Charcot–Leyden crystals (CLCs) are Galectin-10 protein crystals that can form after eosinophils degranulate. CLCs can appear and persist in tissues from patients with eosinophilic disorders, such as asthma, allergic reactions, and fungal and helminthic infections. Despite abundant reports of their occurrence in human disease, the inflammatory potential of CLCs has remained unknown. In this article, we show that CLCs induce the release of the proinflammatory cytokine IL-1β upon their phagocytosis by primary human macrophages in vitro. Chemical inhibition and small interfering RNA knockdown of NLRP3 in primary human macrophages abrogated their IL-1β response to CLCs. Using C57BL/6 ASC-mCitrine transgenic inflammasome reporter mice, we showed that the instillation of CLCs into the lungs promoted the assembly of ASC complexes in infiltrating immune cells (neutrophils and inflammatory monocytes) and resulted in IL-1β accumulation into the bronchoalveolar lavage fluid. Our findings reveal that CLCs are recognized by the NLRP3 inflammasome, which may sustain inflammation that follows eosinophilic inflammatory processes.

Modifications in the physicochemical properties of certain proteins, lipids, or metabolites can result in phase transition, leading to crystal formation or nonphysiological aggregation (1, 2). Crystals can arise in tissues when endogenous material, such as minerals, cholesterol, and uric acid, deposit. They can also be introduced from exogenous sources by inhalation (silica, asbestos, airborne particulate matter) or via injections (medical nanomaterials, vaccine adjuvants) (1). Crystalline or aggregated materials are sensed by innate immune cells, which are equipped with germline-encoded pattern recognition signaling receptors (PRRs). Innate immune cells survey tissues and remove accumulated extracellular material, pathogens, and residues of cellular demise. Sensing of crystals induces potent inflammatory responses, a property that has been exploited for adjuvanticity in vaccine formulations. Whereas several membrane-bound PRRs are involved in the recognition and uptake of crystals (3, 4), the cytosolic PRR NLRP3 senses frustrated phagocytosed crystals. NLRP3 activation causes the formation of an inflammasome that leads to activation and release of the proinflammatory cytokine IL-1β (510). IL-1β plays important roles in crystal-driven inflammatory diseases and, together with NLRP3, represents a promising therapeutic target for a range of crystal-mediated diseases (5).

Inflammation occurs not only as a consequence of crystal buildup, but it can itself promote crystallization. During inflammation, immune cells are recruited to the affected tissues. Among them, eosinophils and basophils play important roles in allergic reactions, fungal and helminthic infections, and in the regulation of a variety of autoimmune disorders. These cells can expel highly cytotoxic proteins from their secretory granules, which mediate the marked physiological changes associated with eosinophil and basophil inflammation. A predominant protein present in human eosinophilic and basophilic granules is Galectin-10 (LGASL10), also known as Charcot–Leyden protein (6, 7). Galectin-10 can cluster and form Charcot–Leyden crystals (CLCs), which are colorless elongated hexagonal bipyramidal structures that can reach up to 50 μm in length. Since their first report more than 160 years ago, CLCs have been shown to spontaneously form in vitro from lysates of eosinophils (8) and basophils (9) and have been found to accumulate in tissues from patients with asthma (10), fungal allergic reactions (1113), helminthic infections (14), and myeloid leukemia (1517). Despite abundant reports showing the appearance of CLCs in tissues from patients with eosinophilic disorders, these crystals are still regarded as inert remains of eosinophil activity with unrecognized immune function. Advances in this field have been hindered by the lack of a suitable animal model to study CLC formation in vivo. Mice lack the gene LGASL10, which encodes for Galectin-10. Therefore, it remains to be demonstrated whether CLCs are inflammatory in vitro and in vivo.

In this article, we show that CLCs of variable sizes are promptly phagocytosed by primary human macrophages in vitro, and this leads to the release of the proinflammatory cytokine IL-1β. CLC-induced IL-1β in human macrophages occurred through the activation of the NLRP3 inflammasome. Furthermore, we show that CLCs promoted inflammation in vivo, characterized by the tissue infiltration of neutrophils and inflammatory monocytes, phagocytosis of CLCs by recruited immune cells, and IL-1β release in the tissue lavage fluid. Additionally, CLCs induced the assembly of inflammasomes (ASC specks) in vivo after their instillation into the lungs of ASC-mCitrine transgenic (Tg) reporter mice.

