenyi 130-095-739 
FLT3l 100 ng/ml PeproTech 250-31L 
UCB 35625 20 µM Tocris Biosciences 2757 
Maraviroc 20 µM Tocris Biosciences 3756 
BMS CCR2 22 20 µM Tocris Biosciences 3129 
rNIF 1.2 µg/ml R&D 5845-NF 
BIRT377 5 µg/ml Tocris Biosciences 4776/10 
ICAM 10 µg/ml BioLegend 553004 
Table II.

Primer sequences used in this study

PrimerSequence (5′–3′)
CCR1 ORF Stan1 ACTTTTGGCATCATCACCAG 
CCR1 ORF Stan2 CTCAGATTGTAGGGGGTCCA 
CCR1 ORF QPCR1 GCCCTCATTTCCCCTACAA 
CCR1 ORF QPCR2 CGGCTTTGACCTTCTTCTCA 
CCR2 ORF Stan1 ACCACAGAATCAAAGGAAATGG 
CCR2 ORF Stan2 GTTGCCCACAAAACCAAAGA 
CCR2 ORF QPCR1 TGACAAGGCTCACCATCATC 
CCR2 ORF QPCR2 TCAGTTCATCCACGGCATACT 
CCR3 ORF Stan1 GCCATCCGTCTTATTTTTGTTG 
CCR3 ORF Stan2 ATTTCTTGCTCCCCAGTTGA 
CCR3 ORF QPCR1 ACCTTCGGCTCTTTTTCCAC 
CCR3 ORF QPCR2 TGTTCTTTCCATTTTCTCACCA 
CCR5 ORF Stan1 GGCCAACAATTGCTTTAACCT 
CCR5 ORF Stan2 AGCAAACACAGCATGGACAA 
CCR5 ORF QPCR1 GGATTTTCAAGGGTCAGTTCC 
CCR5 ORF QPCR2 GAAGACCATCATGTTACCCACA 
GAPDH Stan1 TGAACGGGAAGCTCACTGGC 
GAPDH Stan2 TCCACCACCCTGTTGCTGTAG 
GAPDH QPCR1 ATGTGTCCGTCGTGGATCTGAC 
GAPDH QPCR2 GTTGCTGTTGAAGTCGCAGGAG 
PrimerSequence (5′–3′)
CCR1 ORF Stan1 ACTTTTGGCATCATCACCAG 
CCR1 ORF Stan2 CTCAGATTGTAGGGGGTCCA 
CCR1 ORF QPCR1 GCCCTCATTTCCCCTACAA 
CCR1 ORF QPCR2 CGGCTTTGACCTTCTTCTCA 
CCR2 ORF Stan1 ACCACAGAATCAAAGGAAATGG 
CCR2 ORF Stan2 GTTGCCCACAAAACCAAAGA 
CCR2 ORF QPCR1 TGACAAGGCTCACCATCATC 
CCR2 ORF QPCR2 TCAGTTCATCCACGGCATACT 
CCR3 ORF Stan1 GCCATCCGTCTTATTTTTGTTG 
CCR3 ORF Stan2 ATTTCTTGCTCCCCAGTTGA 
CCR3 ORF QPCR1 ACCTTCGGCTCTTTTTCCAC 
CCR3 ORF QPCR2 TGTTCTTTCCATTTTCTCACCA 
CCR5 ORF Stan1 GGCCAACAATTGCTTTAACCT 
CCR5 ORF Stan2 AGCAAACACAGCATGGACAA 
CCR5 ORF QPCR1 GGATTTTCAAGGGTCAGTTCC 
CCR5 ORF QPCR2 GAAGACCATCATGTTACCCACA 
GAPDH Stan1 TGAACGGGAAGCTCACTGGC 
GAPDH Stan2 TCCACCACCCTGTTGCTGTAG 
GAPDH QPCR1 ATGTGTCCGTCGTGGATCTGAC 
GAPDH QPCR2 GTTGCTGTTGAAGTCGCAGGAG 
Table III.

List of Abs and fluorophores used in the study

MarkerFluorophoreSupplierClone Number
CD11b v500 BD Biosciences m1/70 
MHCII BV421 BioLegend i-a/i-e 
CD11c Allophycocyanin BioLegend n418 
CD86 PerCP BioLegend gl-1 
CD40 FITC BioLegend 3-23 
CD80 PE BioLegend 16-10A1 
F480 Pe/Cy7 BioLegend bm8 
CD11a FITC BioLegend m17/4 
CD11a PE BioLegend 2D7 
Live/dead Allophycocyanin/Cy7 Invitrogen 1965980 
Foxp1 PE Miltenyi REA682 
CCL19 AF647 Almac CAF-6 
OVA AF488 Thermo Fisher 34781 
Fc block — Miltenyi 130-092-575 
MarkerFluorophoreSupplierClone Number
CD11b v500 BD Biosciences m1/70 
MHCII BV421 BioLegend i-a/i-e 
CD11c Allophycocyanin BioLegend n418 
CD86 PerCP BioLegend gl-1 
CD40 FITC BioLegend 3-23 
CD80 PE BioLegend 16-10A1 
F480 Pe/Cy7 BioLegend bm8 
CD11a FITC BioLegend m17/4 
CD11a PE BioLegend 2D7 
Live/dead Allophycocyanin/Cy7 Invitrogen 1965980 
Foxp1 PE Miltenyi REA682 
CCL19 AF647 Almac CAF-6 
OVA AF488 Thermo Fisher 34781 
Fc block — Miltenyi 130-092-575 

Flow cytometric analysis of both the suspended and the adherent fraction in WT GM-CSF cultures revealed four different leukocyte populations characterized on the basis of CD11c and MHC class II (MHCII) expression (Fig. 1C, Supplemental Fig. 3). Further flow cytometric analysis of CCR7 expression (Supplemental Fig. 4A), Ag uptake assays (Supplemental Fig. 4B), and costimulatory molecule expression (Supplemental Fig. 4C, Table IV) identified these subsets as undifferentiated monocytes (CD11cMHCII), macrophages (CD11clow, MHCIIlow), immature DCs (CD11c+MHCIIint), and mature DCs (CD11c+MHCII++).

