C3a exerts multiple biologic functions through activation of its cognate C3a receptor. C3−/− and C3aR−/− mice have been instrumental in defining important roles of the C3a/C3aR axis in the regulation of acute and chronic inflammatory diseases, including ischemia/reperfusion injury, allergic asthma, autoimmune nephritis, and rheumatoid arthritis. Surprisingly little is known about C3aR expression and function in immune and stromal cells. To close this gap, we generated a floxed tandem-dye Tomato (tdTomato)–C3aR reporter knock-in mouse, which we used to monitor C3aR expression in cells residing in the lung, airways, lamina propria (LP) of the small intestine, brain, visceral adipose tissue, bone marrow (BM), spleen, and the circulation. We found a strong expression of tdTomato-C3aR in the brain, lung, LP, and visceral adipose tissue, whereas it was minor in the spleen, blood, BM, and the airways. Most macrophage and eosinophil populations were tdTomato-C3aR+. Interestingly, most tissue eosinophils and some macrophage populations expressed C3aR intracellularly. BM-derived dendritic cells (DCs), lung-resident cluster of differentiation (CD) 11b+ conventional DCs (cDCs) and monocyte-derived DCs, LP CD103+, and CD11b+ cDCs but not pulmonary CD103+ cDCs and splenic DCs were tdTomato-C3aR+. Surprisingly, neither BM, blood, lung neutrophils, nor mast cells expressed C3aR. Similarly, all lymphoid-derived cells were tdTomato-C3aR, except some LP-derived type 3 innate lymphoid cells. Pulmonary and LP-derived epithelial cells expressed at best minor levels of C3aR. In summary, we provide novel insights into the expression pattern of C3aR in mice. The floxed C3aR knock-in mouse will help to reliably track and conditionally delete C3aR expression in experimental models of inflammation.

Exogenous and endogenous threats can activate the complement cascade through canonical and noncanonical pathways (13). Both pathways drive the proteolytic cleavage of C3 into C3b and the generation of the anaphylatoxin C3a, which exerts a wide range of pro- and anti-inflammatory functions (4). Most C3a-induced functions require the binding to its cognate C3a receptor, which belongs to the large family of G protein–coupled receptors (5, 6). Binding of C3a to C3aR leads to rapid C3aR internalization that depends on phosphorylation of serine and threonine residues at the receptor C terminus (7, 8) and activation of pertussis toxin–sensitive G proteins (9).

Northern blot analysis revealed strong expression of the human C3aR in lung, spleen, small intestine, bone marrow (BM), low level expression in the brain, but absence in circulating leukocytes (5, 6). The strong pulmonary expression of C3aR was confirmed in mouse tissue. In contrast, C3aR expression was absent in the murine brain and in the spleen (10, 11). The strong pulmonary expression of C3aR was assigned to immune and stromal cells (12, 13). Among human pulmonary cells of myeloid origin, eosinophils were reported to express C3aR (5, 9). In asthmatic patients, C3aR expression has been reported on endothelial, epithelial, and smooth muscle cells (14). Furthermore, upregulation of C3aR in airway epithelial cells and smooth muscle cells was shown in asthmatics and in experimental allergy models (12, 14).

Binding studies, flow cytometric analyses and functional studies suggest C3aR expression in human neutrophils (15, 16), monocytes (16, 17), eosinophils (16, 18, 19), basophils (20, 21), and mast cells (MCs) (22, 23). The role of C3a as a chemoattractant for human neutrophils is controversial (4, 18). In mice, some reports suggested the expression of C3aR on neutrophils (24, 25), and BM-derived dendritic cells (BMDCs) (26). Reports regarding the expression of C3aR on human T cells are controversial. Earlier studies failed to detect C3aR surface expression in naive T cells (16, 27). More recently, low C3aR surface expression was described on naive human T cells, but C3aR was upregulated upon cluster of differentiation (CD) 3 and CD28 stimulation (28). Also, C3ar1 mRNA expression was reported in naive CD4+Foxp3+ natural regulatory T cells, which was enhanced in response to CD3 and CD28 stimulation (29). Another study found intracellular C3aR expression in naive human T cells and translocation to the cell surface following CD3 with or without CD46 stimulation (30). In mice, one study reported surface expression of C3aR in naive T cells (31).

At this point, most of the expression and functional data related to the C3a/C3aR axis have been obtained with human cells. Direct evidence for the expression of C3aR in murine immune and stromal cells under steady-state or upon inflammatory conditions is limited, which is mainly due to the lack of C3aR-specific mAbs. Several studies used polyclonal or poorly characterized mAbs raised against C3aR (13, 3234). In this study, we report the generation of a floxed tandem-dye Tomato (tdTomato)–C3aR reporter knock-in mouse. The reporter strain contains the coding sequence for a tdTomato-C3aR self-processing polyprotein, flanked by two loxP sites. Using this mouse, we evaluated C3aR expression in myeloid and lymphoid cells from the circulation and several tissues, including the lung, airways, lamina propria (LP) of the small intestine, spleen, visceral adipose tissue (VAT), and the brain. We identified various tdTomato-C3aR+ cell populations. They were evaluated for C3aR surface expression using commercially available anti-C3aR Abs, tested for their specificity using cells from C3aR-deficient mice. This approach provided detailed insights into the C3aR expression pattern in immune and stromal cells under steady-state conditions.

Monoclonal allophycocyanin-Cy7–labeled Ab against CD11b (M1/70), PerCP-Cy5.5–labeled Ab against CD8a (53-6.7), allophycocyanin or V450-labeled Ab against Ly6G (1A8), V450-labeled Ab against CD45R (RA3-6B2), FITC-labeled Ab against CD90.2 (30-H12), allophycocyanin-H7–labeled Ab against CD19 (1D3), and Brilliant Violet (BV)421–labeled Ab against Siglec-F (E50-2440) and CD43 (1G10) were all purchased from BD Biosciences. PerCP-Cy5.5–labeled Ab against Ly6C (RB6-8C5); allophycocyanin-labeled Ab against CD115 (AFS98), CD11c (N418), NK1.1 (PK136), CD4 (GK1.5), and CD11b (M1/70); PE-Cy7–labeled Ab against CD25 (PC61.5) and CD4 (RM4-5); eFluor (eF)450–labeled Ab against CD19 (1D3), CD3e (145-2C11), and CD49b (DX5); Alexa Fluor (AF)488–labeled Ab against CD3 (17A2); allophycocyanin-eF780–labeled Ab against MHC class II (MHCII) (M5/144.15.2); BV510-labeled Ab against CD11b (M1/70); BV711-labeled Ab against CD64 (X54-5/7.1); BV421-labeled Ab against F4/80 (BM8); and PerCP-Cy5.5–labeled Ab against CD103 (2E7) were all purchased from BioLegend. PerCP-Cy5.5–labeled Ab against CD3 (17A2), CD5 (53-73), CD27 (LG.7F9), NK1.1 (PK136), TCR-β (H57-597), and CD11b (M1/70); allophycocyanin-eF780–labeled Ab against CD11c (N418), B220 (RA3-6B2), and CD49b (DX5); eF450-labeled Ab against CD25 (PC61.5) and CD317 (ebio129c); FITC-labeled Ab against CD11c (N418); AF700-labeled Ab against CD11b (M1/70); PE-Cy5–labeled Ab against CD127 (A7R34); and PE-Cy7 IgM (11/41) were purchased from eBioscience (Affymetrix). For surface and intracellular staining, a rat unlabeled C3aR-specific Ab (14D4) was purchased from Hycult Biotech. Binding of this Ab was detected using a secondary allophycocyanin-labeled F(ab′)2 anti-rat Ab (Cell Signaling Technology). Additionally, we used a goat unlabeled C3aR-specific Ab (D20) from Santa Cruz Biotechnology, the binding of which was detected using a secondary allophycocyanin-labeled F(ab′)2 anti-goat Ab (Santa Cruz Biotechnology).

RBC lysis (RBCL) buffer was prepared by using 155 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA (all from Sigma-Aldrich). BSA and an LP dissociation kit were purchased from Miltenyi Biotec. Recombinant murine GM-CSF was from PeproTech. DNase I for total RNA isolation was from Fermentas. Liberase TL was from Roche Diagnostics, and DNase I for cell isolation was from Sigma-Aldrich. FBS, RPMI 1640 medium, and HBSS were all from PAA Laboratories. Dulbecco’s PBS (D-PBS), l-glutamine, penicillin, and streptomycin were all from Life Technologies.

C57BL/6 wild-type (WT) mice were obtained from Janvier. BALB/c WT mice were purchased from the Charles River Laboratories. C3ar1−/−, C3ar1−/−/C5ar1−/−, and tdTomato-C3ar1fl/fl mice in BALB/c background were bred in the animal facility of the University of Lübeck as described (35). All animals were used at 8–12 wk of age and handled in accordance with the appropriate institutional and national guidelines. Animals were used for organ removal according to protocols approved by the local authorities of the Animal Care and Use Committee (Ministerium für Landwirtschaft, Energiewende, Umwelt und ländliche Räume, Kiel, Germany). The experimental allergic asthma study and the i.p. injection of thioglycollate were reviewed and approved by the Schleswig-Holstein state authorities (numbers V312-7224.122-39 [37-2/13] and V242-81505/2016 [19-2/2017], respectively). All experiments were performed by certified personnel.

Experimental allergic asthma was induced as previously described (36). Briefly, BALB/c WT and tdTomato-C3ar1fl/fl mice were anesthetized by i.p. injection with ketamine (Ketavet; Pfizer) and xylazine (Rompun; Bayer) and sensitized intratracheally with 100 μg of house dust mite (HDM) extract (Greer Laboratories, lot not. 262538) in 50 μl of PBS on days 0, 7, 14, and 21. Seventy-two hours after the final intratracheal HDM challenge on day 21, bronchoalveolar lavage (BAL) fluid and lung tissue were harvested for tdTomato-C3ar and C3aR expression.

BALB/c WT and tdTomato-C3ar1fl/fl mice were injected i.p. with 1 ml of 3% thioglycollate medium (BD Biosciences). Seventy-two hours later, peritoneal (PE) cell exudates were collected using 5 ml of ice-cold PBS. Subsequently, cells were identified as F4/80+ cells by flow cytometry and used for the assessment of C3a-mediated mobilization of intracellular Ca2+.

The tdTomato-C3aR knock-in mice were generated by gene targeting. A scheme of the targeting strategy is shown in Fig. 1A. The targeting vector, the embryonic stem cells, and the targeted mice were generated by Ozgene (Perth, Australia). Our strategy was to insert tdTomato in frame with the coding sequence of C3ar1 and simultaneously flank the tdTomato-C3ar1 cassette with two loci of loxP sites. A construct that was the precursor to the final targeting construct was generated by the sequential cloning of six cloned fragments into a final recipient plasmid housing a neomycin selectable marker cassette. Primers used to amplify the fragments also encoded the restriction enzyme sites required for the assembly of the fragments generated. All fragments were housed in the Surf2 vector backbone (Ozgene). The first fragment was a 3.1-kb fragment encoding the 3′ half of the 3′ homology arm. This fragment was amplified from BALB/c genomic DNA using primers 1471_46 (5′-CTAAACTGTGCATCCTAGGTTCAACTTC-3′) and 1471_56 (5′-TAAGCATTGGTAATTCGAAGCAGGCTGTAAAGTCTTGGCATGATGTG-3′). The second fragment was a 3.1-kb fragment encoding the 5′ half of the 3′ homology arm. This fragment was amplified from BALB/c genomic DNA using primers 1471_47 (5′-TAAGCATTGGTAATTCGAATGACCGTCACATAACCCAGGGATGGA-3′) and 1471_57 (5′-CTAATTAATTAAGGGAACCTAAGCAGAGTTCTCA-3′). The third fragment was a 1.4-kb fragment encoding the floxed region of exon 2 of C3ar1. This fragment was amplified from BALB/c genomic DNA using primers 1471_42 (5′-TAAGCATTGGTAATTAATTAACAATTGATAACTTCGTATAGCATACATTATACGAAGTTATAGGGCCACATCTTCACACATCTGTACT-3′) and 1471_52 (5′-CTAAGCGGCCGCCACCATGGAGTCTTTCGATGCTGACAC-3′) where primer 1471_42 also incorporated the loxP sequence. The fourth fragment was a 0.6-kb fragment encoding an internal ribosomal entry site (IRES). This fragment was obtained from one of Ozgene’s proprietary cloning vectors. The fifth fragment was a 1.4-kb fragment encoding tdTomato and was obtained from one of Ozgene’s proprietary cloning vectors. The sixth fragment was a 3.0-kb fragment encoding the 5′ homology arm. This fragment was amplified from BALB/c genomic DNA using primers 1471_45 (5′-CTAACTGATAGACAGTTTCAGGTCTATGC-3′) and 1471_55 (5′-TAAGCATTGGTAACCAAATGTCCCAGCTTCCATTTA-3′). The vector backbone into which this amplicon was introduced contained the neomycin selection cassette. The six fragments were assembled sequentially as follows: fragment 1 was excised by digestion with BstBI and ligated into fragment 2 that had been digested with the same enzyme. Combined fragments 1 and 2 were excised from the resulting construct by digestion with PacI and ligated into fragment 3, which had been digested with the same enzyme. Combined fragments 1, 2, and 3 were excised from the resulting construct by digestion with NotI and ligated into fragment 4, which had been digested with the same enzyme. Combined fragments 1, 2, 3, and 4 were excised from the resulting construct by digestion with FseI and ligated into fragment 5, which had been digested with the same enzyme. Combined fragments 1, 2, 3, 4, and 5 were excised from the resulting construct by digestion with AscI and ligated into fragment 6, which had been digested with the same enzyme, resulting in the vector 1471_pTV. To construct the final targeting vector, 1471_pTV was digested with MfeI and KpnI to excise the region encoding tdTomato_IRES_exon2_loxP. A synthetic fragment (Blue Heron Biotech) encoding tdTomato_P2A_exon2_loxP digested with MfeI and KpnI was ligated into 1741_pTV in place of the excised fragment. We replaced the IRES by the porcine teschovirus-1 2A (P2A) because of its short length, high cleavage efficiency, and the stoichiometric expression of multiple proteins flanking the 2A peptide (37, 38). The resulting targeting construct encoded a 6.2-kb 5′ homology arm and a 3.0-kb 3′ homology arm flanking the floxed coding region of exon 2 where tdTomato_P2A had been knocked in at the ATG of exon 2.

For genotyping, we used ear biopsies. Extraction of DNA from tissue was performed using the KAPAExpress extraction kit from Peqlab Biotechnologie following the manufacturer’s instructions. To amplify the different DNA fragments, we used following primers: GK342, 5′-AACAACAGAAGTAGGGAGGTGTAA-3′; GK45, 5′-TCCCAATAGACAAGTGAGACCAA-3′ (both from Eurofins Scientific). The PCRs were run using the following conditions: 95°C for 3 min, followed by 35 cycles at 95°C for 15 s, 62°C for 15 s, and 72°C for 60 s, followed by 72°C for 120 s. Then, the samples were loaded to a 1.0% sodium borate agarose gel. Amplification products were detected by GelRed staining (Biotrend).