Our findings uncover immune stimulatory features of CLCs and shed new light on the likely consequences caused by inflammatory sequelae that follow eosinophil infiltration in tissues.

Ultrapure LPS from Escherichia coli 0111:B4 and nigericin were from Invitrogen, and DRAQ5 was from eBioscience. Cytochalasin D was purchased from Sigma, and CA-074 Me from Enzo Life Sciences. The caspase-1 inhibitor Belnacasan (VX-765) was from Selleck Chemicals. The NLRP3-specific inhibitor CRID3 (also known as Cytokine release inhibitory drug 3 or MCC950) was purchased from Sigma-Aldrich. Homogeneous time resolved fluorescence (HTRF) kits for human IL-1β and TNF-α were from Cisbio Bioassays and were used according to the manufacturer’s instructions. Cholesterol crystals were prepared as previously described (18). Briefly, crystals were prepared from a 2-mg/ml cholesterol solution in 1-propanol. Crystallization was induced by addition of 1.5 vol of endotoxin-free water. Crystals were dried and resuspended in sterile PBS.

C57BL/6 mice were acquired from The Jackson Laboratory (stock no. 000664). C57BL/6 ASC-mCitrine Tg mice [B6.Cg-Gt(ROSA)26Sortm1.1(CAG-Pycard/mCitrine*,−CD2*)Dtg/J] were acquired from The Jackson Laboratory (stock no. 030744). Animals were female and age-matched and were from the C57BL/6 background. Mice were housed in pathogen-free conditions and were handled in accordance with the Guide for the Care and Use of Laboratory Animals of the U.S. National Institutes of Health and the Institutional Animal Care and Use Committee of the University of Massachusetts Medical School.

The AML14.3D10 cell line was kindly gifted from M. Sheehan, C.C. Paul, and M.A. Baumann (Wright State University, Dayton, OH). AML14.3D10 cells were cultured in RPMI 1640 medium (Life Technologies) containing 10% FBS (Invitrogen), 1% penicillin/streptomycin, 1× GlutaMAX, and 1× sodium pyruvate (both from Life Technologies) (complete medium).

Buffy coats from healthy donors were obtained according to protocols accepted by the institutional review board at the University of Bonn (local ethics votes Lfd. Nr. 075/14). Primary human macrophages were obtained through differentiation of CD14+ monocytes in a medium complemented with 500 U/ml rhGM-CSF (ImmunoTools) for 3 d. In brief, human PBMCs were obtained from buffy coats of healthy donors by density gradient centrifugation in Ficoll–Paque PLUS (GE Healthcare). PBMCs were incubated at 4°C with magnetic microbeads conjugated to monoclonal anti-human CD14 Abs (Miltenyi Biotec). CD14+ monocytes were thereby magnetically labeled and isolated using a MACS column placed in a magnetic field as indicated by the manufacturer (Miltenyi Biotec). Monocyte-derived macrophages were cultivated in complete medium during all experiments.