Table IV.

Marker and costimulatory molecule expression on the four main cellular subpopulations

MarkersMonocyteMacrophageImmature DCsMature DCs
CD11b 
CD11c − Low ++ 
MHCII − Low ++ 
CD40 − − Low Low 
CD80 − Low ++ 
CD86 − − Low ++ 
F480 Low − − 
Ly6C − − − 
CCR7 − − ++ 
CD115 − − 
CD206 − Low − 
CD64 Low − − − 
Arg-1 − Low − − 
Inducible NO synthase (after LPS) − ++ − 
CD103 − − − − 
CD8a − − − − 
Phagocytosis (Ova) − ++ ++ 
Ag processing (pHrodo) − Low ++ 
MarkersMonocyteMacrophageImmature DCsMature DCs
CD11b 
CD11c − Low ++ 
MHCII − Low ++ 
CD40 − − Low Low 
CD80 − Low ++ 
CD86 − − Low ++ 
F480 Low − − 
Ly6C − − − 
CCR7 − − ++ 
CD115 − − 
CD206 − Low − 
CD64 Low − − − 
Arg-1 − Low − − 
Inducible NO synthase (after LPS) − ++ − 
CD103 − − − − 
CD8a − − − − 
Phagocytosis (Ova) − ++ ++ 
Ag processing (pHrodo) − Low ++ 

iCCR−/− GM-CSF cultures have a significantly lower cell yield for all subpopulations (Fig. 1Di), with the biggest decrease observed in the macrophage (CD11clowMHCIIlow) subset. In addition, ∼40% of cells were undifferentiated CD11cMHCII monocytes, compared with 20% of cells in WT cultures (Fig. 1Dii). Furthermore, iCCR cultured cells express less Ki67 than WT cells, suggesting reduced proliferative activity (Fig. 1E). Finally, levels of cell death in the cultures were extremely low and indistinguishable between WT and iCCR cultures (data not shown). Overall, these data show that iCCR deficiency is associated with a small reduction in proliferation but a major reduction in monocytic differentiation in these in vitro cultures.

Thus, iCCR-deficient monocytes have an impaired ability to differentiate into macrophages and DCs in GM-CSF–supplemented cultures, and this is associated with lack of formation of clusters of proliferating and differentiating cells.

To determine whether the absence of chemokine receptors could also affect DC function, we collected the suspended cell fraction (enriched for immature and mature DCs) from both WT and ICCR−/− cultures at day 6, and CD11c+ DCs were isolated by positive selection using CD11c-labeled magnetic beads. DCs were then pulsed with three different forms of OVA Ag (either bead-bound, soluble, or OVA peptide fragments) and coincubated with T cells obtained from the lymph nodes of OT-1 and OT-2 mice. Levels of Ag presentation and T cell activation were assessed by analyzing IL-2 levels in the supernatant.

We observed a reduction in IL-2 levels secreted from both CD4 and CD8 T cells coincubated with iCCR−/− DCs (Fig. 2Ai, ii), but, in both cases, only when DCs were primed with bbOVA. IL-2 levels were identical between WT and iCCR−/− if DCs were exposed to sOVA, or preprocessed OVA peptides (323–339) and (257–264), which bind directly to MHCII and MHC class I (MHCI), respectively. Flow cytometric analysis of WT and iCCR−/− immature and mature DCs showed that they express identical levels of costimulatory molecules CD40, CD80, and CD86 (Fig. 2Bi, ii), and have the same Ag-processing capacity (inferred by similar levels of endosomal acidification; (Fig. 2Biii).

FIGURE 2.

FLT3L DC and CSF-1 macrophage differentiation and function are unaffected by iCCR deletion.

(A) Graphs showing ability to present Ag (Ai) and cross-present Ag (Aii) by magnetically sorted WT and iCCR−/− DCs coincubated with OT-2 and OT-1 T-cells (inferred via IL-2 levels). DCs were loaded with either bbOVA (simulates particulate Ag), sOVA, or control ovalbumin peptides OVA(323–339) and OVA(257–264). Results elaborated from an average of four independent GM-CSF cultures for both WT and iCCR−/− samples. Values shown on graph represent the midrange for each condition: bbOVA (50% OVA: 50% BSA), sOVA (50 μg/ml), OVA(323–339) (2 μg/ml), and OVA(257–264) (0.5 μg/mL). Unpaired t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (B) Costimulatory molecule expression by immature (Bi) andmature DCs (Bii) from WT and ICCR−/− cultures. (Biii) Representative histograms compare WT (red) and ICCR−/− (blue) mature DC ability with process Ag, assessed by pHrodo fluorescence (control: purple). (Ci) Representative FACS plots showing CD11c MHCII subpopulations in WT and iCCR−/− cultures after 9 d of growth under FLT3L. (Cii) IL-12p70 and IL-10 secretion by WT (black) and iCCR−/− (white) FLT3L DCs measured via ELISA in 96-well plates (100,000 cells/well) before and after overnight LPS stimulation (n = 4). (Ciii) Representative histograms showing uptake of OVA-488 by FLT3L DCs (CD11c+MHCII++) (WT: red, iCCR−/−: blue) before and after overnight LPS stimulation (n = 4). (Di) Representative FACS plots showing CD11b+F480+ cells in WT and iCCR cultures after 7 d of growth under CSF-1. (Dii) Representative histograms showing uptake of OVA-488 by macrophages (CD11b+F480+) (WT: red, iCCR: blue). (Diii) Costimulatory molecule expression (MHCII, CD40, CD80, CD86) in WT (black) and iCCR−/− (white) macrophages (CD11b+F480+) after stimulation with LPS overnight (n = 3). (Div) Cytokine secretion (TNF-α, IL-6, and IL-1b) measured via ELISA by WT (black) and iCCR−/− macrophages (gray) in 96-well plates (100,000 cells/well) after overnight LPS stimulation (n = 4). **p < 0.01, ***p < 0.001.