Mice were killed by cervical dislocation under anesthesia or after CO2 exposure and the heart was perfused with 10 ml of ice-cold PBS before organ removal. For BM preparation, femurs and tibias were removed, placed in PBS on ice, and subsequently flushed with PBS. RBCs were removed by incubating the cells in RBCL buffer for 3 min. The reaction was terminated by addition of a large volume of PBS. BMDCs were generated as described (39). Briefly, BM cells were differentiated for 9 d with GM-CSF (20 ng/ml); BMDCs were defined as CD11c+CD11b+MHCIIhiCD115 cells. To obtain BM-derived plasmacytoid DCs (pDCs), BM cells were differentiated for 9 d in the presence of Flt3L (200 ng/ml). To generate BM-derived macrophages (BMMs), we cultured BM cells for 6 d in the presence of L929 cell–conditioned medium (40). Alternatively, BM cells were cultured in GM-CSF (20 ng/ml) for 9 d and identified as CD11c+CD11b+CD115+MHClo BMMs as described (41). For pulmonary cell preparation, lungs were harvested, digested using 0.25 mg/ml Liberase TL and 0.5 mg/ml DNase I in pure RPMI 1640 medium for 45 min at 37°C, and single-cell suspensions were prepared as described (42). For VAT cell preparations, perigonadal fat tissue was harvested, taking care not to remove the gonads, and minced into small pieces. The tissue was digested using 0.25 mg/ml Liberase TL and 0.5 mg/ml DNase I in pure RPMI 1640 medium for 45 min at 37°C. The cell suspension was filtered through a nylon mesh (40 μm) and washed with 10 ml of RPMI 1640 medium complemented with 10% FCS containing 0.5 mg/ml DNase I. Cells were centrifugated at 350 × g for 10 min at 4°C, resuspended in PBS, transferred into a fresh 15-ml tube, and washed once more. BAL fluid samples were obtained as described (39). To collect cells from the PE cavity, mice were lavaged with 5 ml of ice-cold PBS. Collected cells were washed once with PBS. RBCs were removed by incubating the cells in RBCL buffer for 3 min and then washed with PBS as described (42). Blood was collected by cardiac puncture and immediately transferred into tubes containing 10 mM EDTA to prevent coagulation. Diluted blood samples were incubated several times in RBCL buffer to remove RBCs and then washed with PBS as described (42). Isolation of cells from the spleen, mediastinal lymph nodes, and the Peyer’s patches from the small intestine was performed by mechanical disruption using a cell strainer (40 μm nylon; BD Biosciences) and a plunger of a 5-ml syringe (BD Biosciences). The cell strainer was flushed three times with 5 ml of PBS. Cells were then incubated in RBCL buffer for 3 min and finally washed with PBS. LP cell suspension was obtained using an LP dissociation kit (Miltenyi Biotec), following the manufacturer’s recommendation. Briefly, the small intestine was removed from the mice and quickly freshly collected and cleared from feces, residual fat, and Peyer’s patches. Then, they were cut longitudinally and washed once with HBSS supplemented with EDTA (5 mM) and once with HBSS only. The tissue was digested using the manufacturer’s enzyme mixture in HBSS with Ca2+ and Mg2+ for 30 min at 37°C and homogenized with the gentleMACS dissociator (Miltenyi Biotec). After the last washing step, the cell number was determined using a Neubauer chamber, after which the cells were resuspended in PBS containing 1% BSA. The viability of the collected cells was determined by trypan blue exclusion or a Live/Dead cell viability assay (eBioscience). Small intestine epithelial cells were obtained by incubating freshly isolated tissue, cleared from feces, residual fat, and Peyer’s patches, in HBSS supplemented with EDTA (5 mM) for 30 min. Cell suspension was cleared from clumps through a 100-μm cell strainer before blocking. Brain tissue was collected, sliced into pieces, and digested with Liberase TL (0.1 mg/ml) and DNase I (0.1 mg/ml) for 30 min at 37°C. Brain homogenates were passed through a 40-μm cell strainer and the cell suspension was washed and fractionated on a 30–75% discontinuous Percoll gradient (GE Healthcare) for 30 min at 1350 × g without brake. The interphase-containing cells were collected, washed with D-PBS, and the cellular composition was analyzed.

Phenotypic characterization and determination of tdTomato-C3aR expression of tissue and circulating cells was performed using a BD FACSAria III cell sorter (BD Biosciences) and a MoFlo Legacy (Beckman Coulter). The markers and gating strategies used to identify the myeloid, lymphoid, and stromal cells in the different tissues and the circulation are summarized in Supplemental Fig. 3C–H and in the text. Flow cytometric data were analyzed using FlowJo 10 (Tree Star).

All cells that stained positive for tdTomato-C3aR were also assessed for C3aR expression on the cell surface using two different C3aR-specific Abs (clones 14D4 and D20). In some cells, we also determined intracellular expression of C3aR. For surface staining with the rat C3aR-specific mAb 14D4 (Hycult Biotech), cells were first incubated with PBS supplemented with 20% FCS (PBS20) for 30 min at 4°C, after which the cells were stained with the Ab in PBS20 for 30 min at 4°C. After washing with PBS20, cells were stained with an F(ab′)2 anti-rat allophycocyanin Ab (Cell Signaling Technology). After three washes with PBS20, cells were resuspended in PBS/BSA containing Fc Block (eBioscience) and stained for cell-specific markers. The surface staining with a goat polyclonal C3aR-specific Ab (D20) was done in PBS/BSA buffer, together with BMM-specific Abs (F4/80 and CD11b) in the presence of Fc Block. After washing with PBS/BSA, cells were stained with an F(ab′)2 anti-goat allophycocyanin Ab (Santa Cruz Biotechnology). For intracellular staining, cells from WT, C3ar1−/− or tdTomato-C3ar1fl/fl mice were first fixed in 1.5% paraformaldehyde, then resuspended in a saponin (0.2%) buffer containg 20% FCS instead of PBS20. After 30 min, cells were stained with the 14D14 Ab in saponin (0.2%) buffer containg 20% FCS instead of PBS20. After washing with PBS20, cells were stained with an F(ab′)2 anti-rat allophycocyanin Ab (Cell Signaling Technology). After three washes with PBS20, cells were resuspended in PBS/BSA containing Fc Block (eBioscience) and stained for cell-specific markers. C3aR surface and intracellular expression was measured by flow cytometry.

The spleen was harvested and dissociated by mechanical disaggregation. After RBCL, cells were counted and whole spleen cells were used for in vitro T cell activation. To stimulate CD4+ T cells, a 96-well cell culture plate (Greiner Bio-One International) was coated with unconjugated anti-CD3 (17A2; eBioscience) in 150 μl of PBS at a final concentration of 2 μg/ml. The plate was kept at 37°C, 5% CO2 for at least 2 h before discarding the coating solution and adding the cell suspension. T cells were cultured in presence of 10 μg/ml anti-CD28 (37.51; eBioscience) for 5 d in complete RPMI 1640 culture medium supplemented with 10% FBS, 2 mM l-glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. Cells were subsequently analyzed by flow cytometry for tdTomato-C3aR expression.

T and B cells as well as neutrophils from different compartments were sorted using a BD FACSAria III or a MoFlo Legacy cell sorter. Total RNA was isolated using TRIzol reagent (Thermo Fisher Scientific) according to the manufacturer’s instructions. After DNAse I (Fermentas) treatment of the RNA, reverse transcription reaction was performed using a RevertAid first-strand cDNA synthesis kit (Thermo Fisher Scientific). PCR was done using the following primers: β-actin, 5′-GCACCACACCTTCTACAATGAG-3′ and 5′-AAATAGCACAGCCTGGATAGCAAC-3′, C3ar1, 5′-TCGATGCTGACACCAATTCAA-3′ and 5′-TCCCAATAGACAAGTGAGACCAA-3′. The temperature profiles were 95°C for 3 min, followed by 34 cycles at 95°C for 30 s, 57°C for both targets for 30 s, and 72°C for 30 s. Amplification products were separated using a 2% sodium borate agarose gel. Amplification products were detected by GelRed staining (Biotrend).

WT and tdTomato-C3ar1fl/fl mice were sacrificed by isoflurane inhalation (Baxter), and blood was removed from pulmonary vasculature by perfusion with 5 ml of 37°C HEPES-Ringer buffer (10 mM HEPES, 136.4 mM NaCl, 5.6 mM KCl, 1 mM MgCl2, 2.2 mM, CaCl2, 11 mM glucose [pH 7.4]) containing 300 μl of heparin-natrium (25000; Ratiopharm) via the right ventricle. Then, the airways were filled via the cannulated trachea with 3% low melting point agarose (Bio-Rad Laboratories) dissolved in HEPES-Ringer buffer. The lungs were removed en bloc and transferred into ice-cold HEPES-Ringer buffer to solidify the agarose. The lungs were cut as precision-cut lung slices into 300-μm-thick slices using a Vibratome (VT1200S; Leica). For fixation and freezing, slices were placed in 1% paraformaldehyde (Sigma-Aldrich), then washed three times in D-PBS and incubated overnight with 20% d(+)-saccharose (Carl Roth) and frozen at −20°C until further use.

Immunohistochemical stainings were performed in 24-well plates (tissue culture OrPlate; Orange Scientific) in the dark to protect the fluorophores from light. Precision-cut lung slices were defrosted at room temperature on a shaker and washed three times in 1 ml of TBS for 10 min. Primary antibodies were diluted in 500 μl of TBS and incubated on a shaker overnight at room temperature. The next day, the slices were washed three times in TBS for 10 min. The slices were transferred to an object slide (Superfrost; Menzel Gläser), dried carefully, and coverslipped with Mowiol as mounting medium (Mowiol 4-88 [Höchst], 200 mM Tris buffer [pH 8.5; Roth], glycerin [Merck]). Images were acquired with an LSM 710 confocal laser-scanning microscope (Carl Zeiss) with a ×20/0.5 M27 objective and an immersion oil objective ×40/1.30 oil differential interference contrast M27 (Carl Zeiss). Emitted light was detected by three wavelength-separated photomultiplier tubes at 505–534, 555–593, and 657–723 nm. Zen 2011 (Carl Zeiss) was used as acquisition software. Image processing was conducted using Imaris software (Bitplane). mAbs used for staining were AF647-labeled Ab against Siglec-F (E50-2440) from BD Biosciences, AF488-labeled Ab against CD11c (N418), and AF647-labeled Ab against Clara cell protein 10 (CC10).

Imaging of fixed lung slices was performed using the TriM Scope II multiphoton microscope (LaVision BioTec) equipped with a XLUMPLFL ×20 W/0.95 water immersion objective (Olympus). Images were acquired at 740 and 1100 nm performed by a Mai Tai HP (Spectra-Physics) and an InSight DeepSee (Spectra-Physics). Emitted light was detected by three wavelength-separated photomultiplier tubes (Hamamatsu) at 435–495, 495–560, and >560 nm. Imspector Pro (LaVision BioTec) was used as acquisition software. Image processing was conducted using Imaris software (Bitplane) and ImageJ (National Institutes of Health).

Sorted BM neutrophil and PE macrophage lysates from WT, tdTomato-C3ar1fl/fl, and C3ar1−/−/C5ar1−/− animals were obtained by resuspending the cells in Laemmli buffer preheated at 95°C. Lysates were homogenized through a needle and heated at 95°C before loading. Cell lysates were separated by SDS-PAGE according to standard procedures using a Mini-Protean TGX precast gradient gel 4–12% (Bio-Rad Laboratories). Proteins were transferred onto Trans-Blot nitrocellulose membrane (Bio-Rad Laboratories) using a Trans-Blot SD system (Bio-Rad Laboratories). Western blot analysis was performed according to standard procedures. Briefly, after blocking, the membrane was incubated overnight with a polyclonal Ab raised against C3aR (clone D20; Santa Cruz Biotechnology) in TBS plus 5% Blotto low-fat dry milk (Rockland Immunochemicals) plus 0.1% Tween 20. After washing (TBS plus 0.1% Tween 20), the membrane was incubated with an anti-goat IgG-HRP (1:2500 in TBS plus 5% Rockland Immunochemicals Blotto low-fat dry milk plus 0.1% Tween 20; Santa Cruz Biotechnology) for 1 h at room temperature. After washing, detection was performed using the Immun-Star WesternC kit (Bio-Rad Laboratories) and Fusion SL (Vilbert Lourmat).

PE cells (1 × 107 cells/ml) from WT or tdTomato-C3ar1fl/fl mice were resuspended in PBS and allowed to rest for 5 min at 37°C. Cells were left untreated or stimulated either with 1, 10, and 100 nM C3a (Hycult Biotech) for 3 min at 37°C, or with 10 nM for 1, 3, and 9 min at 37°C. The reaction was stopped by addition of paraformaldehyde (1.5% final concentration) and cells were immediately placed on ice. After 30 min fixation, cells were additionally blocked for 30 min with PBS supplemented with PBS20, and then stained first with anti-C3aR mAb (clone 14D4; Hycult) in PBS20 for 30 min at 4°C. After washing with PBS20, cells were stained with an F(ab′)2 anti-rat allophycocyanin Ab (Cell Signaling Technology). After three washes with PBS20, they were resuspended in PBS/BSA containing Fc Block (eBioscience) and stained with an F4/80-specific Ab. Expression of C3aR in F4/80+ cells was determined by flow cytometry.

The C3a-induced increase in intracellular Ca2+ concentration ([Ca2+]i) was determined, as described (42, 43), using thioglycollate-elicited F4/80+ PE macrophages. Briefly, cells were collected in 5 ml of PBS and loaded with 5 mM of the Ca2+-sensitive fluorophore Fluo-4AM (Molecular Probes) according to the manufacturer’s recommendations. Cells were stained with F4/80 for 20 min in PBS/BSA. F4/80hi cells were then analyzed on the LSR II flow cytometer. The background signal was recorded for 30 s. Then, C3a (37 nM) was added, and recording continued for another 90 s. The increase in [Ca2+]i was calculated by assessment of the maximal Ca2+ peak using the kinetic plug-in tool of the FlowJo software (version 9; Tree Star, Ashland, OR). The mean background signal recorded during the 30 s was subtracted from the Ca2+ peak signal to determine the final change in relative mean fluorescence intensity (ΔMFI). Alternatively, thioglycollate-elicited PE macrophages were allowed to adhere on glass coverslips for 2 h before loading with Fluo-4AM as indicated. Glass slides were washed and the assay was performed in PBS. Images were obtained using the Olympus FV 1000 confocal microscope (Olympus, Germany) with a ×40 objective. Image analysis and capturing was performed using the FluoView 2.1c software.

Statistical analysis was performed using GraphPad Prism (version 5.0c; GraphPad Software). Statistical differences between groups were assessed by Student t test. Comparisons involving multiple groups were first analyzed by ANOVA. In case of significant differences (p < 0.05), all groups were post hoc compared pairwise by a Tukey test. A p value <0.05 was considered significant (*p < 0.05, **p < 0.01, ***p < 0.001).