CLCs were isolated following an adapted version of the protocol published elsewhere (19). All steps were carried out at 4°C to avoid protein degradation and under sterile conditions using a class II biosafety cabinet and sterile reagents. At least 600 × 106 AML14.3D10 cells were washed twice with PBS (Life Technologies) and resuspended in hypotonic (0.15% NaCl) PBS complemented with complete protease inhibitor mixture (Roche) (lysis buffer). For every milliliter of cell pellet, 1 ml of lysis buffer was used for resuspension of the cells. Resuspended cells were then lysed at 4°C using a Dounce tissue grinder. Thirty strokes were necessary for lysing >90% of the cells, according to inclusion of trypan blue stain. DNA and big membranes were removed by centrifugation of the lysates at 1000 × g for 10 min at 4°C. The pellets were resuspended in 500 μl of lysis buffer, and the suspension was again centrifuged at 1000 × g for 10 min at 4°C to recover any soluble protein that could be trapped within the pellet of membranes. The supernatants from both centrifugation steps were combined and further centrifuged at 40,000 × g for 20 min at 4°C to remove organelle membranes and cellular debris. The pellets were discarded, and supernatants were centrifuged again at 40,000 × g for 30 min at 4°C. The supernatants were combined, and the volume was reduced to ∼1 ml by centrifugation at 3200 × g in an Amicon Ultra-15 centrifugal filter with a membrane with a nominal m.w. limit of 10 kDa (Millipore). The resultant concentrated protein lysate contained CLCs together with the remainder of the cellular membranes and other protein precipitates. CLCs were separated by centrifugation at 750 × g for 5 min at 4°C. Supernatants were kept for 16 h at 4°C, and they were later on examined for the existence of further CLCs. This second set of CLCs was separated by centrifugation at 750 × g for 5 min at 4°C and mixed with the previous set. Combined CLCs were resuspended in cold PBS and thoroughly washed (a minimum of six times) until the CLCs were mostly free of contaminants when observed under the microscope. After all washing steps, CLCs were counted and diluted in complete medium to be used as stimulus for in vitro experiments, or in PBS for in vivo experiments.

Protein-enriched samples during the process of CLC generation were mixed with NuPAGE LDS Sample Buffer (4×) and with NuPAGE Sample Reducing Agent (10×) (both from Thermo Fisher Scientific) followed by heating at 85°C for 10 min to denature and reduce all the proteins present in the samples.

Denatured and reduced proteins were separated in a NuPAGE 12% Bis-Tris protein gel (Thermo Fisher Scientific) under electrophoretic conditions in MES buffer. After electrophoresis, proteins in the gel were stained with Coomassie Brilliant Blue R-250 dye (Thermo Fisher Scientific) for 10 min. Gels were destained in destaining solution (15% isopropanol plus 10% acetone in water) for 16 h. Alternatively, proteins in the gel were transferred into an Immobilon-FL PVDF membrane (Millipore). Nonspecific binding was blocked with 3% BSA in TBS for 1 h, followed by overnight incubation with a rabbit mAb directed against human Galectin-10 (EPR11197, dilution 1:10,000; Abcam) in 3% BSA in TBS with 0.1% Tween 20 (TBS-T). Membranes were washed three times in TBS-T and incubated for 2 h with an IRDye 680RD donkey anti-rabbit Ab (LI-COR) in TBS-T. Membranes were washed two times with TBS-T and a last time with TBS before measuring fluorescent signal in an Odyssey infrared imager (model 9120).

Cell viability of human macrophages was assayed using the CellTiter-Blue Cell Viability Assay (Promega) as indicated by the manufacturer. In short, after experimental treatment, 1 × 105 cells were incubated at 37°C, 5% CO2, and humidified conditions with 50 μl of 1× CellTiter-Blue Reagent. After 1 h, fluorescent emission at 590 nm was measured from the cells and correlated to the number of viable cells.

Mature IL-1β in cell-free supernatants was detected in capillaries by Simple Western size assay using the Wes module (ProteinSimple, San Jose, CA) as indicated by the manufacturer. In brief, four parts of cell-free supernatants were mixed with one part of master mix (ProteinSimple) containing a fluorescently labeled standard and DTT (40 mM) and then heated at 95°C for 5 min. The samples, biotinylated ladder, anti–IL-1β biotinylated detection Ab (no. 840169 from DuoSet Human IL-1 beta/IL-1F2 ELISA kit DY201, 1:50 dilution; R&D systems), streptavidin–HRP, chemiluminescent substrate, and wash buffer were loaded into the microplates containing separation and stacking matrices. After microplate loading, the separation electrophoresis and immunodetection steps took place in the capillary system of the Wes module.

Cells were fixed in PBS containing 4% methanol-free formaldehyde (Thermo Fisher Scientific) for 15 min at room temperature. After fixation, membranes were stained with wheat germ agglutinin (WGA)–Alexa Fluor 555 (Invitrogen) at 5 μg/ml in PBS for 10 min at room temperature. Fixed cells were incubated for 1 h at room temperature with PBS containing 10% goat serum, 1% FBS, and 0.5% Triton X-100 (blocking/permeabilizing buffer) and then incubated overnight at 4°C with 0.1 μg/ml mouse anti-ASC mAb (clone HASC-71; BioLegend) directly labeled with Alexa Fluor 647 or its respective isotype control, also labeled with Alexa Fluor 647. Cells were washed three times with blocking/permeabilizing buffer. Nuclei were stained with DRAQ5 (1:5000; Thermo Fisher Scientific).