FIGURE 2.

FLT3L DC and CSF-1 macrophage differentiation and function are unaffected by iCCR deletion.

(A) Graphs showing ability to present Ag (Ai) and cross-present Ag (Aii) by magnetically sorted WT and iCCR−/− DCs coincubated with OT-2 and OT-1 T-cells (inferred via IL-2 levels). DCs were loaded with either bbOVA (simulates particulate Ag), sOVA, or control ovalbumin peptides OVA(323–339) and OVA(257–264). Results elaborated from an average of four independent GM-CSF cultures for both WT and iCCR−/− samples. Values shown on graph represent the midrange for each condition: bbOVA (50% OVA: 50% BSA), sOVA (50 μg/ml), OVA(323–339) (2 μg/ml), and OVA(257–264) (0.5 μg/mL). Unpaired t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (B) Costimulatory molecule expression by immature (Bi) andmature DCs (Bii) from WT and ICCR−/− cultures. (Biii) Representative histograms compare WT (red) and ICCR−/− (blue) mature DC ability with process Ag, assessed by pHrodo fluorescence (control: purple). (Ci) Representative FACS plots showing CD11c MHCII subpopulations in WT and iCCR−/− cultures after 9 d of growth under FLT3L. (Cii) IL-12p70 and IL-10 secretion by WT (black) and iCCR−/− (white) FLT3L DCs measured via ELISA in 96-well plates (100,000 cells/well) before and after overnight LPS stimulation (n = 4). (Ciii) Representative histograms showing uptake of OVA-488 by FLT3L DCs (CD11c+MHCII++) (WT: red, iCCR−/−: blue) before and after overnight LPS stimulation (n = 4). (Di) Representative FACS plots showing CD11b+F480+ cells in WT and iCCR cultures after 7 d of growth under CSF-1. (Dii) Representative histograms showing uptake of OVA-488 by macrophages (CD11b+F480+) (WT: red, iCCR: blue). (Diii) Costimulatory molecule expression (MHCII, CD40, CD80, CD86) in WT (black) and iCCR−/− (white) macrophages (CD11b+F480+) after stimulation with LPS overnight (n = 3). (Div) Cytokine secretion (TNF-α, IL-6, and IL-1b) measured via ELISA by WT (black) and iCCR−/− macrophages (gray) in 96-well plates (100,000 cells/well) after overnight LPS stimulation (n = 4). **p < 0.01, ***p < 0.001.

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The defect in Ag presentation (and cross-presentation) thus appears to stem from the inability of iCCR−/− DCs to acquire large particulate Ags, suggesting a defect in phagocytosis and not Ag processing or presentation.

To determine whether the impaired differentiation of iCCR monocytes in GM-CSF cultures reflected a general block in differentiation, or one that was selectively associated with inflammatory conditions, we examined macrophages and DCs differentiated under different cytokine conditions. Notably, these experiments demonstrated that the earlier phenotype was restricted to GM-CSF–derived macrophages and DCs. DCs generated from WT and iCCR−/− BM cultures containing FLT3-L were identical in terms of subpopulations, costimulatory molecule expression, cytokine secretion profile, and resting and LPS-induced CCR7 expression (Fig. 2C). Similarly, CSF-1–derived macrophages from WT and iCCR−/− BM were also identical in terms of yield, costimulatory molecule expression, Ag uptake ability, and cytokine secretion profile, both before and after overnight stimulation with LPS (Fig. 2D).

Thus, the iCCRs appear to selectively regulate cell clustering and differentiation in GM-CSF BM-derived cultures, and this phenotype is not seen in cultures generating noninflammatory DCs and macrophages.

To determine which receptors may be responsible for the clustering phenotype, we examined iCCR expression in differentiating GM-CSF cultures using PCR. This revealed strong expression of CCR1, CCR2, and CCR5 in the cultures with upregulation over the time course of differentiation (Fig. 3A). CCR3 expression was already very low in freshly isolated BM cells and became undetectable as maturation progressed (Fig. 3B). CCR3 was therefore not included in our subsequent analyses. Thus, transcriptomic analysis reveals that CCR1, CCR2, and CCR5 are prominently expressed in GM-CSF–differentiating cultures, suggesting that one (or more) of these receptors could be involved in cluster formation.

FIGURE 3.

Clustered cells upregulate expression of CCR1, CCR2, and CCR5 in differentiating GM-CSF cultures and secrete their cognate ligands.

(A) CCR1 (green), CCR2 (red), CCR3 (blue), and CCR5 (purple) expression in clustered cells from WT GM-CSF cultures at days 3, 5, and 7. (B) Change in iCCR expression in GM-CSF cultures, from freshly extracted bone marrow cells (day 0) to day 7. Gray area highlights days of rapid cluster formation in GM-CSF cultures (days 2.5–5). Ordinary one-way ANOVA was performed to determine statistical significance, with a p value of 0.05 determined as significant. (C) Cytokine secretion of CCL2 and CCL5 in WT (black) and iCCR−/− (white) cultures before and after administration of 100 ng/ml LPS for 12 h, as measured via ELISA. Results elaborated from an average of at least three independent GM-CSF cultures for both WT and iCCR−/− samples. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. **p < 0.01, ***p < 0.001, ****p < 0.0001.

FIGURE 3.