We aimed at generating a tdTomato-C3aR reporter knock-in mouse that allows for conditional deletion of the C3ar1 gene. Therefore, we cloned a tdTomato cassette directly upstream of the 5′ end of exon 2 of the C3ar1 gene. To allow for conditional deletion of the C3ar1 gene, we placed two loxP sites upstream of the tdTomato coding sequence and downstream of exon 2 of the C3ar1 gene (Fig. 1A). We obtained several embryonic stem cells that incorporated tdTomato and the two loxP sites into the C3ar1 gene locus, one of which was successfully used to produce heterozygous (tdTomato-C3ar1fl/+) and homozygous (tdTomato-C3ar1fl/fl) floxed tdTomato-C3aR knock-in mice (Fig. 1B). A first screen of tdTomato-C3aR tissue expression revealed a strong expression in the brain, lung, LP of the small intestine, and VAT (Fig. 1C). In contrast, the expression of tdTomato-C3aR in the spleen, blood, BM, and the airways was at best minor (Fig. 1D).

FIGURE 1.

Generation and initial characterization of the floxed tdTomato-C3aR knock-in mouse. (A) Schematic of the gene-targeting strategy. WT exon 2 of C3ar1 was replaced with a cassette encoding a fusion protein comprising tdTomato-C3aR flanked with loxP sites by homologous recombination. (B) PCR-based phenotyping of WT, heterozygous tdTomato-C3ar1fl/+, and homozygous tdTomato-C3ar1fl/fl mice. The primer combination GK342:GK45 amplifies a 400-bp DNA fragment in BALB/c WT and tdTomato-C3aRfl/+ mice and a 2000-bp DNA fragment in tdTomato-C3ar1fl/fl mice. The arrows point toward the amplified PCR fragments. (C and D) Flow cytometric analysis of the tdTomato signal in (C) brain, lung, VAT, and LP of the small intestine; and (D) spleen, BAL, blood, and BM from WT and tdTomato-C3ar1fl/fl mice.

FIGURE 1.

Generation and initial characterization of the floxed tdTomato-C3aR knock-in mouse. (A) Schematic of the gene-targeting strategy. WT exon 2 of C3ar1 was replaced with a cassette encoding a fusion protein comprising tdTomato-C3aR flanked with loxP sites by homologous recombination. (B) PCR-based phenotyping of WT, heterozygous tdTomato-C3ar1fl/+, and homozygous tdTomato-C3ar1fl/fl mice. The primer combination GK342:GK45 amplifies a 400-bp DNA fragment in BALB/c WT and tdTomato-C3aRfl/+ mice and a 2000-bp DNA fragment in tdTomato-C3ar1fl/fl mice. The arrows point toward the amplified PCR fragments. (C and D) Flow cytometric analysis of the tdTomato signal in (C) brain, lung, VAT, and LP of the small intestine; and (D) spleen, BAL, blood, and BM from WT and tdTomato-C3ar1fl/fl mice.

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In previous studies, C3aR expression has been found in Gr-1+ cells that have been considered as neutrophils (32, 44). However, Gr-1 is not only expressed on neutrophils but broadly on other immune cells, including eosinophils, monocytes, DCs, T cells, NK cells, and NKT cells. High expression of Ly6G can be used to reliably identify neutrophils (45). Therefore, we first assessed the expression of tdTomato-C3aR in Ly6Ghi neutrophils from BM, blood, and lung tissue using tdTomato as a surrogate for C3aR expression. Surprisingly, neither BM, blood, nor lung neutrophils expressed C3aR (Fig. 2A). Also, C3aR surface staining of BALB/c or C57BL/6 BM neutrophils with C3aR-specific mAb 14D4 were negative (Supplemental Fig. 1A). Previously, it was reported that Gr-1+ cells express C3aR based on Western immunoblot analysis and by flow cytometry (24). Re-evaluating the Ab used in that study (D20), we found no C3aR surface staining in BM neutrophils (Supplemental Fig. 1A) but a band in Western blot analysis that also appeared in BM neutrophils isolated from C3ar1−/−C5ar1−/− mice (Supplemental Fig. 1B). To exclude a neutrophil-specific proteolysis of the protein, we also determined the mRNA expression of C3ar1 in WT and tdTomato-C3ar1fl/fl mice. We found no C3ar1 transcripts in Ly6G+-sorted blood- and lung-derived neutrophils and a very faint band in Ly6G+ BM neutrophils. In contrast, C3ar1 mRNA was strongly expressed in GM-CSF–differentiated BMDCs/BMMs used as positive control (Supplemental Fig. 1C). These data demonstrate that mouse neutrophils in the circulation and in pulmonary tissue do not express C3aR, whereas C3aR is at best marginally expressed in BM neutrophils at the mRNA level.

FIGURE 2.

Low level of tdTomato-C3aR expression in airway and tissue neutrophils during the effector phase of HDM-driven allergic asthma. Flow cytometric assessment of tdTomato-C3aR expression in different neutrophil populations is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (A) Flow cytometric analysis of the tdTomato signal in Ly6G+ neutrophils from BM, blood, and lung. (B) Strong influx of neutrophils in the airways during the allergic effector phase. BAL fluid cells were first gated as Siglec-FCD11c cells (left histogram). Within the population, neutrophils were identified as Ly6G+ cells (right histogram). The graph on the right shows the number of airway neutrophils in the indicated groups. Values shown are the mean ± SEM; n = 4 PBS groups and n = 6 HDM groups. Statistical differences between groups were assessed by a Student t test. (C) tdTomato-C3aR (left) and C3aR surface (right) expression in BAL neutrophils. Values shown are the mean ± SEM; n = 3 for tdTomato-C3aR expression in WT and tdTomato-C3ar1fl/fl mice; n = 4–6 for C3aR surface expression in WT (open symbols) and tdTomato-C3ar1fl/fl (closed symbols) mice treated with PBS (circles) or HDM (squares). Statistical differences between groups were assessed by Student t test. (D) tdTomato-C3aR and C3aR surface expression in lung tissue neutrophils. Shown is the gating strategy to identify Ly6G+ neutrophils in lung tissue (two histograms upper row), tdTomato-C3aR expression (lower left histogram and graph), and surface C3aR expression (lower right histogrgam and graph). Values shown are the mean ± SEM; n = 3 for tdTomato expression in WT and tdTomato-C3ar1fl/fl mice; n = 4–6 for C3aR surface expression in WT (open symbols) and tdTomato-C3ar1fl/fl (closed symbols) mice treated with PBS (circles) or HDM (squares). Statistical differences between groups were assessed by Student t test. *p < 0.05, **p < 0.01.

FIGURE 2.

Low level of tdTomato-C3aR expression in airway and tissue neutrophils during the effector phase of HDM-driven allergic asthma. Flow cytometric assessment of tdTomato-C3aR expression in different neutrophil populations is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (A) Flow cytometric analysis of the tdTomato signal in Ly6G+ neutrophils from BM, blood, and lung. (B) Strong influx of neutrophils in the airways during the allergic effector phase. BAL fluid cells were first gated as Siglec-FCD11c cells (left histogram). Within the population, neutrophils were identified as Ly6G+ cells (right histogram). The graph on the right shows the number of airway neutrophils in the indicated groups. Values shown are the mean ± SEM; n = 4 PBS groups and n = 6 HDM groups. Statistical differences between groups were assessed by a Student t test. (C) tdTomato-C3aR (left) and C3aR surface (right) expression in BAL neutrophils. Values shown are the mean ± SEM; n = 3 for tdTomato-C3aR expression in WT and tdTomato-C3ar1fl/fl mice; n = 4–6 for C3aR surface expression in WT (open symbols) and tdTomato-C3ar1fl/fl (closed symbols) mice treated with PBS (circles) or HDM (squares). Statistical differences between groups were assessed by Student t test. (D) tdTomato-C3aR and C3aR surface expression in lung tissue neutrophils. Shown is the gating strategy to identify Ly6G+ neutrophils in lung tissue (two histograms upper row), tdTomato-C3aR expression (lower left histogram and graph), and surface C3aR expression (lower right histogrgam and graph). Values shown are the mean ± SEM; n = 3 for tdTomato expression in WT and tdTomato-C3ar1fl/fl mice; n = 4–6 for C3aR surface expression in WT (open symbols) and tdTomato-C3ar1fl/fl (closed symbols) mice treated with PBS (circles) or HDM (squares). Statistical differences between groups were assessed by Student t test. *p < 0.05, **p < 0.01.

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A recent study showed that C3aR in neutrophils is upregulated under proinflammatory conditions (46). The C3a/C3aR plays an important role in the effector phase of experimental allergic asthma (14, 47, 48). Thus, we analyzed dTomato-C3aR expression in airway and lung neutrophils in HDM-driven allergic asthma. Seventy-two hours after the final HDM challenge, we observed a marked and significant increase in neutrophil numbers in the airways of HDM-treated mice as compared with PBS controls (Fig. 2B). Such neutrophils showed a slight but significant increase in the tdTomato-C3aR signal when compared with WT mice. However, we found no C3aR surface expression in WT or tdTomato-C3aR neutrophils (Fig. 2C). Similarly, lung neutrophils showed an increase in the tdTomato-C3aR signal, but no C3aR surface expression in WT or tdTomato-C3aR neutrophils (Fig. 2D). Thus, airway and lung tissue neutrophils upregulate C3aR expression during the effector phase of HDM-driven allergic asthma.

F4/80 is considered a key lineage marker for macrophages (49). Alveolar and tissue-associated alveolar macrophages (AMs) are characterized by the expression of Siglec-F (50). Surprisingly, airway and tissue-associated AMs did not express the tdTomato signal (Supplemental Fig. 2A, 2B). Furthermore, we found no tdTomato signal in Siglec-F+CD11c+ AMs by confocal microscopy (Supplemental Fig. 2C). In line with this finding, we found no C3ar1 mRNA expression in AMs as determined by RT-PCR (Supplemental Fig. 2D). In contrast, F4/80+Siglec-F macrophages differentiated with M-CSF (Fig. 3A) or GM-CSF from BM cells (Fig. 3B) or PE macrophages (Fig. 3C) as well as macrophages from the LP of the small intestine (Fig. 3D) or from VAT (Fig. 3E) showed a strong tdTomato signal. Similarly, brain macrophages (Fig. 3F) were homogeneously positive for tdTomato-C3aR. In contrast, M-CSF–derived BMMs (Fig. 3A), VAT macrophages (Fig. 3E), and microglia cells can be divided into C3aR+ and C3aR subpopulations.

FIGURE 3.

Siglec-FF4/80+ macrophages express dTomato-C3aR and C3aR. Flow cytometric assessment of tdTomato-C3aR expression in different macrophage populations. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (AF) tdTomato-C3aR and C3aR expression in (A) M-CSF–derived F4/80+CD11b+ BM macrophages (M-CSF BMM); (B) GM-CSF–derived BM macrophage population (GM-CSF BMM) that was identified as CD115+MHCIIlo cells within the CD11c+CD11b+ cell population; (C) F4/80+CD11b+ PE macrophages; (D) LineageCD11c+CD11b+MHCII+F4/80+ macrophages from the LP of the small intestine; (E) Siglec-FF4/80+CD11b+ macrophages from VAT; and (F) CD45hiF4/80+CD11b+ macrophages and CD45int microglia.

FIGURE 3.

Siglec-FF4/80+ macrophages express dTomato-C3aR and C3aR. Flow cytometric assessment of tdTomato-C3aR expression in different macrophage populations. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (AF) tdTomato-C3aR and C3aR expression in (A) M-CSF–derived F4/80+CD11b+ BM macrophages (M-CSF BMM); (B) GM-CSF–derived BM macrophage population (GM-CSF BMM) that was identified as CD115+MHCIIlo cells within the CD11c+CD11b+ cell population; (C) F4/80+CD11b+ PE macrophages; (D) LineageCD11c+CD11b+MHCII+F4/80+ macrophages from the LP of the small intestine; (E) Siglec-FF4/80+CD11b+ macrophages from VAT; and (F) CD45hiF4/80+CD11b+ macrophages and CD45int microglia.

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To determine whether tdTomato-C3aR expression correlates with C3aR protein expression, WT and tdTomato+ macrophages were stained with C3aR-specific mAb 14D4. The C3aR specificity was verified by flow cytometry using C3aR-deficient cells as control and by immunofluorescence microscopy using PE macrophages and GM-CSF–differentiated BMMs/BMDCs from WT and C3ar1−/− mice (Supplemental Fig. 2E). The tdTomato-C3aR expression matched the C3aR surface expression using mAb 14D4 in some but not all macrophage populations. PE macrophages and M-CSF–derived BMMs showed a strong C3aR surface expression (Fig. 3A, 3C). Furthermore, GM-CSF BMMs and brain microglia expressed C3aR at their surface, although at lower levels (Fig. 3B, 3F). In contrast, brain, LP, and VAT macrophages stained C3aR (Fig. 3D–F), suggesting intracellular localization. Indeed, we found C3aR expression in permeabilized VAT macrophages (Fig. 3E). Unfortunately, high background staining in permeabilized LP and brain macrophages precluded intracellular assessment of C3aR in such cells. In summary, our data demonstrate that only F4/80+Siglec-F macrophages express C3aR and that their C3aR surface expression is heterogeneous in the investigated organs.

The BM and VAT are considered reservoirs of eosinophils (51, 52). We found that BM eosinophils do not express tdTomato-C3aR (Fig. 4A). In contrast, eosinophils from blood, lung, LP, and VAT showed a clear tdTomato signal (Fig. 4B–E). The tdTomato-C3aR expression was strongest in VAT and lung tissue eosinophils. PE MCs did not express a tdTomato-C3aR signal (Fig. 4F).

FIGURE 4.

Circulating and most tissue-residing eosinophils express tdTomato-C3aR intracellularly. Flow cytometric determination of tdTomato-C3aR expression in eosinophils is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface or intracellular expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (AF) tdTomato-C3aR and/or C3aR expression in Siglec-F+ (A) BM, (B) blood, (C) lung, (D) LP lineageCD11c+CD11b+MHCIIloF4/80lo, (E) VAT F4/80int eosinophils, and (F) FcεRI+ST2+ PE MCs.

FIGURE 4.

Circulating and most tissue-residing eosinophils express tdTomato-C3aR intracellularly. Flow cytometric determination of tdTomato-C3aR expression in eosinophils is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface or intracellular expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (AF) tdTomato-C3aR and/or C3aR expression in Siglec-F+ (A) BM, (B) blood, (C) lung, (D) LP lineageCD11c+CD11b+MHCIIloF4/80lo, (E) VAT F4/80int eosinophils, and (F) FcεRI+ST2+ PE MCs.

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To correlate the tdTomato-C3aR with C3aR surface expression, we stained all tdTomato+ eosinophils with the C3aR Ab. Surprisingly, we found no surface expression of C3aR on almost all eosinophil populations (Fig. 4B, 4C, 4E), except eosinophils from the LP of the small intestine (Fig. 4D). However, after permeabilization, all tdTomato+ eosinophil populations and their WT counterparts stained C3aR+, whereas eosinophils from C3ar1−/− mice stained negative (Fig. 4B, 4C, 4E). Thus, lung and VAT eosinophils as well as circulating eosinophils express the C3aR intracellularly, whereas LP eosinophils express the C3aR on the cell surface.