Confocal microscopy was combined with fluorescence microscopy using a Leica True Confocal Scanner SP5 Single Molecule Detection confocal system (Leica Microsystems, Wetzlar, Germany). Images were acquired using a 63× objective with a numerical aperture of 1.2 and analyzed using the Volocity 6.01 software (PerkinElmer, Waltham, MA). For better visibility, in some instances, brightness and contrast were adjusted on all images within one experiment and applied equally to isotype controls and staining conditions.

Primary human macrophages were electroporated with small interfering RNAs (siRNAs) using a Neon Transfection System (MPK5000; Invitrogen). For each electroporation reaction, 1.2 × 106 cells (10 μl) were mixed with 75 pmol (1.5 μl) of either Silencer Negative Control siRNA (no. 1: 4390843, no. 2: 4390846; Thermo Fisher Scientific) or Silencer NLRP3-targeted siRNA (4392420, no. 1: s41554, no. 2: s41556; Thermo Fisher Scientific). The mixture was taken up into a 10-μl Neon Pipette Tip (Thermo Fisher Scientific) and electroporated using the following settings: 1400 V, 20 ms, two pulses. Electroporated cells were transferred to RPMI medium supplemented with 10% FCS without antibiotics, counted, and seeded into tissue culture plates in the presence of 125 U/ml rhGM-CSF. After 16 h of incubation, cells were challenged to experimental treatment.

C57BL/6 mice were exposed to crystals by the peritoneal injection of 100 μl of PBS containing either 1 × 106 CLCs, 100 μg of silica crystals, or nothing. After 6 h, mice were euthanized, and 6 ml of lavage solution (RPMI plus 3 mM EDTA) was injected into the peritoneum of mice as described elsewhere (20). After massaging the distended peritoneal cavity, the peritoneal lavage fluid (PELF) was recovered and separated into a cellular fraction and a liquid fraction by centrifugation at 350 × g at 4°C for 5 min. These cells were assayed in flow cytometry to determine neutrophil recruitment and also observed under the microscope (Leica SP5) to identify cells that have phagocytosed CLCs.

C57BL/6 ASC-mCitrine mice were instilled at a dose of 1 mg of silica crystals or 3.5 × 106 CLCs in 60 μl of PBS. The sample suspension was intratracheally instilled once to isoflurane-anesthetized animals by a syringe through a catheter inserted into the airway. A control group was instilled with PBS alone. Mice were sacrified 6 h after instillation, and 3 ml of bronchoalveolar lavage fluid (BALF) was collected to determine the total cell count, neutrophil (7AAD, CD11b+, Ly6G+) counts, and cytokine levels by HTRF.

Cells recovered in the PELF or BALF were resuspended in 2 ml of staining solution (PBS plus 2% FCS) and counted with a hemocytometer. For neutrophil and monocyte identification, 250 μl of cells was stained with fluorochrome-labeled Abs against CD11b (clone M1/70; BioLegend), Ly-6G (clone 1A8; BD Biosciences), and Ly-6C (clone ER-MP20; Bio-Rad Laboratories), or their respective fluorochrome-labeled isotype Ig. After staining with 7AAD (BD Biosciences) for exclusion of dead cells, cells were analyzed in a MACSQuant Analyzer 10 flow cytometer.

The liquid fraction of the BALF was concentrated using Amicon Ultra-2 ml 10K centrifugal filters. The amounts of mouse IL-1β and mouse TNF-α in the concentrated BALF were quantified using an HTRF assay kit (Cisbio Bioassays) and normalized to the volume of concentrated lavage.