Clustered cells upregulate expression of CCR1, CCR2, and CCR5 in differentiating GM-CSF cultures and secrete their cognate ligands.

(A) CCR1 (green), CCR2 (red), CCR3 (blue), and CCR5 (purple) expression in clustered cells from WT GM-CSF cultures at days 3, 5, and 7. (B) Change in iCCR expression in GM-CSF cultures, from freshly extracted bone marrow cells (day 0) to day 7. Gray area highlights days of rapid cluster formation in GM-CSF cultures (days 2.5–5). Ordinary one-way ANOVA was performed to determine statistical significance, with a p value of 0.05 determined as significant. (C) Cytokine secretion of CCL2 and CCL5 in WT (black) and iCCR−/− (white) cultures before and after administration of 100 ng/ml LPS for 12 h, as measured via ELISA. Results elaborated from an average of at least three independent GM-CSF cultures for both WT and iCCR−/− samples. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. **p < 0.01, ***p < 0.001, ****p < 0.0001.

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For these receptors to be able to contribute to DC and macrophage differentiation, a source of their ligands has to be available. We therefore next examined expression of chemokine ligands for the iCCRs in developing GM-CSF culture supernatants. This revealed functionally relevant levels of CCL2 (ligand for CCR2) and CCL5 (ligand binding to CCR1 and CCR5), both at rest and after addition of LPS (Fig. 3C). Although the levels of CCL5 were identical in WT and iCCR−/− cultures, CCL2 levels in iCCR−/− cultures were twice as high and remained elevated after exposure to LPS. This difference may be attributed to the known ability of CCR2 to rapidly internalize, and destroy, ligand (15, 16), leading to effective ligand scavenging, which therefore is not seen in the iCCR−/− cultures. Thus, these data demonstrate that ligands for the iCCRs are easily detectable in DC cultures.

We next used the formation of proliferating clusters as a surrogate readout to assess the impact of single iCCR knockouts on monocyte differentiation in GM-CSF cultures. Interestingly, the “cluster-less” phenotype was not observed in GM-CSF cultures grown from BM from single CCR1, CCR2, or CCR5 knockout mice (Fig. 4A), suggesting potential redundancy in the way that these receptors regulate GM-CSF–dependent monocyte differentiation. Due to the genomic proximity of the iCCRs, it is not yet possible to generate mice with select combinations of deletions of these receptors by crossing individual knockout mice. Therefore, to further analyze the roles for receptor combinations in regulating differentiation in GM-CSF–derived cultures, we opted to use well-characterized pharmacological blockers of receptor function. Addition of a combination of antagonists to CCR1, CCR2, and CCR5 to WT GM-CSF cultures recapitulated the iCCR−/− “cluster-less” phenotype with a marked reduction in clustering of the differentiating cells (Fig. 4B).

FIGURE 4.

Cluster formation in WT, single-knockout, iCCR−/−, and WT + iCCR antagonists GM-CSF cultures.

(A) Representative bright-field images of proliferation clusters in GM-CSF cultures at ×40 original magnification from WT, CCR1−/−, CCR2−/−, CCR5−/−, and iCCR−/− bone marrow at day 4. (B) Representative bright-field images of proliferation clusters in GM-CSF cultures at ×40 original magnification from WT, iCCR−/−, and WT cultures treated with iCCR antagonist cocktail from day 0.

FIGURE 4.

Cluster formation in WT, single-knockout, iCCR−/−, and WT + iCCR antagonists GM-CSF cultures.

(A) Representative bright-field images of proliferation clusters in GM-CSF cultures at ×40 original magnification from WT, CCR1−/−, CCR2−/−, CCR5−/−, and iCCR−/− bone marrow at day 4. (B) Representative bright-field images of proliferation clusters in GM-CSF cultures at ×40 original magnification from WT, iCCR−/−, and WT cultures treated with iCCR antagonist cocktail from day 0.

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Different combinations of antagonists were then added to WT cultures to determine the importance of particular iCCRs in cluster development and differentiation. In keeping with the data in (Fig. 4A, none of the individual receptor blockers alone had any significant effect on differentiation in these cultures. Strikingly, however, cluster number decreased significantly if two or more iCCRs were blocked simultaneously (Fig. 5A), and the effects were most pronounced when CCR5 inhibition was involved.

FIGURE 5.

Cluster numbers are reduced on simultaneous inhibition of multiple iCCRs.

(A) Representative bright-field images of proliferation clusters in GM-CSF cultures (12-well plates) at ×40 original magnification from WT cells treated with different combinations of chemokine antagonists, with graph showing cluster quantification for each condition. The same WT bone marrow was used for each culture, each condition was repeated in triplicate, and five fields of view were taken from each well. Each value in the graph represents an average of at least 15 points. Cluster quantification was performed with Fiji Software’s particle analysis by selecting clusters >1800 μm2. Ordinary one-way ANOVA with Dunnett’s multiple comparisons test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (B) Representative bright-field images of proliferation clusters in GM-CSF cultures (12-well plates) at ×40 original magnification from CCR2−/− bone marrow cells treated with different combinations of chemokine antagonists, with graph showing cluster quantification for each condition. The same CCR2−/− bone marrow was used for each culture, each condition was repeated in triplicate, and five fields of view taken from each well. Each value in the graph represents an average of at least 15 points. Cluster quantification was performed with Fiji Software’s particle analysis by selecting clusters >1800 μm2. Ordinary one-way ANOVA with Dunnett’s multiple comparisons test was performed to determine statistical significance, with a p value of 0.05 determined as significant. *p < 0.05, **p < 0.01.

FIGURE 5.

Cluster numbers are reduced on simultaneous inhibition of multiple iCCRs.