During the past years, several DC subsets in lymphoid and nonlymphoid compartments have been defined (53). Currently, three distinct DC subsets can be distinguished comprising conventional DCs (cDCs), inflammatory monocyte-derived DCs (moDCs), and plasmacytoid DCs (pDCs). In mice, cDCs are further subdivided into CD11b+ and CD8-like subsets. Following this differentiation scheme, CD11b+ cDCs comprise splenic CD11b+CD8 DCs and the pulmonary CD11b+ cDCs, whereas the CD8a-like cDCs subset is composed of splenic CD8a+CD11b and pulmonary CD103+CD11b cDCs (54). At steady-state, lungs harbor a minor population of moDCs, which increases under inflammatory conditions (55). Similarly, in the LP of the small intestine, CD11b+ and CD103+ cDCs subpopulations have been described (56). In contrast, DCs in the VAT are poorly defined. We found an F4/80Siglec-FMHCII+CD11c+CD11b+ population in the VAT that we consider CD11b+ cDCs but no CD8-like cDC equivalent (data not shown). Finally, we observed a mixed BMDC/BMM population, as recently described by Helft et al. (41), with 20–30% CD11c+CD11b+CD115MHCII+ BMDCs and 70–80% CD11c+CD11b+CD115+ BMMs, when BM cells were cultured in GM-CSF–supplemented medium.

In the lung, only CD11b+ cDCs and moDCs showed a tdTomato signal, whereas CD103+ cDCs and pDCs did not (Fig. 5A, 5E). In contrast, both splenic DC subsets stained negative for the tdTomato signal (Supplemental Fig. 3A). Furthermore, we detected a strong tdTomato-C3aR signal in GM-CSF–derived BMDCs (Fig. 5B). In the LP of the small intestine, CD11b+ and CD103+ cDCs both expressed tdTomato-C3aR (Fig. 5C), although the expression level was somewhat higher in the CD11b+ cDC population. The VAT DC subset showed no tdTomato signal (Fig. 5D).

FIGURE 5.

Mucosal surface-associated DCs express tdTomato-C3aR and C3aR. Flow cytometric assessment of tdTomato-C3aR and C3aR surface or intracellular expression in different DC subpopulations. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface or intracellular expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (A) tdTomato-C3aR and C3aR expression in different pulmonary DC subsets. After excluding Siglec-F+ eosinophils and AMs as well as lineage+ cells (CD19, CD49b, CD3e, Ly6G), DCs were identified as CD11c+MHCII+ cells. In the DC population, CD103+ cDCs and CD11b+ DCs could be distinguished. The latter population was further subdivided into CD11b+ cDCs and moDCs. (B) tdTomato-C3aR and C3aR expression in GM-CSF–derived BMDCs that were identified as CD115MHCIIhi cells within the CD11c+CD11b+ population. (C) tdTomato-C3aR and C3aR expression in CD11c+CD11bMHCII+CD103+ and CD11c+CD11b+CD103loF4/80 DCs of the LP. (D) tdTomato-C3aR expression in F4/80Siglec-FCD11c+MHCII+ DCs from VAT. (E) tdTomato-C3aR expression in different pDC populations. In LP pDCs, C3aR surface staining was also performed. The gating strategy is depicted in Supplemental Fig. 3C–H.

FIGURE 5.

Mucosal surface-associated DCs express tdTomato-C3aR and C3aR. Flow cytometric assessment of tdTomato-C3aR and C3aR surface or intracellular expression in different DC subpopulations. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface or intracellular expression, the dashed lines depict the staining of cells from C3ar1−/− mice. (A) tdTomato-C3aR and C3aR expression in different pulmonary DC subsets. After excluding Siglec-F+ eosinophils and AMs as well as lineage+ cells (CD19, CD49b, CD3e, Ly6G), DCs were identified as CD11c+MHCII+ cells. In the DC population, CD103+ cDCs and CD11b+ DCs could be distinguished. The latter population was further subdivided into CD11b+ cDCs and moDCs. (B) tdTomato-C3aR and C3aR expression in GM-CSF–derived BMDCs that were identified as CD115MHCIIhi cells within the CD11c+CD11b+ population. (C) tdTomato-C3aR and C3aR expression in CD11c+CD11bMHCII+CD103+ and CD11c+CD11b+CD103loF4/80 DCs of the LP. (D) tdTomato-C3aR expression in F4/80Siglec-FCD11c+MHCII+ DCs from VAT. (E) tdTomato-C3aR expression in different pDC populations. In LP pDCs, C3aR surface staining was also performed. The gating strategy is depicted in Supplemental Fig. 3C–H.

Close modal

In the next step, we assessed tdTomato-C3aR expression in different tissue pDCs and Flt3L-induced pDCs. The pDCs are strong producers of IFN-α upon viral infection (57) and have been identified in the spleen (58), lung (59), LP, Peyer’s patches (60), and in the adipose tissue (61). Additionally, 10–20% pDCs differentiate from BM cells in response to Flt3L stimulation (62). Using CD11b, CD11c, B220, mPDCA-1, and Siglec-H as markers, we identified pDCs in all tissues outlined above using the gating strategies provided in Supplemental Fig. 3C–H. Among the different tissue-residing pDCs tested, only a small fraction of the LP pDCs stained positive for tdTomato-C3aR (Fig. 5E).

Counterstaining of pulmonary tdTomato+ DCs with mAb 14D4 showed that moDCs but not CD11b+ cDCs (Fig. 5A) express C3aR at their cell surface. However, CD11b+ cDCs express C3aR intracellularly in low amounts as shown in permeabilized cells (Fig. 5A, CD11b+ intracellular panel). By confocal microscopy, the tdTomato signal colocalized with some but not all CD11b+CD11c+ cells (Supplemental Fig. 3B), confirming that most tdTomato-C3aR+ cells in the airways were resident moDCs. In the small intestine, LP CD11b+ cDCs showed a clear surface expression of C3aR, whereas C3aR expression in CD103+ cDCs was very weak (Fig. 5C). Finally, we found weak C3aR surface expression in GM-CSF–derived BMDCs (Fig. 5B). Similarly, we counterstained tdTomato-C3aR+ pDCs and found that LP pDCs express C3aR at the cell surface (Fig. 5E). From all DC populations tested, moDCs were the DC population with the strongest C3aR surface expression. In summary, we demonstrate that small intestinal LP cDCs and pDCs, lung moDCs, and GM-CSF–derived BMDCs express C3aR. Furthermore, pulmonary CD11b+ cDCs show low intracellular C3aR expression. In contrast, neither CD11b+ splenic DCs nor pulmonary and splenic CD8-like cDCs express C3aR.

The expression of the anaphylatoxin receptors by murine lymphoid cells is controversial (2931, 42, 63). In this study, we used the tdTomato-C3aR reporter mouse to monitor C3aR expression in B and T cells in the circulation, different secondary lymphoid organs including the spleen, lymph nodes and Peyer’s patches, as well as the lung. Additionally, we assessed C3aR expression in B-1 cell subsets residing in the PE cavity (64). We found no tdTomato-C3aR+ B or T cells in the circulation or any of the tissues under steady-state conditions (Fig. 6A–E, 6G–I). In line with this finding, we observed no C3ar1 mRNA expression in splenic T and B cells (Supplemental Fig. 4). Next, we stimulated naive splenic T cells using anti-CD3 and anti-CD28 Abs. We found a strong proliferative response of the T cells (data not shown). However, this treatment did not result in the induction of tdTomato-C3aR expression in the T cells (Fig. 6F).

FIGURE 6.

Circulating and tissue-resident B and T lymphocytes do not express tdTomato-C3aR. Flow cytometric assessment of tdTomato-C3aR expression in different B and T lymphocyte populations is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. tdTomato-C3aR expression in B (AD) and T lymphocytes (EH) from the circulation and different tissues is shown. tdTomato-C3aR expression in (A) splenic CD19+B220+ B lymphocytes, (B) circulating CD19+ B cells, and (C) PE B1 lymphocytes is shown. After exclusion of CD11bhi macrophages, B-1 lymphocytes were identified as CD19+IgM+CD43+ cells. They were further divided into CD5+ B-1a and CD5 B-1b lymphocytes. (D) tdTomato-C3aR expression in CD19+B220+ B lymphocytes from mediastinal lymph nodes. (E and F) tdTomato-C3aR expression in (E) naive splenic CD49bCD3+ T lymphocytes or in (F) T lymphocytes stimulated in vitro for 5 d in the presence of CD3- and CD28-specific Abs. (G–I) tdTomato-C3aR expression in CD3+CD4+ T lymphocytes from (G) blood, (H) mediastinal lymph nodes, and (I) lung tissue. mLN, mediastinal lymph node.

FIGURE 6.

Circulating and tissue-resident B and T lymphocytes do not express tdTomato-C3aR. Flow cytometric assessment of tdTomato-C3aR expression in different B and T lymphocyte populations is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. tdTomato-C3aR expression in B (AD) and T lymphocytes (EH) from the circulation and different tissues is shown. tdTomato-C3aR expression in (A) splenic CD19+B220+ B lymphocytes, (B) circulating CD19+ B cells, and (C) PE B1 lymphocytes is shown. After exclusion of CD11bhi macrophages, B-1 lymphocytes were identified as CD19+IgM+CD43+ cells. They were further divided into CD5+ B-1a and CD5 B-1b lymphocytes. (D) tdTomato-C3aR expression in CD19+B220+ B lymphocytes from mediastinal lymph nodes. (E and F) tdTomato-C3aR expression in (E) naive splenic CD49bCD3+ T lymphocytes or in (F) T lymphocytes stimulated in vitro for 5 d in the presence of CD3- and CD28-specific Abs. (G–I) tdTomato-C3aR expression in CD3+CD4+ T lymphocytes from (G) blood, (H) mediastinal lymph nodes, and (I) lung tissue. mLN, mediastinal lymph node.

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In the lung and intestine, innate lymphoid cells (ILCs) play important roles in the regulation of tolerance (65). Three different ILC subsets have been described: group 1 includes ILC1 and NK cells, group 2 consists of ILC2, and group 3 consists of ILC3 (65). NK cells and ILC2 reside in the lung. NK cells are also found in the spleen. ILC2 are further recruited upon activation by alarmins (65). ILC3 are not present in the lung but, together with NK and ILC2, can be found in high numbers in the LP of the small intestine (66). Splenic NK cells did not stain positive for tdTomato-C3aR (Fig. 7A). In the lung, we observed low numbers of NK cells and ILC2 but neither ILC1 nor ILC3 (Fig. 7B, 7C), both of which stained negative for tdTomato-C3aR. In contrast to the lung, LP of the small intestine contained some tdTomato-C3aR+ ILCs. Whereas NK, ILC1 (Fig. 7D), and ILC2 (Fig. 7E) were tdTomato-C3aR, a subset of lineageCD90.2+CD25CD127+ ILC3 cells showed a positive tdTomato signal (Fig. 7E). C3aR surface staining of ILC3 using the C3aR-specific mAb 14D4 was negative, suggesting that C3aR is either expressed intracellularly or that the sensitivity of flow cytometry is too low to detect its expression (Fig. 7E). In summary, our results show that most ILCs do not express C3aR at steady-state. Interestingly, a subgroup of LP ILC3 expresses low numbers of C3aR.

FIGURE 7.

Expression of tdTomato-C3aR in ILC3 but not in ILC1, NK, or ILC2 from LP of the small intestine. Flow cytometric evaluation of tdTomato-C3aR expression in group 1–3 ILCs is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface expression, the dashed lines depict the staining of cells from C3ar1−/− mice. tdTomato-C3aR expression in (A) splenic NK cells, (B) pulmonary NK cells, (C) pulmonary ILC2, (D) CD127+CD49b ILC1 and CD127CD49b+ NK cells from the LP, and (E) CD90.2+CD25+CD127+ ILC2 and CD90.2+CD25CD127+ ILC3 from the LP of the small intestine is shown.

FIGURE 7.

Expression of tdTomato-C3aR in ILC3 but not in ILC1, NK, or ILC2 from LP of the small intestine. Flow cytometric evaluation of tdTomato-C3aR expression in group 1–3 ILCs is shown. The gray histograms show the WT signal; the black lines correspond to the signal in the tdTomato-C3ar1fl/fl mice. In case of C3aR surface expression, the dashed lines depict the staining of cells from C3ar1−/− mice. tdTomato-C3aR expression in (A) splenic NK cells, (B) pulmonary NK cells, (C) pulmonary ILC2, (D) CD127+CD49b ILC1 and CD127CD49b+ NK cells from the LP, and (E) CD90.2+CD25+CD127+ ILC2 and CD90.2+CD25CD127+ ILC3 from the LP of the small intestine is shown.

Close modal

The expression of C3aR was reported in steady-state lung epithelium (13). To explore the expression of the tdTomato reporter in the different lung epithelial populations, we examined them by two-photon and confocal microscopy. Examination of 300-μm-thick sections of WT and tdTomato-C3aRfl/fl lungs by two-photon microscopy revealed several tdTomato-C3aR+ cells in the bronchial compartment of the airways but only a vey small number of tdTomato-C3aR+ cells in the alveolar compartment (Fig. 8A). Confocal microscopy showed that tdTomato-C3aR+ cells in the airways were not epithelial nonciliated bronchiolar secretory cells (Clara cells), because the tdTomato signal did not colocalize with the CC10 molecule (67) (Fig. 8B). Consistent with the very low numbers of tdTomato-C3aR+ cells in the alveolar compartment (Fig. 8A), CD45autofluorecence+MHCII+ cells, which correspond to the alveolar type II cells (68), stained negative for tdTomato-C3aR (Fig. 8C). Finally, we found that a small group within the epithelial cells from the small intestine expresses tdTomato-C3aR (Fig. 8D).

FIGURE 8.

Minor tdTomato-C3aR expression in epithelial cells from the small intestine. (A) Assessment of tdTomato-C3aR expression in the lung and the airways of WT and tdTomato-C3ar1fl/fl mice by two-photon microscopy (original magnification ×20). Shown is the fluorescence signal of tdTomato-C3aR (red) from autofluorescent connective tissue cells of the airways (green) and from the alveolar epithelial cells (turquoise). (B) Determination of the spatial localization of tdTomato-C3aR expression in lung tissue by confocal microscopy (original magnification ×40). Lung sections from tdTomato-C3ar1fl/fl mice were stained with CC10-specific Ab to identify Clara cells. (C) tdTomato-C3aR expression in CD45autofluorescence+MHCII+ type II alveolar epithelial cells. The gray histogram shows the WT signal; the black line corresponds to the signal in cells from tdTomato-C3ar1fl/fl mice. (D) tdTomato-C3aR expression in CD45CD3autofluorescence+ epithelial cells from the LP of the small intestine. Shown is the signal obtained in cells from WT and tdTomato-C3ar1fl/fl mice. The panel on the right quantifies the tdTomato-C3aR signal in n = 4–5 mice from each group. *p < 0.05.

FIGURE 8.