Unless otherwise stated, all data are pooled from a minimum of three independent experiments carried out on macrophages from different donors. For clarity, individual experiments are depicted as symbols in graphs. Each symbol represents average values from technical triplicates from an individual donor. Unless indicated otherwise in figure legends all data are graphed as either box plots showing median and quartiles, or floating bars showing the mean and minimum to maximum values from pooled experiments. The significance of differences between groups was evaluated by one-way ANOVA with repeated measures with Holm–Sidak postcomparison test and Geisser–Greenhouse correction. Statistical analysis was carried out with GraphPad Prism (version 6.0). Data were considered significant when p < 0.05 (*), 0.01 (**), 0.001 (***), or 0.0001 (****).

To investigate the inflammatory potential of CLCs, we produced CLCs from whole-cell lysates of AML14.3D10 cells, a subclone of the AML14 cell line that was established from a 68-y-old patient with FAB M2 acute myeloid leukemia (21). Both the parental AML14 cell line and the AML14.3D10 subclone are established tools for the study of eosinophil biology (22). We produced CLCs in vitro by incubating precleared lysates of AML14.3D10 cells at 4°C for 16 h (Fig. 1A, Supplemental Fig. 1A) as previously described (19). The resulting crystals displayed the hexagonal bipyramidal morphology characteristic of CLCs (Fig. 1B). Coomassie Brilliant Blue staining of a gel loaded with SDS-denatured crystals revealed that CLC preparations contained mainly one protein (Fig. 1C, Supplemental Fig. 1A), and immunoblotting with specific anti–Galecin-10 mAb showed that the CLC preparations were composed of Galectin-10 (Fig. 1D, Supplemental Fig. 1A).

FIGURE 1.

Production and characterization of CLCs. (A) Schematics of the generation of CLCs from AML14.3D10 cells. (B) Wide-field imaging of a representative pure CLC fraction generated as shown in (A). Scale bar, 400 μm. Insert is an original magnification ×20 of the area outlined at top. (C) Coomassie Brilliant Blue–stained protein gel and (D) immunoblot for Galectin-10 of representative fractions of lysates of AML14.3D10 cells, impure CLCs, and pure CLCs that were generated as shown in (A). Data are representative of two independent experiments.

FIGURE 1.

Production and characterization of CLCs. (A) Schematics of the generation of CLCs from AML14.3D10 cells. (B) Wide-field imaging of a representative pure CLC fraction generated as shown in (A). Scale bar, 400 μm. Insert is an original magnification ×20 of the area outlined at top. (C) Coomassie Brilliant Blue–stained protein gel and (D) immunoblot for Galectin-10 of representative fractions of lysates of AML14.3D10 cells, impure CLCs, and pure CLCs that were generated as shown in (A). Data are representative of two independent experiments.

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We next generated primary human monocyte-derived macrophages (hMDMs) differentiated from CD14+monocytes isolated from the PBMCs from buffy coats of healthy donors. These cells were primed with LPS and exposed to CLCs, and their phagocytic potential against the CLCs was assessed by microscopy. We noticed that hMDMs promptly engulfed CLCs of variable sizes (Fig. 2A, Supplemental Video 1).

FIGURE 2.

CLCs induce the secretion of IL-1β from primary human macrophages. (A) Confocal imaging of LPS-primed (2 ng/ml for 3 h) primary hMDMs stimulated with purified CLCs for 6 h. Plasma membrane (red, WGA–Alexa Fluor 555), nuclei (blue, DRAQ5), and CLCs (green, laser reflection). Scale bars, 4 μm (top panel), 8 μm (bottom panel). Data are from one representative out of three independent experiments. (B) HTRF measurement of IL-1β and TNF-α from the supernatants of hMDMs that were either left untreated, primed with LPS, or primed with LPS and stimulated for 6 h with the indicated ratios of CLCs per macrophage, the indicated concentrations of silica crystals, or cholesterol crystals, or for 1.5 h with nigericin (Nig). Box plots show median and quartile values from four donors pooled from independent experiments with different CLC preparations. Individual buffy coat donors are represented by different symbols. (C) HTRF measurement of IL-1β in hMDMs primed as in (A) and stimulated with CLCs (two per cell) or silica crystals (100 μg/ml) for the indicated time points. Floating bars (with mean and minimum to maximum values) are shown from pooled data from two independent experiments. (D) Cell viability assay of hMDMs treated as in (B) (n = 2). Floating bars (with mean and minimum to maximum values) are shown from pooled data from two independent experiments. (E) Simple Western assay for human IL-1β in cell-free supernatants of hMDMs from the same donors as in (B) (n = 4).