(A) Representative bright-field images of proliferation clusters in GM-CSF cultures (12-well plates) at ×40 original magnification from WT cells treated with different combinations of chemokine antagonists, with graph showing cluster quantification for each condition. The same WT bone marrow was used for each culture, each condition was repeated in triplicate, and five fields of view were taken from each well. Each value in the graph represents an average of at least 15 points. Cluster quantification was performed with Fiji Software’s particle analysis by selecting clusters >1800 μm2. Ordinary one-way ANOVA with Dunnett’s multiple comparisons test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (B) Representative bright-field images of proliferation clusters in GM-CSF cultures (12-well plates) at ×40 original magnification from CCR2−/− bone marrow cells treated with different combinations of chemokine antagonists, with graph showing cluster quantification for each condition. The same CCR2−/− bone marrow was used for each culture, each condition was repeated in triplicate, and five fields of view taken from each well. Each value in the graph represents an average of at least 15 points. Cluster quantification was performed with Fiji Software’s particle analysis by selecting clusters >1800 μm2. Ordinary one-way ANOVA with Dunnett’s multiple comparisons test was performed to determine statistical significance, with a p value of 0.05 determined as significant. *p < 0.05, **p < 0.01.

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To minimize any negative effects on cluster development caused by the simultaneous exposure to multiple iCCR antagonists, we repeated the experiment using GM-CSF–derived cultures from CCR2−/− mice supplemented with antagonists of the other receptors. With CCR2 already knocked out, the addition of a single other chemokine receptor antagonist was enough to reduce cluster formation, and this defect was again most profound when a CCR5 antagonist was added to the cultures (Fig. 5B). In addition, there was no difference in cluster number between CCR2−/− cultures treated with CCR1 + CCR5 or all three antagonists simultaneously, indicating that the absence of cluster formation is the result of iCCR inhibition, and not chemokine receptor antagonist toxicity.

Thus, these antagonist-based studies demonstrate the requirement for multiple simultaneous receptor blockades for impairment of monocyte differentiation in GM-CSF cultures indicating redundancy of chemokine receptor function in this context.

Integrins facilitate cell-to-cell and cell-to-extracellular matrix interactions (1719) and can be activated to a high-affinity/avidity state on leukocytes by chemokine receptor “inside-out signaling” after ligand binding. Integrin engagement has been shown to promote monocyte differentiation (20) into macrophages and is an important mediator of particle clearance in phagocytosis (21).

mAbs to integrins LFA-1 or CD31 can also inhibit Langerhans-type DC cluster formation in vitro (22). We therefore hypothesized that reduced iCCR signaling–dependent integrin engagement could be associated with impaired cell-to-cell communication and phagocytosis, and therefore reduced DC and macrophage differentiation, in GM-CSF–dependent iCCR−/− cultures.

Integrin expression and activation levels were therefore analyzed using flow cytometry on monocytes (Fig. 6A) derived from WT, CCR2−/−, and iCCR−/− GM-CSF cultures and WT cultures treated with the chemokine receptor antagonist mixture. Two different Abs were used, one detecting both the high- and low-affinity versions of LFA-1 (clone M174), and another capable of preferentially binding to the low-affinity version (clone 2D7) (23). Overall expression of both low- and high-affinity forms of LFA-1, as detected using the M174 Ab, was significantly lower in iCCR−/− and WT + antagonist CD11cMHCII monocytes compared with WT and CCR2−/− monocytes (Fig. 6A). However, 2D7 analysis demonstrated that LFA-1+ monocytes from iCCR−/− and WT + antagonist cultures had a higher mean fluorescent intensity (MFI) for the low-affinity version of LFA-1 compared with monocytes from WT cultures, indicating that not only do undifferentiated monocytes have decreased integrin expression when iCCRs are absent or inhibited, but the LFA-1 that is expressed is preferentially of the low-affinity form.

FIGURE 6.

Integrin expression and adhesion to ICAM-1 by WT and iCCR−/− GM-CSF–derived cells.

(Ai) Representative FACS plots showing the WT and iCCR−/− monocyte (CD11cMHCII) population analyzed for integrin expression (blue box). (Aii) Expression of integrin LFA-1 (M174 and 2D7 clone) on CD11cMHCII monocytes from GM-CSF cultures from WT, CCR2−/−, ICCR−/−, and WT+ antagonist-treated bone marrow cells, measured as MFI via flow cytometry. Results elaborated from an average of at least three independent GM-CSF cultures for both WT and iCCR−/− samples. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (B) Representative bright-field images at ×100 original magnification of WT and iCCR cells on ICAM-1–coated coverslips. Graph summarizes cell number counts in bright-field images detected through ‘Particle Count’ program on Fiji Image Software (number of cells >50 μm2). Experiment was performed in triplicates, with five fields of view taken for each coverslip, resulting in an average of 15 total measurements for both WT and iCCR−/−. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Ci) Representative images (×40 original magnification) and graph showing cluster number at day 5 in WT GM-CSF cultures (12-well plate) treated from day 0 with integrin antagonists NIF and BIRT377, with media changed every second day. The same WT bone marrow was used for each culture, each condition was repeated in triplicate, and four fields of view were taken from each well for a total of 12 points per condition. Ordinary one-way ANOVA with Dunnett’s multiple comparisons test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Cii) Effect of integrin antagonists on the cellular yields for each CD11c/MHCII subpopulation on WT GM-CSF cultures. Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Di) Phagocytic uptake of 3-μm latex beads by WT, iCCR−/−, and integrin antagonist-treated DCs. DCs were incubated with latex beads at a ratio of 200:1 (beads/DC) for 12 h with LPS at 37°C. Values show number of beads per DC, with >100 individual cells counted from 10 fixed fields of view. Representative bright-field images at ×100 original magnification show internalized beads in WT, BIRT377, and iCCR−/− DCs. (Dii) Foxp1 expression in clustered monocytes (CD11clowMHCIIlow) from WT, iCCR−/−, and CCR2−/− GM-CSF cultures, measured as MFI via intracellular flow cytometry. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

FIGURE 6.