Minor tdTomato-C3aR expression in epithelial cells from the small intestine. (A) Assessment of tdTomato-C3aR expression in the lung and the airways of WT and tdTomato-C3ar1fl/fl mice by two-photon microscopy (original magnification ×20). Shown is the fluorescence signal of tdTomato-C3aR (red) from autofluorescent connective tissue cells of the airways (green) and from the alveolar epithelial cells (turquoise). (B) Determination of the spatial localization of tdTomato-C3aR expression in lung tissue by confocal microscopy (original magnification ×40). Lung sections from tdTomato-C3ar1fl/fl mice were stained with CC10-specific Ab to identify Clara cells. (C) tdTomato-C3aR expression in CD45autofluorescence+MHCII+ type II alveolar epithelial cells. The gray histogram shows the WT signal; the black line corresponds to the signal in cells from tdTomato-C3ar1fl/fl mice. (D) tdTomato-C3aR expression in CD45CD3autofluorescence+ epithelial cells from the LP of the small intestine. Shown is the signal obtained in cells from WT and tdTomato-C3ar1fl/fl mice. The panel on the right quantifies the tdTomato-C3aR signal in n = 4–5 mice from each group. *p < 0.05.

Close modal

To assess whether the genomic construct of tdTomato with C3aR alters the functionality of the C3aR, we compared functional responses of both receptors side by side. Stimulation of C3aR with C3a results in rapid internalization (7, 69). As we found strong C3aR surface expression in PE macrophages, we used this cell population to determine C3a-driven C3aR internalization. PE macrophages were identified in PE lavage as F4/80+C3aR+ cells. WT and tdTomato-C3ar1fl/fl macrophages express similar levels of C3aR at their surface (Fig. 9A). When we incubated WT and tdTomato-C3ar1fl/fl macrophages with increasing concentrations of C3a, we observed a rapid and similar dose-dependent decrease of C3aR surface expression in WT and tdTomato-C3ar1fl/fl macrophages. The maximum of C3aR internalization was reached 3 min after C3a stimulation using a concentration of 10 nM C3a (Fig. 9B), although most of the fluorescence signal disappeared within 1 min, suggesting a rapid internalization of the C3aR upon C3a stimulation. Between 1 and 3 min after C3a administration, the fluorescence signal further declined although at a much lower pace. Eventually, the normalized C3aR MFI decreased to 30% of unstimulated controls (Fig. 9C).

FIGURE 9.

C3a drives rapid C3aR internalization and mobilization of intracellular calcium in PE macrophages from WT and tdTomato-C3ar1fl/fl mice. (A) Determination of C3aR surface expression in F4/80+ PE macrophages from WT (gray histogram) and tdTomato-C3ar1fl/fl mice (black line) by flow cytometry. PE macrophages from C3ar1−/− mice (dashed line) served as controls. Data show representative histograms of the C3aR signal in unstimulated PE macrophages as compared with 1, 10, and 100 nM C3a stimulation for 3 min at 37°C. (B) Comparison of C3aR surface expression in PE macrophages from WT and tdTomato-C3ar1fl/fl mice in response to stimulation with increasing concentrations of C3a (1, 10, 100 nM) for 3 min at 37°C. Shown is the ΔMFI of C3aR staining, which is defined as the MFI obtained by C3aR staining of cells from WT or tdTomato-C3ar1fl/fl mice corrected by the C3aR staining obtained with PE macrophages from C3aR-deficient mice. Values shown are the mean ± SEM; n = 6 per group. (C) Comparison of C3aR surface expression before as well as 1, 3, and 9 min after stimulation with 10 nM C3a at 37°C. Shown is the ΔMFI of C3aR staining. Values shown are the mean ± SEM; n = 5–6 per group. (D) C3a-mediated increase of [Ca2+]i in F4/80+ thioglycollate-elicited PE macrophages from WT mice (gray line), tdTomato-C3ar1fl/fl mice (black line), or C3ar1−/−C5ar1−/− (dashed line) mice. Shown is the ΔMFI of the fluorescence peak. Values shown are the mean ± SEM; n = 7 per group. Statistical differences between groups were assessed by a Student t test. (E) Micrcoscopic evaluation of the C3a-mediated change of [Ca2+]i in thioglycollate-elicited PE macrophages from WT and tdTomato-C3ar1fl/fl mice. Adherent thioglycollate-elicited PE macrophages from both groups were loaded with Fluo4-AM and challenged with C3a (37 nM). The images show the fluorescence emission in the FITC (Fluo4 emission) and PE channels (tdTomato), as wells as from polarized light before and 6 s after C3a stimulation (×40 objective). Scale bar, 10 μm. Data are representative of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 9.

C3a drives rapid C3aR internalization and mobilization of intracellular calcium in PE macrophages from WT and tdTomato-C3ar1fl/fl mice. (A) Determination of C3aR surface expression in F4/80+ PE macrophages from WT (gray histogram) and tdTomato-C3ar1fl/fl mice (black line) by flow cytometry. PE macrophages from C3ar1−/− mice (dashed line) served as controls. Data show representative histograms of the C3aR signal in unstimulated PE macrophages as compared with 1, 10, and 100 nM C3a stimulation for 3 min at 37°C. (B) Comparison of C3aR surface expression in PE macrophages from WT and tdTomato-C3ar1fl/fl mice in response to stimulation with increasing concentrations of C3a (1, 10, 100 nM) for 3 min at 37°C. Shown is the ΔMFI of C3aR staining, which is defined as the MFI obtained by C3aR staining of cells from WT or tdTomato-C3ar1fl/fl mice corrected by the C3aR staining obtained with PE macrophages from C3aR-deficient mice. Values shown are the mean ± SEM; n = 6 per group. (C) Comparison of C3aR surface expression before as well as 1, 3, and 9 min after stimulation with 10 nM C3a at 37°C. Shown is the ΔMFI of C3aR staining. Values shown are the mean ± SEM; n = 5–6 per group. (D) C3a-mediated increase of [Ca2+]i in F4/80+ thioglycollate-elicited PE macrophages from WT mice (gray line), tdTomato-C3ar1fl/fl mice (black line), or C3ar1−/−C5ar1−/− (dashed line) mice. Shown is the ΔMFI of the fluorescence peak. Values shown are the mean ± SEM; n = 7 per group. Statistical differences between groups were assessed by a Student t test. (E) Micrcoscopic evaluation of the C3a-mediated change of [Ca2+]i in thioglycollate-elicited PE macrophages from WT and tdTomato-C3ar1fl/fl mice. Adherent thioglycollate-elicited PE macrophages from both groups were loaded with Fluo4-AM and challenged with C3a (37 nM). The images show the fluorescence emission in the FITC (Fluo4 emission) and PE channels (tdTomato), as wells as from polarized light before and 6 s after C3a stimulation (×40 objective). Scale bar, 10 μm. Data are representative of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001.

Close modal

Furthermore, we determined C3a-induced mobilization of [Ca2+]i in thioglycollate-elicited PE macrophages from WT and tdTomato-C3ar1fl/fl mice. Stimulation of F4/80+ macrophages with C3a resulted in a transient increase in [Ca2+]i in cells from WT and tdTomato-C3ar1fl/fl mice (Fig. 9D). The amplitude of the response, measured as the ΔMFI, was similar in WT and tdTomato-C3ar1fl/fl macrophages. In line with the flow cytometric analysis, microscopic assessment of the C3a-mediated change in [Ca2+]i confirmed the rapid and transient influx of calcium into macrophages from WT and tdTomato-C3ar1fl/fl macrophages. We found a peak of [Ca2+]i 6 s after C3a stimulation (Fig. 9E). In contrast, we observed no change in [Ca2+]i in thioglycollate-elicited PE macrophages from C3ar1−/−C5ar1−/− mice upon C3a challenge (data not shown), either by flow cytometry or by confocal microscopy. In conclusion, our findings demonstrate that the C3aR expression in tdTomato-C3ar1fl/fl and WT macrophages is similar. Furthermore, C3a-mediated internalization of C3aR and mobilization of intracellular calcium follows the same pattern in WT and tdTomato-C3ar1fl/fl macrophages, suggesting that the C3aR in tdTomato-C3ar1fl/fl mice is functional.

Although the mouse C3aR was cloned 20 y ago (11), a detailed understanding of C3aR expression in tissue-residing immune and stromal cells is still lacking (9). In this study, we generated a knock-in reporter mouse strain, designed to express a fluorescent tdTomato complex and C3aR. To avoid interference with C3aR signal transduction, we added the tdTomato sequence to the 5′ end of C3ar1, because structural data have shown that the N-terminal part of the C3aR does not bind C3a (6, 70). In contrast, the C-terminal part of the C3aR harbors important phosphorylation sites, conserved in human and mice that drive receptor internalization and signal transduction (7). Furthermore, N-terminal extension of C5aR1 with GFP did not affect C5aR1 expression and function (42) whereas C-terminal GFP fusion resulted in a block of C5aR1 translocation from the cytoplasm to the cell surface (63). In agreement with these observations, the C3aR protein from tdTomato-C3ar1fl/fl mice was correctly targeted to the cell surface and fully functional as shown by equivalent C3aR internalization and calcium mobilization in PE macrophages from tdTomato-C3ar1fl/fl and WT mice in response to C3a.

Previous studies showed differences in C3ar1 mRNA expression patterns between mice and humans. In the former, the mRNA was detected mostly in the heart and the lung, and to a lesser extent in the kidney, liver, skeletal muscles, and adipose tissue, but not in the brain and the spleen (10, 11). In humans, C3AR1 mRNA was found in large amounts in lung, spleen, small intestine, peripheral blood leukocytes, s.c. adipose tissue, and at low levels in the kidney and heart but not in the brain (5, 6, 71). Our data obtained with the tdTomato-C3aR knock-in mouse confirmed the presence of C3aR expression in the lung and VAT. Surprisingly, in contrast to previous mRNA studies, we observed a strong tdTomato signal in the brain. Until now, C3ar1 mRNA expression had been evaluated solely at the level of the whole mouse brain (10, 11). Differences in the isolation procedure may account for the different expression pattern observed in these studies as compared with results obtained with the tdTomato-C3aR reporter mouse. However, in humans, the low C3aR expression in whole brain neglects high C3AR1 mRNA expression in defined regions and underrepresented cell populations of the brain (5). Furthermore, we found tdTomato-C3aR expression in the LP of the small intestine, whereas the number of tdTomato-C3aR+ cells in the spleen was at best minor. Also, the number of tdTomato-C3aR+ cells in BAL, blood, and BM was very low. The cell-specific determination of C3aR expression revealed that cells of the myeloid lineage such as macrophages, eosinophils, and DCs and in much lower frequency ILC3 from the small intestine but not other ILCs, B or T lymphocytes express this anaphylatoxin receptor.

In agreement with our observations that only a few tdTomato-C3aR+ cells were present in the BM and in strong contrast to human data (5, 16, 69, 72), we did not observe tdTomato-C3aR+ neutrophils in that compartment. Furthermore, we observed at best a very minor expression of C3ar1 mRNA in sorted BM neutrophils from WT or tdTomato-C3ar1fl/fl mice. In agreement with this finding, we found no surface or intracellular C3aR expression in WT BM neutrophils. In the literature, direct evidence for C3aR expression in mouse neutrophils is scarce. In a study in which Gr-1 was used to identify neutrophils (24), the authors described C3aR surface and intracellular C3aR expression using the D20 Ab. Using this Ab, we found no C3aR expression in Ly6G+ neutrophils, suggesting that C3aR expression in Gr-1+ cells resulted from other cells that express Gr-1 such as monocytes (45, 73). Furthermore, our data suggest that the D20 Ab recognizes a protein that is also present in the absence of C3aR expression. In line with our findings, two other reports found no C3aR expression in BM neutrophils (33, 46). Interestingly, C3aR was upregulated in response to LPS in low- and high-density neutrophils in a small intestinal tumor model and promoted neutrophil extracellular trap formation (46). In agreement with these observations, we found upregulation of tdTomato-C3aR expression in airway and tissue neutrophils during the effector phase of HDM-driven allergic asthma. However, C3aR was not expressed at the cell surface, suggesting either intracellular localization of C3aR or a very low level of surface expression that is below the detection limit of the assay.

Our data show that the major cell population expressing C3aR under steady-state conditions are macrophages. In agreement with earlier studies (74, 75), we found that all PE macrophages from WT and tdTomato-C3ar1fl/fl mice express C3aR at the cell surface. Our data also confirm C3aR expression in VAT macrophages (76), microglia (77, 78), and in BMMs (17, 24). Additionally, we are reporting, to our knowledge for the first time, the expression of C3aR in macrophages of the brain and the LP of the small intestine, although C3aR is not expressed at the cell surface. Interestingly, we observed an inverse correlation between the expression of C3aR and Siglec-F in macrophages. Indeed, all F4/80+Siglec-F macrophages were tdTomato-C3aR+, whereas Siglec-F+ macrophages stained negative. Siglec-F is a C-type lectin, whose function is poorly understood. Recently, the pulmonary glycoprotein Muc5b was identified as a ligand for Siglec-F that induces apoptosis of eosinophils (79). Of note, the C3a/C3aR axis has been reported to drive the production of a similar mucin, Muc5a, from Clara cells (12). The AM population can be divided into two subsets, comprising AMs residing in the alveolar space and a sessile fraction of AMs, attached to the alveolar epithelium via connexin 43 (80). Our data show that both populations stain negative for tdTomato-C3aR, confirming a previous report showing that pulmonary leukocytes, but not AMs, express C3aR (34).

In addition to macrophages, we found that eosinophils from different organs express tdTomato-C3aR and C3aR. Earlier studies showed that C3a is expressed on human eosinophils (81), triggers reactive oxygen species and calcium flux, and is chemotactic for human blood eosinophils (1820, 82). Our data demonstrate that blood eosinophils express tdTomato-C3aR. Similarly, we demonstrate intracellular C3aR expression in pulmonary and VAT eosinophils. Only eosinophils from the LP of the small intestine expressed C3aR on the cell surface. Eosinophils from the BM, the site of eosinophil development and differentiation (51), stained C3aR. BM eosinophils are immature and require activation by IL-5 to egress and home to tissues via the bloodstream (51). Our observation that eosinophils start to express C3aR intracellularly in the blood, which is enhanced in the lung and the VAT and already translocates to the cell surface in LP eosinophils, suggests that C3aR may also play important roles in eosinophil activation. Furthermore, the intestine (51) and the VAT (52) have been described as retention sites for eosinophils, indicating that C3aR may play a role in their retention, at least in the LP.

Our data confirm a previous report (83) demonstrating that PE MCs do not express C3aR at steady-state, although functional studies suggest that C3a activates MCs from skin and the small intestine (84, 85). In line with this notion, variable and complex C3aR expression has been shown in human MCs. Human skin MCs (23, 86), CD34+-derived human MCs (87), the human MC line HMC-1 (88), and LAD1 and 2 (89) have been reported to express C3aR (87) whereas BM-derived human MCs do not (90).