FIGURE 2.

CLCs induce the secretion of IL-1β from primary human macrophages. (A) Confocal imaging of LPS-primed (2 ng/ml for 3 h) primary hMDMs stimulated with purified CLCs for 6 h. Plasma membrane (red, WGA–Alexa Fluor 555), nuclei (blue, DRAQ5), and CLCs (green, laser reflection). Scale bars, 4 μm (top panel), 8 μm (bottom panel). Data are from one representative out of three independent experiments. (B) HTRF measurement of IL-1β and TNF-α from the supernatants of hMDMs that were either left untreated, primed with LPS, or primed with LPS and stimulated for 6 h with the indicated ratios of CLCs per macrophage, the indicated concentrations of silica crystals, or cholesterol crystals, or for 1.5 h with nigericin (Nig). Box plots show median and quartile values from four donors pooled from independent experiments with different CLC preparations. Individual buffy coat donors are represented by different symbols. (C) HTRF measurement of IL-1β in hMDMs primed as in (A) and stimulated with CLCs (two per cell) or silica crystals (100 μg/ml) for the indicated time points. Floating bars (with mean and minimum to maximum values) are shown from pooled data from two independent experiments. (D) Cell viability assay of hMDMs treated as in (B) (n = 2). Floating bars (with mean and minimum to maximum values) are shown from pooled data from two independent experiments. (E) Simple Western assay for human IL-1β in cell-free supernatants of hMDMs from the same donors as in (B) (n = 4).

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The proinflammatory cytokine IL-1β is a key driver of crystal-induced inflammation in several chronic inflammatory diseases (2328). We therefore investigated whether phagocytosis of CLCs resulted in the release of IL-1β by hMDMs. We found that, similar to the well-established inflammasome activators silica and cholesterol crystals, exposure of LPS-primed hMDMs to CLCs resulted in a dose- and time-dependent release of IL-1β from these cells (Fig. 2B, 2C). Similar to reports using other crystals, LPS priming was required for the release of IL-1β by in vitro CLC-stimulated hMDMs (Supplemental Table I). Consistent with inflammasome activation, CLCs induced cell death of hMDMs (Fig. 2D). Immunoblotting for IL-1β confirmed the presence of the cleaved and bioactive form of this cytokine in cell-free supernatants of CLC-stimulated hMDMs (Fig. 2E).

Phagocytosis of crystalline material and insoluble protein aggregates can ultimately result in lysosomal damage and the release of lysosomal content into the cell cytosol. Lysosomal damage can activate NLRP3, which then recruits the adapter protein ASC and caspase-1, a protein complex that enables maturation of pro–IL-1β by caspase-1–mediated proteolysis (23). In line with these findings, pretreatment of hMDMs with the actin polymerization inhibitor cytochalasin D impaired their phagocytic activity and diminished their capacity to release IL-1β in response to CLC stimulation, indicating that phagocytosis is required for CLC-induced IL-1β release by hMDMs (Fig. 3A, 3B). As expected, cytochalasin D inhibited the IL-1β release mediated by silica crystals, whereas the response to nigericin, a soluble inflammasome activator that acts as an antiporter of H+ and K+, remained unaffected.

FIGURE 3.

Phagocytosis of CLCs and lysosomal damage precede IL-1β production from primary human macrophages. (A) Confocal imaging of LPS-primed hMDMs that were pretreated or not with 5 mM of cytochalasin D 30 min before incubation with CLCs (two per cell). Images are representative of three experiments. (B) HTRF measurement of IL-1β from the supernatants of LPS-primed hMDMs that were pretreated with increasing concentrations of cytochalasin D (0, 0.3, 0.6, 1.25, 2.5, or 5 mM). (C) HTRF measurement of IL-1β from the supernatants of LPS-primed hMDMs that were pretreated with increasing concentrations of the cathepsin-B inhibitor CA-074 Me (0, 1.25, 2.5, 5, 10, or 15 mM). Floating bars (with mean and minimum to maximum values) are shown from pooled data from three independent experiments. Each symbol represents the values from different donors (n = 3).