Integrin expression and adhesion to ICAM-1 by WT and iCCR−/− GM-CSF–derived cells.

(Ai) Representative FACS plots showing the WT and iCCR−/− monocyte (CD11cMHCII) population analyzed for integrin expression (blue box). (Aii) Expression of integrin LFA-1 (M174 and 2D7 clone) on CD11cMHCII monocytes from GM-CSF cultures from WT, CCR2−/−, ICCR−/−, and WT+ antagonist-treated bone marrow cells, measured as MFI via flow cytometry. Results elaborated from an average of at least three independent GM-CSF cultures for both WT and iCCR−/− samples. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (B) Representative bright-field images at ×100 original magnification of WT and iCCR cells on ICAM-1–coated coverslips. Graph summarizes cell number counts in bright-field images detected through ‘Particle Count’ program on Fiji Image Software (number of cells >50 μm2). Experiment was performed in triplicates, with five fields of view taken for each coverslip, resulting in an average of 15 total measurements for both WT and iCCR−/−. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Ci) Representative images (×40 original magnification) and graph showing cluster number at day 5 in WT GM-CSF cultures (12-well plate) treated from day 0 with integrin antagonists NIF and BIRT377, with media changed every second day. The same WT bone marrow was used for each culture, each condition was repeated in triplicate, and four fields of view were taken from each well for a total of 12 points per condition. Ordinary one-way ANOVA with Dunnett’s multiple comparisons test was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Cii) Effect of integrin antagonists on the cellular yields for each CD11c/MHCII subpopulation on WT GM-CSF cultures. Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Di) Phagocytic uptake of 3-μm latex beads by WT, iCCR−/−, and integrin antagonist-treated DCs. DCs were incubated with latex beads at a ratio of 200:1 (beads/DC) for 12 h with LPS at 37°C. Values show number of beads per DC, with >100 individual cells counted from 10 fixed fields of view. Representative bright-field images at ×100 original magnification show internalized beads in WT, BIRT377, and iCCR−/− DCs. (Dii) Foxp1 expression in clustered monocytes (CD11clowMHCIIlow) from WT, iCCR−/−, and CCR2−/− GM-CSF cultures, measured as MFI via intracellular flow cytometry. Student t test was performed to determine statistical significance, with a p value of 0.05 determined as significant. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

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To further determine whether the decreased integrin expression and activation in iCCR−/− cultures correlate with decreased cell adhesion, we performed a static adhesion assay by adding WT and iCCR−/− GM-CSF–derived cells (from cultures allowed to grow for 8 d to ensure maximum yield of fully differentiated macrophages and DCs) to coverslips coated with the LFA-1 ligand ICAM-1. iCCR−/− cells were found to adhere to these coverslips in lower numbers (Fig. 6B) compared with WT cells, suggesting that their integrin-dependent interaction with immobilized ICAM-1 is weaker and less stable. Thus, iCCR−/− monocytes, as well as their GM-CSF–differentiated progeny, display impaired adhesiveness associated with reduced integrin activation.

To confirm that lack of integrin activation is indeed responsible for the observed cluster-less phenotype, we cultured GM-CSF–derived cells with antagonists of distinct β2 integrins: CD11a (BIRT377) and CD11b (NIF). As shown in (Fig. 6Ci, almost no clusters were observed in cultures treated with BIRT377 and a combination of BIRT377 and NIF, whereas cultures treated with NIF only showed no difference in cluster numbers compared with untreated cultures. The addition of BIRT377 to WT cultures also resulted in yields of CD11c/MHCII subsets that were similar to what was observed in iCCR−/− cultures, with a very prominent decrease in macrophages and an overall accumulation of undifferentiated monocytes (Fig. 6Cii). Ag uptake, assessed via phagocytosis of latex beads, was also severely affected in BIRT377-treated DCs (Fig. 6Di), suggesting that lack of LFA-1 engagement alone is directly linked to both the cluster-less phenotype and the reduced phagocytic rates seen in iCCR−/− DCs.

In terms of molecular mechanisms that might explain the association of impaired integrin activation and reduced monocyte differentiation, we next compared Foxp1 expression in WT and iCCR−/− GM-CSF–dependent cultures. Foxp1 is a transcriptional repressor involved in monocyte-to-macrophage differentiation, which is negatively regulated by integrins (24). Intracellular staining for Foxp1 revealed higher expression of Foxp1 in iCCR−/− compared with WT and CCR2−/− monocytes (Fig. 6Dii), further suggesting that defective integrin activation and associated downstream signaling are associated with the phenotype observed.

Next, we wanted to determine whether monocyte-derived DCs in vivo also express CCR1, CCR2, and CCR5, or whether this coexpression is an exclusive in vitro phenotype. To test this, we isolated CD11b+ DCs from the inflamed site of iCCR reporter mice (25) in three different inflammation models: LPS lung challenge, air pouch model containing inflammatory mediator Carrageenan, and acute skin inflammation using imiquimod. In all three inflammatory models, a large portion of CD11b+CD11c+MHCII++ DCs coexpressed CCR1, CCR2, and CCR5 (Fig. 7A), confirming expression in vivo and suggesting these receptors may play an important role in DC differentiation and function in vivo.

FIGURE 7.

In vivo WT and iCCR−/− DC and macrophage maturation.