The third population largely expressing tdTomato-C3aR was DCs. In vitro–generated DC populations of human (17, 91, 92) and murine origin have been reported to express C3aR (26, 73). We found that GM-CSF–differentiated BM cells, which contained macrophages and DCs (41), were tdTomato-C3aR+ and expressed C3aR at the cell surface. In tissue, we observed that some pulmonary and LP-resident DCs express tdTomato-C3aR and C3aR at the surface, that is, moDCs and CD11b+ cDCs in the lung as well as CD103+, CD11b+ cDCs, and pDCs in the LP of the small intestine. At this point, the function of C3aR on mucosal cDCs and pDCs is unclear. Importantly, LP CD103+ cDCs produce retinoic acid and TGF-β, both of which are important for the generation of Foxp3+ regulatory T cells (93). Also, they drive the differentiation of IgA-producing plasma cells, whereas CD11b+ cDCs promote Th17 cell differentiation (94). In the lung, CD11b+ cDCs are critical drivers of Th2/Th17 maladaptive immune response in allergic asthma (50, 55). Previous studies have shown that C3aR signaling in BMDCs is important for Ag uptake and T cell activation (26), indicating that the C3a/C3aR axis in tissue DCs may have similar functions. Lymphoid tissue DCs and most pDC populations stained negative for tdTomato-C3aR.

Interestingly, some subsets of macrophages, DCs, and most of the tissue eosinophils expressed C3aR intracellularly. Recent studies already reported in human cells the intracellular activation of the C3/C3a/C3aR signaling axis, either through the conversion of C3 into C3a/C3b in vesicles where C3aR resides (30), or through the activation of NLRP3 inflammasome in the cytosol leading to the secretion of IL-1β by such cells (95). Importantly, IL-1β plays an important role in the regulation of the experimental allergic asthma phenotype through its impact on Th2/Th17 immune responses (96). In line with this notion, human eosinophils secrete IL-1β and drive the production of IL-17A by CD4+ T cells (97).

The expression of C3aR in lung parenchymal cells is unclear. Using RNA hybridization of lung sections (13) or immunohistochemical C3aR staining with polyclonal Abs, the expression of C3aR was reported on bronchial (13, 98, 99), alveolar epithelial cells and on smooth muscle cells (12, 13). In contrast, another study demonstrated C3aR expression solely on leukocytes using C3aR-specific mAb 14D4 for surface staining (34). In line with this observation, our confocal microscopy and flow cytometric data provide no evidence for C3aR expression on naive alveolar or airway epithelial cells. Thus, C3aR expression in the lung is restricted to myeloid cells. This does not exclude that C3aR might be induced on epithelial cells under inflammatory conditions. Similar to the lung, most epithelial cells in the intestine were tdTomato-C3aR. Only a minor fraction expressed the C3aR.

Finally, we assessed the expression of the C3aR in lymphoid cells, which is still controversial. In previous studies, the expression of C3aR was described in activated human T lymphocytes (27, 100), but not in B lymphocytes (27). However, more recent studies reported that C3aR is expressed intracellularly by naive and in vitro–activated human T lymphocytes (30). Furthermore, several studies reported weak expression of both C3aR and C5aR1 in naive and activated mouse T lymphocytes (31) and an important role for the development of regulatory T cells (29, 101). However, we and others have failed in detecting C5aR1 in B, T, and NKT cells using a GFP-C5aR1 reporter mouse at steady-state or upon activation (42, 63). Our data obtained with the tdTomato-C3aR knock-in mouse demonstrate no C3aR expression in naive B and T lymphocytes. Also, stimulation of naive T lymphocytes with anti-CD3/CD28 mAbs did not induce C3aR expression. In addition to B and T lymphocytes, we assessed C3aR expression in ILCs, which play important roles in mucosal immunology (65). ILCs are divided into three groups containing ILC1 and NK cells (group 1), ILC2 (group 2), and ILC3 (group 3). We found no tdTomato-C3aR signal in group 1 and 2 ILCs. In contrast, we observed that a small fraction of ILC3 in the LP of the small intestine expressed tdTomato-C3aR. Transcriptome analysis revealed that ILC3 are a heterogeneous population of cells that produce several cytokines, including IL17-A/F and IL-22 in response to myeloid cell–derived IL-1β and IL-23 (102).

Finally, we demonstrate that the C3aR in tdTomato-C3ar1fl/fl knock-in mice is fully functional. Using WT and the tdTomato-C3aR PE macrophages, we show that C3a triggers C3aR internalization. The concentration of C3a, needed to drive C3aR internalization, was similar to that reported for human cells (69), but the maximum internalization in mouse cells already occurred in response to 10 nM C3a as compared with 100 nM using human granulocytes. Similarly, we observed a more rapid internalization of C3aR compared with human cells (69), where most of the C3aR signal was lost only 10 min after C3a stimulation. Additionally, we observed a similar rapid and transient increase in intracellular calcium concentration in thioglycollate-elicited PE macrophages from WT and tdTomato-C3ar1fl/fl.

Collectively, our findings provide detailed and novel insights into steady-state C3aR expression in myeloid, lymphoid, and pulmonary parenchymal cells. Our data clearly demonstrate a heterogeneous C3aR expression pattern in macrophages, eosinophils, and DCs, depending on tissue distribution and the cellular subsets within such tissues. Furthermore, our results show spatial heterogeneity of C3aR expression, that is, intracellular and surface expression. Finally, we provide evidence that naive neutrophils, B and T lymphocytes, most ILCs, as well as pulmonary epithelial cells do not express C3aR. The tdTomato-C3aR will prove useful to track C3aR expression and cell-specifically delete C3aR in experimental models of allergic asthma (14, 73), emphysema (103), ischemia/reperfusion injury (104), cancer (46), inflammatory neurologic (105) and renal diseases (106), as well as infections (84, 107) among others, where the C3a/C3aR axis has been demonstrated to play important roles. It will also be instrumental to define the role of C3aR in VGF peptide TLQP-21–mediated functions, as this peptide can also bind to C3aR, at least in rodents (108).

We thank E. Strerath, G. Köhl, D. Theil, L. Pohl, and C. Lehmann for excellent technical assistance.

This work was supported by Deutsche Forschungsgemeinschaft Grants IRTG 1911 (projects A1 and A2 to Y.L., J.K., and P.K., respectively), IRTG 1911 (projects B1 and B2 to C.M.K. and J.K.), and KO 1245/3-1 (to J.K.). L.N.A. was supported by the National Council for Technological and Scientific Development/Science without Borders (Brazil).

The online version of this article contains supplemental material.

Abbreviations used in this article:

AF

Alexa Fluor

AM

alveolar macrophage

BAL

bronchoalveolar lavage

BM

bone marrow

BMDC

BM-derived dendritic cell

BMM

BM-derived macrophage

BV

Brilliant Violet

[Ca2+]i

intracellular Ca2+ concentration

CC10

Clara cell protein 10

CD

cluster of differentiation

DC

dendritic cell

D-PBS

Dulbecco’s PBS

eF

eFluor

HDM

house dust mite

ILC

innate lymphoid cell

IRES

internal ribosome entry site

LP

lamina propria

MC

mast cell

MFI

mean fluorescence intensity

ΔMFI

change in relative mean fluorescence intensity

MHCII

MHC class II

moDC

monocyte-derived DC

PBS20

PBS supplemented with 20% FCS

pDC

plasmacytoid DC

PE

peritoneal

RBCL

RBC lysis

tdTomato

tandem-dye Tomato

VAT

visceral adipose tissue

WT

wild-type.