FIGURE 3.

Phagocytosis of CLCs and lysosomal damage precede IL-1β production from primary human macrophages. (A) Confocal imaging of LPS-primed hMDMs that were pretreated or not with 5 mM of cytochalasin D 30 min before incubation with CLCs (two per cell). Images are representative of three experiments. (B) HTRF measurement of IL-1β from the supernatants of LPS-primed hMDMs that were pretreated with increasing concentrations of cytochalasin D (0, 0.3, 0.6, 1.25, 2.5, or 5 mM). (C) HTRF measurement of IL-1β from the supernatants of LPS-primed hMDMs that were pretreated with increasing concentrations of the cathepsin-B inhibitor CA-074 Me (0, 1.25, 2.5, 5, 10, or 15 mM). Floating bars (with mean and minimum to maximum values) are shown from pooled data from three independent experiments. Each symbol represents the values from different donors (n = 3).

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To investigate whether lysosomal damage is involved in CLC-induced inflammasome activation, we pretreated hMDMs with CA-074 Me, a compound that inhibits lysosomal damage–mediated NLRP3 activation (23). CA-074 Me inhibition blocked CLC-induced IL-1β release of hMDMs (Fig. 3C), suggesting that lysosomal damage is a likely mechanism by which CLCs can induce IL-1β secretion by macrophages.

Inflammasome assembly can be visualized by the transition of the adapter protein ASC from a diffused cytosolic localization into a dot-like structure called the ASC speck (24, 25). We therefore performed immunofluorescence staining of human ASC in LPS-primed hMDMs stimulated with CLCs. Using fluorochrome-conjugated anti-human ASC Abs or isotype-matched fluorochrome-conjugated IgGs, we revealed the formation of ASC specks in hMDMs that phagocytosed CLCs in vitro (Fig. 4A).

FIGURE 4.

CLCs activate the NLRP3 inflammasome in human macrophages. (A) Confocal imaging of LPS-primed (2 ng/ml for 3 h) hMDMs stimulated with CLCs (two per macrophage) for 6 h. Cells were fixed and stained directly with Alexa Fluor 647 (AF647)–labeled anti-ASC Ab or equal amounts of AF647-IgG–matched isotype control mAbs. Plasma membrane (red, WGA–Alexa Fluor 555), nucleus (deep blue, Hoechst 34580), ASC (green, AF647), and CLCs (light blue, laser reflection). Scale bars 23.0 μm, insets 5.0 μm. Data are representative of two independent experiments. (B) HTRF measurement of IL-1β in LPS-primed (2 ng/ml for 3 h) human macrophages that were left untreated or further stimulated with CLCs (two per macrophage, 6 h), silica crystals (100 μg/ml, 6 h), cholesterol crystals (250 μg/ml, 6 h), nigericin (10 μM, 1.5 h), or PrgI (2 μg/ml, 3 h) together with LFn-PA (0.5 μg/ml) in the presence of increasing concentrations of the NLRP3 inhibitor CRID3 (0, 2.5, 5, or 10 μM) or the caspase-1 inhibitor VX-765 (0, 5, 10, or 30 μM). Box plots showing median and quartiles are shown from pooled data from four independent experiments. Each symbol represents values from a different donor. The same IL-1β data points for the CLC-treated hMDMs in the absence of inhibitors are represented in (B) and (C). (D) HTRF measurement of IL-1β in hMDMs that were transiently transfected through electroporation with two different scrambled (siRNA Ctrl.1 and 2) or two NLRP3-specific siRNAs (siRNA N3.1 and 2). Cells were stimulated as in (B). Nonelectroporated (Non-Elect.) hMDMs were assessed as control. Floating bars (with mean and minimum to maximum values) are shown from pooled data from at least four independent experiments. Each symbol represents values from a different donor (n = 7).

FIGURE 4.