(A) CCR1 (green), CCR2 (red), and CCR5 (purple) expression in CD11b+CD11c+MHCII++ DCs collected from the inflamed site in three different inflammation models on an iCCR reporter strain: air pouch model (inflammatory mediator: Carrageenan), LPS lung challenge (inflammatory mediator: LPS), and skin inflammation (inflammatory mediator: imiquimod. Associated pie charts show preferred iCCR combination by CD11b+CD11c+MHCII++ DCs, with multiple colors representing coexpression (CCR2: red, CCR1: green, CCR5: purple; n > 4. (Bi) Stitched ×40 bright-field image (and representative ×100 image) of H&E staining of a WT and iCCR−/− granuloma. The white box of ×100 WT granuloma slice highlights a multinucleated giant cell. Associated representative FACS plots show lack of CD11c+ MHCII+ cells. (Bii) Summary of all leukocyte subsets found in WT and iCCR−/− granulomas (n = 4). Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Ci) Representative FACS plots and associated graph, showing undifferentiated Ly6C++ monocytes (red arrow) and differentiated Ly6ClowCD11c+ cells in the air pouch fluid of WT mice injected with WT or iCCR−/− monocytes. (Cii) Representative bright-field images showing no cellular aggregation in air pouch fluid collected from WT mice injected with either WT or iCCR−/− monocytes (×40 original magnification). (Di) Toluidine blue whole-mount staining of air pouch membrane of WT mice injected with either WT or iCCR−/− monocytes (×40 original magnification). (Dii) Graph summarizing cellular yields recovered from the air pouch membrane of WT mice injected with either WT or iCCR−/− monocytes (CD11c+MHCIIint DCs, CD11c+MHCII++ DCs, and CD11b+CD64+ macrophages) (n = 6); Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Diii) Graph summarizing yields of DCs and macrophages in the lymph nodes and spleen of WT mice injected with either WT or iCCR−/− monocytes. Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. *p < 0.05, **p < 0.01.

FIGURE 7.

In vivo WT and iCCR−/− DC and macrophage maturation.

(A) CCR1 (green), CCR2 (red), and CCR5 (purple) expression in CD11b+CD11c+MHCII++ DCs collected from the inflamed site in three different inflammation models on an iCCR reporter strain: air pouch model (inflammatory mediator: Carrageenan), LPS lung challenge (inflammatory mediator: LPS), and skin inflammation (inflammatory mediator: imiquimod. Associated pie charts show preferred iCCR combination by CD11b+CD11c+MHCII++ DCs, with multiple colors representing coexpression (CCR2: red, CCR1: green, CCR5: purple; n > 4. (Bi) Stitched ×40 bright-field image (and representative ×100 image) of H&E staining of a WT and iCCR−/− granuloma. The white box of ×100 WT granuloma slice highlights a multinucleated giant cell. Associated representative FACS plots show lack of CD11c+ MHCII+ cells. (Bii) Summary of all leukocyte subsets found in WT and iCCR−/− granulomas (n = 4). Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Ci) Representative FACS plots and associated graph, showing undifferentiated Ly6C++ monocytes (red arrow) and differentiated Ly6ClowCD11c+ cells in the air pouch fluid of WT mice injected with WT or iCCR−/− monocytes. (Cii) Representative bright-field images showing no cellular aggregation in air pouch fluid collected from WT mice injected with either WT or iCCR−/− monocytes (×40 original magnification). (Di) Toluidine blue whole-mount staining of air pouch membrane of WT mice injected with either WT or iCCR−/− monocytes (×40 original magnification). (Dii) Graph summarizing cellular yields recovered from the air pouch membrane of WT mice injected with either WT or iCCR−/− monocytes (CD11c+MHCIIint DCs, CD11c+MHCII++ DCs, and CD11b+CD64+ macrophages) (n = 6); Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. (Diii) Graph summarizing yields of DCs and macrophages in the lymph nodes and spleen of WT mice injected with either WT or iCCR−/− monocytes. Student t test with Welch’s correction was performed to determine statistical significance, with a p value of 0.05 determined as significant. *p < 0.05, **p < 0.01.

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To attempt to analyze the effect of iCCR DC recruitment and differentiation in vivo, we used two distinct models of inflammation. For the first instance we examined granuloma formation in response to CFA injection. WT and iCCR−/− mice were injected s.c. with CFA, and the resulting granuloma was excised after 3 wk and analyzed using histology and flow cytometry to determine macrophage and DC numbers and the presence of proliferating clusters. H&E staining on excised granulomas revealed densely packed cells in iCCR−/− mice (Fig. 7Bi). However, further flow cytometric analysis revealed that the vast majority of these cells were neutrophils, with a complete absence of macrophages, DCs, and eosinophils even after 3 wk of sustained localized inflammation (Fig. 7Bii).

A clear problem with this model is that monocyte migration is severely impaired in both acute and chronic models of inflammation in iCCR−/− mice. We therefore next decided to bypass the requirement for iCCR-dependent migration by injecting sorted Ly6C++ monocytes from both WT and iCCR−/− bone marrow, along with GM-CSF, into the air pouch of WT animals to determine whether iCCR deficiency is associated with reduced differentiation and yields of macrophages and DCs. Three days after the monocyte injection, the fluid in the air pouch, together with the air pouch membrane, draining inguinal lymph node, and spleen were collected for analysis. Small undifferentiated CD45lowLy6C++CD11c were still detectable in the air pouch fluid of WT mice injected with WT and ICCR−/− (Fig. 7Ci). However, there was no difference in the number of recovered monocytes and no difference in the number of differentiated Ly6ClowCD11c+ cells in the air pouch fluid (Fig. 7Cii), suggesting that both WT and iCCR−/− monocytes were able to differentiate, although the efficiency of differentiation is essentially impossible to ascertain in this model. Furthermore, we could not detect clustered cells in the air pouch fluid (Fig. 7Ciii), suggesting that the clustering phenotype may not be detectable in the “fluid” environment of the air pouch. Whole-mount histology of the air pouch membrane also did not reveal any clustered cells (Fig. 7Di), although the increase in CD11c+MHCIIint and CD11c+MHCII++ cells observed in both WT and iCCR−/− monocyte-injected mice suggests monocytes have indeed managed to differentiate appropriately (Fig. 7Dii), Again, the efficiency of differentiation is impossible to determine in this model. Notably, a higher number of DCs was detected in the draining inguinal lymph node of both WT and iCCR−/− monocyte-injected mice (but not the spleen), suggesting that DC differentiation in both conditions was associated with an upregulation of CCR7 required for lymph node homing (Fig. 7Diii).