1
Klos
,
A.
,
A. J.
Tenner
,
K. O.
Johswich
,
R. R.
Ager
,
E. S.
Reis
,
J.
Köhl
.
2009
.
The role of the anaphylatoxins in health and disease.
Mol. Immunol.
46
:
2753
2766
.
2
Kolev
,
M.
,
G.
Le Friec
,
C.
Kemper
.
2014
.
Complement—tapping into new sites and effector systems.
Nat. Rev. Immunol.
14
:
811
820
.
3
Verschoor
,
A.
,
C. M.
Karsten
,
S. P.
Broadley
,
Y.
Laumonnier
,
J.
Köhl
.
2016
.
Old dogs—new tricks: immunoregulatory properties of C3 and C5 cleavage fragments.
Immunol. Rev.
274
:
112
126
.
4
Coulthard
,
L. G.
,
T. M.
Woodruff
.
2015
.
Is the complement activation product C3a a proinflammatory molecule? Re-evaluating the evidence and the myth.
J. Immunol.
194
:
3542
3548
.
5
Ames
,
R. S.
,
Y.
Li
,
H. M.
Sarau
,
P.
Nuthulaganti
,
J. J.
Foley
,
C.
Ellis
,
Z.
Zeng
,
K.
Su
,
A. J.
Jurewicz
,
R. P.
Hertzberg
, et al
.
1996
.
Molecular cloning and characterization of the human anaphylatoxin C3a receptor.
J. Biol. Chem.
271
:
20231
20234
.
6
Crass
,
T.
,
U.
Raffetseder
,
U.
Martin
,
M.
Grove
,
A.
Klos
,
J.
Köhl
,
W.
Bautsch
.
1996
.
Expression cloning of the human C3a anaphylatoxin receptor (C3aR) from differentiated U-937 cells.
Eur. J. Immunol.
26
:
1944
1950
.
7
Settmacher
,
B.
,
C.
Rheinheimer
,
H.
Hamacher
,
R. S.
Ames
,
A.
Wise
,
L.
Jenkinson
,
D.
Bock
,
M.
Schaefer
,
J.
Köhl
,
A.
Klos
.
2003
.
Structure-function studies of the C3a-receptor: C-terminal serine and threonine residues which influence receptor internalization and signaling.
Eur. J. Immunol.
33
:
920
927
.
8
Settmacher
,
U.
,
H. D.
Volk
,
S.
Jahn
,
K.
Neuhaus
,
F.
Kuhn
,
R.
von Baehr
.
1991
.
Characterization of human lymphocytes separated from fetal liver and spleen at different stages of ontogeny.
Immunobiology
182
:
256
265
.
9
Klos
,
A.
,
E.
Wende
,
K. J.
Wareham
,
P. N.
Monk
.
2013
.
International union of basic and clinical pharmacology. [corrected]. LXXXVII. Complement peptide C5a, C4a, and C3a receptors.
Pharmacol. Rev.
65
:
500
543
.
10
Hsu
,
M. H.
,
J. A.
Ember
,
M.
Wang
,
E. R.
Prossnitz
,
T. E.
Hugli
,
R. D.
Ye
.
1997
.
Cloning and functional characterization of the mouse C3a anaphylatoxin receptor gene.
Immunogenetics
47
:
64
72
.
11
Tornetta
,
M. A.
,
J. J.
Foley
,
H. M.
Sarau
,
R. S.
Ames
.
1997
.
The mouse anaphylatoxin C3a receptor: molecular cloning, genomic organization, and functional expression.
J. Immunol.
158
:
5277
5282
.
12
Dillard
,
P.
,
R. A.
Wetsel
,
S. M.
Drouin
.
2007
.
Complement C3a regulates Muc5ac expression by airway Clara cells independently of Th2 responses.
Am. J. Respir. Crit. Care Med.
175
:
1250
1258
.
13
Drouin
,
S. M.
,
J.
Kildsgaard
,
J.
Haviland
,
J.
Zabner
,
H. P.
Jia
,
P. B.
McCray
Jr.
,
B. F.
Tack
,
R. A.
Wetsel
.
2001
.
Expression of the complement anaphylatoxin C3a and C5a receptors on bronchial epithelial and smooth muscle cells in models of sepsis and asthma.
J. Immunol.
166
:
2025
2032
.
14
Drouin
,
S. M.
,
D. B.
Corry
,
T. J.
Hollman
,
J.
Kildsgaard
,
R. A.
Wetsel
.
2002
.
Absence of the complement anaphylatoxin C3a receptor suppresses Th2 effector functions in a murine model of pulmonary allergy.
J. Immunol.
169
:
5926
5933
.
15
Gerardy-Schahn
,
R.
,
D.
Ambrosius
,
D.
Saunders
,
M.
Casaretto
,
C.
Mittler
,
G.
Karwarth
,
S.
Görgen
,
D.
Bitter-Suermann
.
1989
.
Characterization of C3a receptor-proteins on guinea pig platelets and human polymorphonuclear leukocytes.
Eur. J. Immunol.
19
:
1095
1102
.
16
Martin
,
U.
,
D.
Bock
,
L.
Arseniev
,
M. A.
Tornetta
,
R. S.
Ames
,
W.
Bautsch
,
J.
Köhl
,
A.
Ganser
,
A.
Klos
.
1997
.
The human C3a receptor is expressed on neutrophils and monocytes, but not on B or T lymphocytes.
J. Exp. Med.
186
:
199
207
.
17
Asgari
,
E.
,
G.
Le Friec
,
H.
Yamamoto
,
E.
Perucha
,
S. S.
Sacks
,
J.
Köhl
,
H. T.
Cook
,
C.
Kemper
.
2013
.
C3a modulates IL-1β secretion in human monocytes by regulating ATP efflux and subsequent NLRP3 inflammasome activation.
Blood
122
:
3473
3481
.
18
Daffern
,
P. J.
,
P. H.
Pfeifer
,
J. A.
Ember
,
T. E.
Hugli
.
1995
.
C3a is a chemotaxin for human eosinophils but not for neutrophils. I. C3a stimulation of neutrophils is secondary to eosinophil activation.
J. Exp. Med.
181
:
2119
2127
.
19
Elsner
,
J.
,
M.
Oppermann
,
W.
Czech
,
G.
Dobos
,
E.
Schöpf
,
J.
Norgauer
,
A.
Kapp
.
1994
.
C3a activates reactive oxygen radical species production and intracellular calcium transients in human eosinophils.
Eur. J. Immunol.
24
:
518
522
.
20
Bischoff
,
S. C.
,
A. L.
de Weck
,
C. A.
Dahinden
.
1990
.
Interleukin 3 and granulocyte/macrophage-colony-stimulating factor render human basophils responsive to low concentrations of complement component C3a.
Proc. Natl. Acad. Sci. USA
87
:
6813
6817
.
21
Kretzschmar
,
T.
,
A.
Jeromin
,
C.
Gietz
,
W.
Bautsch
,
A.
Klos
,
J.
Köhl
,
G.
Rechkemmer
,
D.
Bitter-Suermann
.
1993
.
Chronic myelogenous leukemia-derived basophilic granulocytes express a functional active receptor for the anaphylatoxin C3a.
Eur. J. Immunol.
23
:
558
561
.
22
Legler
,
D. F.
,
M.
Loetscher
,
S. A.
Jones
,
C. A.
Dahinden
,
M.
Arock
,
B.
Moser
.
1996
.
Expression of high- and low-affinity receptors for C3a on the human mast cell line, HMC-1.
Eur. J. Immunol.
26
:
753
758
.
23
Nilsson
,
G.
,
M.
Johnell
,
C. H.
Hammer
,
H. L.
Tiffany
,
K.
Nilsson
,
D. D.
Metcalfe
,
A.
Siegbahn
,
P. M.
Murphy
.
1996
.
C3a and C5a are chemotaxins for human mast cells and act through distinct receptors via a pertussis toxin-sensitive signal transduction pathway.
J. Immunol.
157
:
1693
1698
.
24
Chen
,
N. J.
,
C.
Mirtsos
,
D.
Suh
,
Y. C.
Lu
,
W. J.
Lin
,
C.
McKerlie
,
T.
Lee
,
H.
Baribault
,
H.
Tian
,
W. C.
Yeh
.
2007
.
C5L2 is critical for the biological activities of the anaphylatoxins C5a and C3a.
Nature
446
:
203
207
.
25
Wu
,
M. C.
,
F. H.
Brennan
,
J. P.
Lynch
,
S.
Mantovani
,
S.
Phipps
,
R. A.
Wetsel
,
M. J.
Ruitenberg
,
S. M.
Taylor
,
T. M.
Woodruff
.
2013
.
The receptor for complement component C3a mediates protection from intestinal ischemia-reperfusion injuries by inhibiting neutrophil mobilization.
Proc. Natl. Acad. Sci. USA
110
:
9439
9444
.
26
Li
,
K.
,
K. J.
Anderson
,
Q.
Peng
,
A.
Noble
,
B.
Lu
,
A. P.
Kelly
,
N.
Wang
,
S. H.
Sacks
,
W.
Zhou
.
2008
.
Cyclic AMP plays a critical role in C3a-receptor-mediated regulation of dendritic cells in antigen uptake and T-cell stimulation.
Blood
112
:
5084
5094
.
27
Werfel
,
T.
,
K.
Kirchhoff
,
M.
Wittmann
,
G.
Begemann
,
A.
Kapp
,
F.
Heidenreich
,
O.
Götze
,
J.
Zwirner
.
2000
.
Activated human T lymphocytes express a functional C3a receptor.
J. Immunol.
165
:
6599
6605
.
28
Cravedi
,
P.
,
J.
Leventhal
,
P.
Lakhani
,
S. C.
Ward
,
M. J.
Donovan
,
P. S.
Heeger
.
2013
.
Immune cell-derived C3a and C5a costimulate human T cell alloimmunity.
Am. J. Transplant.
13
:
2530
2539
.
29
Kwan
,
W. H.
,
W.
van der Touw
,
E.
Paz-Artal
,
M. O.
Li
,
P. S.
Heeger
.
2013
.
Signaling through C5a receptor and C3a receptor diminishes function of murine natural regulatory T cells.
J. Exp. Med.
210
:
257
268
.
30
Liszewski
,
M. K.
,
M.
Kolev
,
G.
Le Friec
,
M.
Leung
,
P. G.
Bertram
,
A. F.
Fara
,
M.
Subias
,
M. C.
Pickering
,
C.
Drouet
,
S.
Meri
, et al
.
2013
.
Intracellular complement activation sustains T cell homeostasis and mediates effector differentiation.
Immunity
39
:
1143
1157
.
31
Strainic
,
M. G.
,
J.
Liu
,
D.
Huang
,
F.
An
,
P. N.
Lalli
,
N.
Muqim
,
V. S.
Shapiro
,
G. R.
Dubyak
,
P. S.
Heeger
,
M. E.
Medof
.
2008
.
Locally produced complement fragments C5a and C3a provide both costimulatory and survival signals to naive CD4+ T cells.
Immunity
28
:
425
435
.
32
Braun
,
M. C.
,
R. Y.
Reins
,
T. B.
Li
,
T. J.
Hollmann
,
R.
Dutta
,
W. A.
Rick
,
B. B.
Teng
,
B.
Ke
.
2004
.
Renal expression of the C3a receptor and functional responses of primary human proximal tubular epithelial cells.
J. Immunol.
173
:
4190
4196
.
33
Kiafard
,
Z.
,
T.
Tschernig
,
S.
Schweyer
,
A.
Bley
,
D.
Neumann
,
J.
Zwirner
.
2007
.
Use of monoclonal antibodies to assess expression of anaphylatoxin receptors in tubular epithelial cells of human, murine and rat kidneys.
Immunobiology
212
:
129
139
.
34
Tschernig
,
T.
,
Z.
Kiafard
,
C.
Dibbert
,
D.
Neumann
,
J.
Zwirner
.
2007
.
Use of monoclonal antibodies to assess expression of anaphylatoxin receptors in rat and murine models of lung inflammation.
Exp. Toxicol. Pathol.
58
:
419
425
.
35
Engelke
,
C.
,
A. V.
Wiese
,
I.
Schmudde
,
F.
Ender
,
H. A.
Ströver
,
T.
Vollbrandt
,
P.
König
,
Y.
Laumonnier
,
J.
Köhl
.
2014
.
Distinct roles of the anaphylatoxins C3a and C5a in dendritic cell–mediated allergic asthma.
J. Immunol.
193
:
5387
5401
.
36
Zhang
,
X.
,
I.
Schmudde
,
Y.
Laumonnier
,
M. K.
Pandey
,
J. R.
Clark
,
P.
König
,
N. P.
Gerard
,
C.
Gerard
,
M.
Wills-Karp
,
J.
Köhl
.
2010
.
A critical role for C5L2 in the pathogenesis of experimental allergic asthma.
J. Immunol.
185
:
6741
6752
.
37
de Felipe
,
P.
,
G. A.
Luke
,
L. E.
Hughes
,
D.
Gani
,
C.
Halpin
,
M. D.
Ryan
.
2006
.
E unum pluribus: multiple proteins from a self-processing polyprotein.
Trends Biotechnol.
24
:
68
75
.
38
Kim
,
J. H.
,
S. R.
Lee
,
L. H.
Li
,
H. J.
Park
,
J. H.
Park
,
K. Y.
Lee
,
M. K.
Kim
,
B. A.
Shin
,
S. Y.
Choi
.
2011
.
High cleavage efficiency of a 2A peptide derived from porcine teschovirus-1 in human cell lines, zebrafish and mice.
PLoS One
6
:
e18556
.
39
Schmudde
,
I.
,
H. A.
Ströver
,
T.
Vollbrandt
,
P.
König
,
C. M.
Karsten
,
Y.
Laumonnier
,
J.
Köhl
.
2013
.
C5a receptor signalling in dendritic cells controls the development of maladaptive Th2 and Th17 immunity in experimental allergic asthma.
Mucosal Immunol.
6
:
807
825
.
40
Weischenfeldt
,
J.
,
B.
Porse
.
2008
.
Bone marrow-derived macrophages (BMM): isolation and applications.
CSH Protoc.
2008
:
pdb.prot5080
.
41
Helft
,
J.
,
J.
Böttcher
,
P.
Chakravarty
,
S.
Zelenay
,
J.
Huotari
,
B. U.
Schraml
,
D.
Goubau
,
C.
Reis e Sousa
.
2015
.
GM-CSF mouse bone marrow cultures comprise a heterogeneous population of CD11c+MHCII+ macrophages and dendritic cells.
Immunity
42
:
1197
1211
.
42
Karsten
,
C. M.
,
Y.
Laumonnier
,
B.
Eurich
,
F.
Ender
,
K.
Bröker
,
S.
Roy
,
A.
Czabanska
,
T.
Vollbrandt
,
J.
Figge
,
J.
Köhl
.
2015
.
Monitoring and cell-specific deletion of C5aR1 using a novel floxed GFP-C5aR1 reporter knock-in mouse.
J. Immunol.
194
:
1841
1855
.
43
Karsten
,
C. M.
,
Y.
Laumonnier
,
J.
Köhl
.
2014
.
Functional analysis of C5a effector responses in vitro and in vivo.
Methods Mol. Biol.
1100
:
291
304
.
44
Glovsky
,
M. M.
,
T. E.
Hugli
,
T.
Ishizaka
,
L. M.
Lichtenstein
,
B. W.
Erickson
.
1979
.
Anaphylatoxin-induced histamine release with human leukocytes: studies of C3a leukocyte binding and histamine release.
J. Clin. Invest.
64
:
804
811
.
45
Daley
,
J. M.
,
A. A.
Thomay
,
M. D.
Connolly
,
J. S.
Reichner
,
J. E.
Albina
.
2008
.
Use of Ly6G-specific monoclonal antibody to deplete neutrophils in mice.
J. Leukoc. Biol.
83
:
64
70
.
46
Guglietta
,
S.
,
A.
Chiavelli
,
E.
Zagato
,
C.
Krieg
,
S.
Gandini
,
P. S.
Ravenda
,
B.
Bazolli
,
B.
Lu
,
G.
Penna
,
M.
Rescigno
.
2016
.
Coagulation induced by C3aR-dependent NETosis drives protumorigenic neutrophils during small intestinal tumorigenesis.
Nat. Commun.
7
:
11037
.
47
Bautsch
,
W.
,
H. G.
Hoymann
,
Q.
Zhang
,
I.
Meier-Wiedenbach
,
U.
Raschke
,
R. S.
Ames
,
B.
Sohns
,
N.
Flemme
,
A.
Meyer zu Vilsendorf
,
M.
Grove
, et al
.
2000
.
Cutting edge: guinea pigs with a natural C3a-receptor defect exhibit decreased bronchoconstriction in allergic airway disease: evidence for an involvement of the C3a anaphylatoxin in the pathogenesis of asthma.
J. Immunol.
165
:
5401
5405
.
48
Humbles
,
A. A.
,
B.
Lu
,
C. A.
Nilsson
,
C.
Lilly
,
E.
Israel
,
Y.
Fujiwara
,
N. P.
Gerard
,
C.
Gerard
.
2000
.
A role for the C3a anaphylatoxin receptor in the effector phase of asthma.
Nature
406
:
998
1001
.
49
Morris
,
L.
,
C. F.
Graham
,
S.
Gordon
.
1991
.
Macrophages in haemopoietic and other tissues of the developing mouse detected by the monoclonal antibody F4/80.
Development
112
:
517
526
.
50
Plantinga
,
M.
,
M.
Guilliams
,
M.
Vanheerswynghels
,
K.
Deswarte
,
F.
Branco-Madeira
,
W.
Toussaint
,
L.
Vanhoutte
,
K.
Neyt
,
N.
Killeen
,
B.
Malissen
, et al
.
2013
.
Conventional and monocyte-derived CD11b+ dendritic cells initiate and maintain T helper 2 cell-mediated immunity to house dust mite allergen.
Immunity
38
:
322
335
.
51
Rothenberg
,
M. E.
,
S. P.
Hogan
.
2006
.
The eosinophil.
Annu. Rev. Immunol.
24
:
147
174
.
52
Wu
,
D.
,
A. B.
Molofsky
,
H. E.
Liang
,
R. R.
Ricardo-Gonzalez
,
H. A.
Jouihan
,
J. K.
Bando
,
A.
Chawla
,
R. M.
Locksley
.
2011
.
Eosinophils sustain adipose alternatively activated macrophages associated with glucose homeostasis.
Science
332
:
243
247
.
53
Guilliams
,
M.
,
F.
Ginhoux
,
C.
Jakubzick
,
S. H.
Naik
,
N.
Onai
,
B. U.
Schraml
,
E.
Segura
,
R.
Tussiwand
,
S.
Yona
.
2014
.
Dendritic cells, monocytes and macrophages: a unified nomenclature based on ontogeny.
Nat. Rev. Immunol.
14
:
571
578
.
54
Mayer
,
C. T.
,
P.
Ghorbani
,
A.
Nandan
,
M.
Dudek
,
C.
Arnold-Schrauf
,
C.
Hesse
,
L.
Berod
,
P.
Stüve
,
F.
Puttur
,
M.
Merad
,
T.
Sparwasser
.
2014
.