Overall, these experiments confirm expression of iCCRs on DCs in vivo but also indicate that demonstrating the in vivo relevance of our in vitro findings is likely to be impractical given the requirement for iCCRs for monocyte recruitment to tissue sites and the artifactual nature of the air pouch.

Macrophage and DC differentiation at inflamed sites is likely to be influenced by, and even dependent on, locally produced inflammatory mediators. One such inflammatory mediator is the cytokine GM-CSF (14), which is produced at inflamed sites and which, in vitro, is associated with differentiation of inflammatory-type macrophages and DCs. Also abundant at inflamed sites are chemokines, which are implicated in recruitment from the circulation, as well as intratissue movement, of leukocytes. We have previously shown that there is a residual level of DC production at inflamed sites in CCR2−/− mice, which have a profound impairment of monocyte recruitment (9). In mice lacking all the iCCRs, residual DC production ceases, suggesting a redundant role for the iCCRs in at least some aspects of DC production. In this study, we have tested the hypothesis that chemokine signaling in monocyte precursors is involved in macrophage and DC differentiation in conditions mimicking an inflamed environment.

Overall, our data reveal, to our knowledge, a novel role for inflammatory chemokine receptors in the development of GM-CSF–derived macrophages and DCs in vitro. Differentiating monocytes express CCR1, CCR2, and CCR5 in the first few days of culture, and chemokines secreted by these cells (e.g., CCL2, CCL5) bind to their cognate chemokine receptors, initiating a signaling cascade. The signal from the iCCRs is relayed to low-affinity integrins (inside-out signaling) (26), which change conformation to a high-affinity and high-avidity state and promote intercellular adhesion resulting in the clustering phenotype observed in vitro. These data are in keeping with previous studies reporting an association between chemokines and the regulation of monocyte integrin expression (27). Furthermore, our data show that chemokine-dependent integrin-based signaling leads to a reduction in Foxp1 expression and promotion of monocyte differentiation. Thus, iCCR signaling activates integrins on monocytes and stabilizes the formation of proliferation clusters allowing developing monocytes to stay in close contact and share signals. This ultimately promotes differentiation and proliferation of monocyte-derived DCs and macrophages. In the absence, or inhibition, of iCCRs, LFA-1 decreases in both expression and affinity, reducing cell-to-cell contact and inhibiting cluster formation, resulting in an accumulation of undifferentiated cells and an inability to phagocytose particulate Ag in mature DCs. It is our assumption that the ligand–receptor interactions that drive this clustering of DCs are paracrine in nature and may reflect a similar mechanism that controls DC/DC and T cell/DC interactions within peripheral tissues and lymph nodes (28, 29).

One of the striking features of this clustering phenotype is that it is redundantly controlled by chemokine receptors and is impaired only on deletion, or antagonism, of more than one receptor. Our interpretation of this is that during differentiation of recruited monocytes into macrophages and monocyte-derived DCs they express multiple iCCRs to allow them to respond to the varying chemokine landscapes that may be associated with different tissues and different infectious and inflammatory agents. Hence the redundancy seen in this crude in vitro context may not be reflected by redundancy in vivo but would allow for ligand–receptor–specific interactions in different tissues and inflammatory environments. In addition, however, this apparently redundant requirement for multiple chemokine receptor ligation may ensure that such clusters only form at sites of active inflammation in which ligands for all three receptors are coexpressed. The common downstream effect is clustering of differentiating monocytic cells to enhance differentiation.

Overall, therefore, our study reveals, to our knowledge, a novel role for chemokines and their receptors in regulating the clustering and differentiation of monocytic cells in vitro under surrogate inflammatory conditions. Crucially, our conclusions are derived entirely from in vitro experimentation. However, this is a necessary restriction of these studies, because mice with compound deletion in chemokine receptor genes have fundamental defects in cellular recruitment, and as shown in (Fig. 7, it is essentially impossible to disentangle recruitment defects from effects on intratissue monocytic differentiation. However, it is clear that clusters of monocytes, macrophages, and DCs do exist in vivo (3032), which, alongside clear evidence of expression of multiple iCCRs by DCs in vivo, suggests that what is observed in vitro is also likely to be applicable in vivo, at least in certain circumstances. In summary, therefore, our data highlight a potentially important role for chemokines and their receptors in cell–cell interactions, which enhances our understanding of their in vivo roles and which should be borne in mind in clinical trials using multiple chemokine receptor antagonists.

We acknowledge the assistance of the Institute of Infection, Immunity and Inflammation Flow Core Facility at the University of Glasgow.

This work was supported by a Wellcome Trust Investigator Award (217093/Z/19/Z to G.J.G.) and an MRC Programme Grant (MR/V010972/1 to G.J.G.).

The online version of this article contains supplemental material.

Abbreviations used in this article

     
  • bbOVA

    bead-bound OVA

  •  
  • BM

    bone marrow

  •  
  • DC

    dendritic cell

  •  
  • iCCR

    inflammatory CC-chemokine receptor

  •  
  • MFI

    mean fluorescent intensity

  •  
  • MHCI

    MHC class I

  •  
  • MHCII

    MHC class II

  •  
  • qPCR

    quantitative PCR

  •  
  • sOVA

    soluble OVA

  •  
  • WT

    wild-type

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The authors have no financial conflicts of interest.

This article is distributed under the terms of the CC BY 4.0 Unported license.

Supplementary data