Selective and efficient generation of functional Batf3-dependent CD103+ dendritic cells from mouse bone marrow.
Blood
124
:
3081
3091
.
55
Hoffmann
,
F.
,
F.
Ender
,
I.
Schmudde
,
I. P.
Lewkowich
,
J.
Köhl
,
P.
König
,
Y.
Laumonnier
.
2016
.
Origin, localization, and immunoregulatory properties of pulmonary phagocytes in allergic asthma.
Front. Immunol.
7
:
107
.
56
Varol
,
C.
,
A.
Vallon-Eberhard
,
E.
Elinav
,
T.
Aychek
,
Y.
Shapira
,
H.
Luche
,
H. J.
Fehling
,
W. D.
Hardt
,
G.
Shakhar
,
S.
Jung
.
2009
.
Intestinal lamina propria dendritic cell subsets have different origin and functions.
Immunity
31
:
502
512
.
57
Gilliet
,
M.
,
W.
Cao
,
Y. J.
Liu
.
2008
.
Plasmacytoid dendritic cells: sensing nucleic acids in viral infection and autoimmune diseases.
Nat. Rev. Immunol.
8
:
594
606
.
58
Nakano
,
H.
,
M.
Yanagita
,
M. D.
Gunn
.
2001
.
CD11c+B220+Gr-1+ cells in mouse lymph nodes and spleen display characteristics of plasmacytoid dendritic cells.
J. Exp. Med.
194
:
1171
1178
.
59
Köhl
,
J.
,
R.
Baelder
,
I. P.
Lewkowich
,
M. K.
Pandey
,
H.
Hawlisch
,
L.
Wang
,
J.
Best
,
N. S.
Herman
,
A. A.
Sproles
,
J.
Zwirner
, et al
.
2006
.
A regulatory role for the C5a anaphylatoxin in type 2 immunity in asthma.
J. Clin. Invest.
116
:
783
796
.
60
Goubier
,
A.
,
B.
Dubois
,
H.
Gheit
,
G.
Joubert
,
F.
Villard-Truc
,
C.
Asselin-Paturel
,
G.
Trinchieri
,
D.
Kaiserlian
.
2008
.
Plasmacytoid dendritic cells mediate oral tolerance.
Immunity
29
:
464
475
.
61
Ghosh
,
A. R.
,
R.
Bhattacharya
,
S.
Bhattacharya
,
T.
Nargis
,
O.
Rahaman
,
P.
Duttagupta
,
D.
Raychaudhuri
,
C. S.
Liu
,
S.
Roy
,
P.
Ghosh
, et al
.
2016
.
Adipose recruitment and activation of plasmacytoid dendritic cells fuel metaflammation.
Diabetes
65
:
3440
3452
.
62
Björck
,
P.
2001
.
Isolation and characterization of plasmacytoid dendritic cells from Flt3 ligand and granulocyte-macrophage colony-stimulating factor-treated mice.
Blood
98
:
3520
3526
.
63
Dunkelberger
,
J.
,
L.
Zhou
,
T.
Miwa
,
W. C.
Song
.
2012
.
C5aR expression in a novel GFP reporter gene knockin mouse: implications for the mechanism of action of C5aR signaling in T cell immunity.
J. Immunol.
188
:
4032
4042
.
64
Kantor
,
A. B.
,
A. M.
Stall
,
S.
Adams
,
L. A.
Herzenberg
,
L. A.
Herzenberg
.
1992
.
Differential development of progenitor activity for three B-cell lineages.
Proc. Natl. Acad. Sci. USA
89
:
3320
3324
.
65
Walker
,
J. A.
,
J. L.
Barlow
,
A. N.
McKenzie
.
2013
.
Innate lymphoid cells—how did we miss them?
Nat. Rev. Immunol.
13
:
75
87
.
66
Qiu
,
J.
,
X.
Guo
,
Z. M.
Chen
,
L.
He
,
G. F.
Sonnenberg
,
D.
Artis
,
Y. X.
Fu
,
L.
Zhou
.
2013
.
Group 3 innate lymphoid cells inhibit T-cell-mediated intestinal inflammation through aryl hydrocarbon receptor signaling and regulation of microflora.
Immunity
39
:
386
399
.
67
Wang
,
H.
,
Y.
Liu
,
Z.
Liu
.
2013
.
Clara cell 10-kD protein in inflammatory upper airway diseases.
Curr. Opin. Allergy Clin. Immunol.
13
:
25
30
.
68
Corbière
,
V.
,
V.
Dirix
,
S.
Norrenberg
,
M.
Cappello
,
M.
Remmelink
,
F.
Mascart
.
2011
.
Phenotypic characteristics of human type II alveolar epithelial cells suitable for antigen presentation to T lymphocytes.
Respir. Res.
12
:
15
.
69
Settmacher
,
B.
,
D.
Bock
,
H.
Saad
,
S.
Gärtner
,
C.
Rheinheimer
,
J.
Köhl
,
W.
Bautsch
,
A.
Klos
.
1999
.
Modulation of C3a activity: internalization of the human C3a receptor and its inhibition by C5a.
J. Immunol.
162
:
7409
7416
.
70
Chao
,
T. H.
,
J. A.
Ember
,
M.
Wang
,
Y.
Bayon
,
T. E.
Hugli
,
R. D.
Ye
.
1999
.
Role of the second extracellular loop of human C3a receptor in agonist binding and receptor function.
J. Biol. Chem.
274
:
9721
9728
.
71
Gupta
,
A.
,
R.
Rezvani
,
M.
Lapointe
,
P.
Poursharifi
,
P.
Marceau
,
S.
Tiwari
,
A.
Tchernof
,
K.
Cianflone
.
2014
.
Downregulation of complement C3 and C3aR expression in subcutaneous adipose tissue in obese women.
PLoS One
9
:
e95478
.
72
Norgauer
,
J.
,
G.
Dobos
,
E.
Kownatzki
,
C.
Dahinden
,
R.
Burger
,
R.
Kupper
,
P.
Gierschik
.
1993
.
Complement fragment C3a stimulates Ca2+ influx in neutrophils via a pertussis-toxin-sensitive G protein.
Eur. J. Biochem.
217
:
289
294
.
73
Zhang
,
X.
,
I. P.
Lewkowich
,
G.
Köhl
,
J. R.
Clark
,
M.
Wills-Karp
,
J.
Köhl
.
2009
.
A protective role for C5a in the development of allergic asthma associated with altered levels of B7-H1 and B7-DC on plasmacytoid dendritic cells.
J. Immunol.
182
:
5123
5130
.
74
Hollmann
,
T. J.
,
D. L.
Haviland
,
J.
Kildsgaard
,
K.
Watts
,
R. A.
Wetsel
.
1998
.
Cloning, expression, sequence determination, and chromosome localization of the mouse complement C3a anaphylatoxin receptor gene.
Mol. Immunol.
35
:
137
148
.
75
Kildsgaard
,
J.
,
T. J.
Hollmann
,
K. W.
Matthews
,
K.
Bian
,
F.
Murad
,
R. A.
Wetsel
.
2000
.
Cutting edge: targeted disruption of the C3a receptor gene demonstrates a novel protective anti-inflammatory role for C3a in endotoxin-shock.
J. Immunol.
165
:
5406
5409
.
76
Mamane
,
Y.
,
C.
Chung Chan
,
G.
Lavallee
,
N.
Morin
,
L. J.
Xu
,
J.
Huang
,
R.
Gordon
,
W.
Thomas
,
J.
Lamb
,
E. E.
Schadt
, et al
.
2009
.
The C3a anaphylatoxin receptor is a key mediator of insulin resistance and functions by modulating adipose tissue macrophage infiltration and activation.
Diabetes
58
:
2006
2017
.
77
Davoust
,
N.
,
J.
Jones
,
P. F.
Stahel
,
R. S.
Ames
,
S. R.
Barnum
.
1999
.
Receptor for the C3a anaphylatoxin is expressed by neurons and glial cells.
Glia
26
:
201
211
.
78
Möller
,
T.
,
C.
Nolte
,
R.
Burger
,
A.
Verkhratsky
,
H.
Kettenmann
.
1997
.
Mechanisms of C5a and C3a complement fragment-induced [Ca2+]i signaling in mouse microglia.
J. Neurosci.
17
:
615
624
.
79
Kiwamoto
,
T.
,
T.
Katoh
,
C. M.
Evans
,
W. J.
Janssen
,
M. E.
Brummet
,
S. A.
Hudson
,
Z.
Zhu
,
M.
Tiemeyer
,
B. S.
Bochner
.
2015
.
Endogenous airway mucins carry glycans that bind Siglec-F and induce eosinophil apoptosis.
J. Allergy Clin. Immunol.
135
:
1329
1340.e1321–1329
.
80
Westphalen
,
K.
,
G. A.
Gusarova
,
M. N.
Islam
,
M.
Subramanian
,
T. S.
Cohen
,
A. S.
Prince
,
J.
Bhattacharya
.
2014
.
Sessile alveolar macrophages communicate with alveolar epithelium to modulate immunity.
Nature
506
:
503
506
.
81
Petering
,
H.
,
J.
Köhl
,
A.
Weyergraf
,
Y.
Dulkys
,
D.
Kimmig
,
R.
Smolarski
,
A.
Kapp
,
J.
Elsner
.
2000
.
Characterization of synthetic C3a analog peptides on human eosinophils in comparison to the native complement component C3a.
J. Immunol.
164
:
3783
3789
.
82
DiScipio
,
R. G.
,
P. J.
Daffern
,
M. A.
Jagels
,
D. H.
Broide
,
P.
Sriramarao
.
1999
.
A comparison of C3a and C5a-mediated stable adhesion of rolling eosinophils in postcapillary venules and transendothelial migration in vitro and in vivo.
J. Immunol.
162
:
1127
1136
.
83
Soruri
,
A.
,
J.
Grigat
,
Z.
Kiafard
,
J.
Zwirner
.
2008
.
Mast cell activation is characterized by upregulation of a functional anaphylatoxin C5a receptor.
BMC Immunol.
9
:
29
.
84
Li
,
E.
,
E. A.
Tako
,
S. M.
Singer
.
2016
.
Complement activation by Giardia duodenalis parasites through the lectin pathway contributes to mast cell responses and parasite control.
Infect. Immun.
84
:
1092
1099
.
85
Schäfer
,
B.
,
A. M.
Piliponsky
,
T.
Oka
,
C. H.
Song
,
N. P.
Gerard
,
C.
Gerard
,
M.
Tsai
,
J.
Kalesnikoff
,
S. J.
Galli
.
2013
.
Mast cell anaphylatoxin receptor expression can enhance IgE-dependent skin inflammation in mice.
J. Allergy Clin. Immunol.
131
:
541
548.e541–549
.
86
Hartmann
,
K.
,
B. M.
Henz
,
S.
Krüger-Krasagakes
,
J.
Köhl
,
R.
Burger
,
S.
Guhl
,
I.
Haase
,
U.
Lippert
,
T.
Zuberbier
.
1997
.
C3a and C5a stimulate chemotaxis of human mast cells.
Blood
89
:
2863
2870
.
87
Venkatesha
,
R. T.
,
E.
Berla Thangam
,
A. K.
Zaidi
,
H.
Ali
.
2005
.
Distinct regulation of C3a-induced MCP-1/CCL2 and RANTES/CCL5 production in human mast cells by extracellular signal regulated kinase and PI3 kinase.
Mol. Immunol.
42
:
581
587
.
88
Ahamed
,
J.
,
R. T.
Venkatesha
,
E. B.
Thangam
,
H.
Ali
.
2004
.
C3a enhances nerve growth factor-induced NFAT activation and chemokine production in a human mast cell line, HMC-1.
J. Immunol.
172
:
6961
6968
.
89
Kirshenbaum
,
A. S.
,
C.
Akin
,
Y.
Wu
,
M.
Rottem
,
J. P.
Goff
,
M. A.
Beaven
,
V. K.
Rao
,
D. D.
Metcalfe
.
2003
.
Characterization of novel stem cell factor responsive human mast cell lines LAD 1 and 2 established from a patient with mast cell sarcoma/leukemia; activation following aggregation of FcεRI or FcγRI.
Leuk. Res.
27
:
677
682
.
90
Erdei
,
A.
,
I.
Pecht
.
1996
.
Complement peptides and mast cell triggering.
Immunol. Lett.
54
:
109
112
.
91
Gutzmer
,
R.
,
M.
Lisewski
,
J.
Zwirner
,
S.
Mommert
,
C.
Diesel
,
M.
Wittmann
,
A.
Kapp
,
T.
Werfel
.
2004
.
Human monocyte-derived dendritic cells are chemoattracted to C3a after up-regulation of the C3a receptor with interferons.
Immunology
111
:
435
443
.
92
Li
,
K.
,
H.
Fazekasova
,
N.
Wang
,
Q.
Peng
,
S. H.
Sacks
,
G.
Lombardi
,
W.
Zhou
.
2012
.
Functional modulation of human monocytes derived DCs by anaphylatoxins C3a and C5a.
Immunobiology
217
:
65
73
.
93
Coombes
,
J. L.
,
K. R.
Siddiqui
,
C. V.
Arancibia-Cárcamo
,
J.
Hall
,
C. M.
Sun
,
Y.
Belkaid
,
F.
Powrie
.
2007
.
A functionally specialized population of mucosal CD103+ DCs induces Foxp3+ regulatory T cells via a TGF-β and retinoic acid-dependent mechanism.
J. Exp. Med.
204
:
1757
1764
.
94
Gross
,
M.
,
T. M.
Salame
,
S.
Jung
.
2015
.
Guardians of the gut—murine intestinal macrophages and dendritic cells.
Front. Immunol.
6
:
254
.
95
Laudisi
,
F.
,
R.
Spreafico
,
M.
Evrard
,
T. R.
Hughes
,
B.
Mandriani
,
M.
Kandasamy
,
B. P.
Morgan
,
B.
Sivasankar
,
A.
Mortellaro
.
2013
.
Cutting edge: the NLRP3 inflammasome links complement-mediated inflammation and IL-1β release.
J. Immunol.
191
:
1006
1010
.
96
Kobayashi
,
T.
,
K.
Iijima
,
J. L.
Checkel
,
H.
Kita
.
2013
.
IL-1 family cytokines drive Th2 and Th17 cells to innocuous airborne antigens.
Am. J. Respir. Cell Mol. Biol.
49
:
989
998
.
97
Esnault
,
S.
,
E. A.
Kelly
,
L. M.
Nettenstrom
,
E. B.
Cook
,
C. M.
Seroogy
,
N. N.
Jarjour
.
2012
.
Human eosinophils release IL-1β and increase expression of IL-17A in activated CD4+ T lymphocytes.
Clin. Exp. Allergy
42
:
1756
1764
.
98
Melendi
,
G. A.
,
S. J.
Hoffman
,
R. A.
Karron
,
P. M.
Irusta
,
F. R.
Laham
,
A.
Humbles
,
B.
Schofield
,
C. H.
Pan
,
R.
Rabold
,
B.
Thumar
, et al
.
2007
.
C5 modulates airway hyperreactivity and pulmonary eosinophilia during enhanced respiratory syncytial virus disease by decreasing C3a receptor expression.
J. Virol.
81
:
991
999
.
99
Thangam
,
E. B.
,
R. T.
Venkatesha
,
A. K.
Zaidi
,
K. L.
Jordan-Sciutto
,
D. A.
Goncharov
,
V. P.
Krymskaya
,
Y.
Amrani
,
R. A.
Panettieri
Jr.
,
H.
Ali
.
2005
.
Airway smooth muscle cells enhance C3a-induced mast cell degranulation following cell-cell contact.
FASEB J.
19
:
798
800
.
100
Ghannam
,
A.
,
J. L.
Fauquert
,
C.
Thomas
,
C.
Kemper
,
C.
Drouet
.
2014
.
Human complement C3 deficiency: Th1 induction requires T cell-derived complement C3a and CD46 activation.
Mol. Immunol.
58
:
98
107
.
101
Strainic
,
M. G.
,
E. M.
Shevach
,
F.
An
,
F.
Lin
,
M. E.
Medof
.
2013
.
Absence of signaling into CD4+ cells via C3aR and C5aR enables autoinductive TGF-β1 signaling and induction of Foxp3+ regulatory T cells.
Nat. Immunol.
14
:
162
171
.
102
Melo-Gonzalez
,
F.
,
M. R.
Hepworth
.
2017
.
Functional and phenotypic heterogeneity of group 3 innate lymphoid cells.
Immunology
150
:
265
275
.
103
Yuan
,
X.
,
M.
Shan
,
R.
You
,
M. V.
Frazier
,
M. J.
Hong
,
R. A.
Wetsel
,
S.
Drouin
,
A.
Seryshev
,
L. Z.
Song
,
L.
Cornwell
, et al
.
2015
.
Activation of C3a receptor is required in cigarette smoke-mediated emphysema.
Mucosal Immunol.
8
:
874
885
.
104
Ducruet
,
A. F.
,
B. G.
Hassid
,
W. J.
Mack
,
S. A.
Sosunov
,
M. L.
Otten
,
D. J.
Fusco
,
Z. L.
Hickman
,
G. H.
Kim
,
R. J.
Komotar
,
J.
Mocco
,
E. S.
Connolly
.
2008
.
C3a receptor modulation of granulocyte infiltration after murine focal cerebral ischemia is reperfusion dependent.
J. Cereb. Blood Flow Metab.
28
:
1048
1058
.
105
Lian
,
H.
,
A.
Litvinchuk
,
A. C.
Chiang
,
N.
Aithmitti
,
J. L.
Jankowsky
,
H.
Zheng
.
2016
.
Astrocyte-microglia cross talk through complement activation modulates amyloid pathology in mouse models of Alzheimer’s disease.
J. Neurosci.
36
:
577
589
.
106
Wenderfer
,
S. E.
,
H.
Wang
,
B.
Ke
,
R. A.
Wetsel
,
M. C.
Braun
.
2009
.
C3a receptor deficiency accelerates the onset of renal injury in the MRL/lpr mouse.
Mol. Immunol.
46
:
1397
1404
.
107
Vasek
,
M. J.
,
C.
Garber
,
D.
Dorsey
,
D. M.
Durrant
,
B.
Bollman
,
A.
Soung
,
J.
Yu
,
C.
Perez-Torres
,
A.
Frouin
,
D. K.
Wilton
, et al
.
2016
.
A complement-microglial axis drives synapse loss during virus-induced memory impairment.
Nature
534
:
538
543
.
108
Hannedouche
,
S.
,
V.
Beck
,
J.
Leighton-Davies
,
M.
Beibel
,
G.
Roma
,
E. J.
Oakeley
,
V.
Lannoy
,
J.
Bernard
,
J.
Hamon
,
S.
Barbieri
, et al
.
2013
.
Identification of the C3a receptor (C3AR1) as the target of the VGF-derived peptide TLQP-21 in rodent cells.
J. Biol. Chem.
288
:
27434
27443
.

The authors have no financial conflicts of interest.

Supplementary data