CLCs activate the NLRP3 inflammasome in human macrophages. (A) Confocal imaging of LPS-primed (2 ng/ml for 3 h) hMDMs stimulated with CLCs (two per macrophage) for 6 h. Cells were fixed and stained directly with Alexa Fluor 647 (AF647)–labeled anti-ASC Ab or equal amounts of AF647-IgG–matched isotype control mAbs. Plasma membrane (red, WGA–Alexa Fluor 555), nucleus (deep blue, Hoechst 34580), ASC (green, AF647), and CLCs (light blue, laser reflection). Scale bars 23.0 μm, insets 5.0 μm. Data are representative of two independent experiments. (B) HTRF measurement of IL-1β in LPS-primed (2 ng/ml for 3 h) human macrophages that were left untreated or further stimulated with CLCs (two per macrophage, 6 h), silica crystals (100 μg/ml, 6 h), cholesterol crystals (250 μg/ml, 6 h), nigericin (10 μM, 1.5 h), or PrgI (2 μg/ml, 3 h) together with LFn-PA (0.5 μg/ml) in the presence of increasing concentrations of the NLRP3 inhibitor CRID3 (0, 2.5, 5, or 10 μM) or the caspase-1 inhibitor VX-765 (0, 5, 10, or 30 μM). Box plots showing median and quartiles are shown from pooled data from four independent experiments. Each symbol represents values from a different donor. The same IL-1β data points for the CLC-treated hMDMs in the absence of inhibitors are represented in (B) and (C). (D) HTRF measurement of IL-1β in hMDMs that were transiently transfected through electroporation with two different scrambled (siRNA Ctrl.1 and 2) or two NLRP3-specific siRNAs (siRNA N3.1 and 2). Cells were stimulated as in (B). Nonelectroporated (Non-Elect.) hMDMs were assessed as control. Floating bars (with mean and minimum to maximum values) are shown from pooled data from at least four independent experiments. Each symbol represents values from a different donor (n = 7).

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In line with a role for the NLRP3 inflammasome in the intracellular sensing of CLCs, specific inhibition of NLRP3 and caspase-1 with the compounds CRID3 (26) and VX-765, respectively, ablated CLC-induced IL-1β release by hMDMs (Fig. 4B, 4C). In contrast, the response to NAIP–NLRC4 inflammasome activation, tested through the delivery of the needle protein of the Salmonella pathogenicity island 1 type III secretion system (PrgI) into the cytosol, was not affected by the NLRP3 inhibitor CRID3, demonstrating specificity. The findings with inhibitors indicate that, similar to silica (23), uric acid (27), and cholesterol crystals (28), CLCs induce IL-1β activation and release from hMDMs by activating the NLRP3 inflammasome.

To confirm that the NLRP3 sensor is involved in the CLC-induced IL-1β response of primary human macrophages, we performed siRNA knockdown experiments targeting NLRP3 transcripts on hMDMs. Cells were electroporated with 75 pmol of two different NLRP3-specific siRNAs (siRNAs N3.1 and N3.2) or equivalent amounts of two control siRNAs (siRNAs Ctrl.1 and 2). NLRP3 knockdown impeded the translation of NLRP3 in eletroporated LPS-treated hMDMs with different efficiencies (Supplemental Fig. 2), without affecting cell viability. Electroporation of hMDMs with both siRNAs directed against NLRP3, but not with control siRNAs, impaired the IL-1β production in response to CLCs and the related NLRP3-activators, silica crystals and nigericin, without interfering with the cellular responses to the NLRC4 activator, PrgI (Fig. 4D). Electroporation alone did not affect the IL-1β response of inflammasome-activated hMDMs. Taken together, these data show that the CLC-induced IL-1β response of human primary macrophages is dependent on the activation of the NLRP3 inflammasome.

One of the main functions of IL-1β is to induce the recruitment of immune cells, such as neutrophils and monocytes, which limits the spread of infection and initiates tissue repair (29). Inflammasome activation in vivo is characterized by strong neutrophil infiltration in tissues (30), and tissue neutrophilia is a hallmark of IL-1–driven autoinflammatory diseases (31, 32). To confirm whether CLCs induce inflammasome activation in vivo and to locally associate CLC uptake with inflammasome activation in the tissue, we performed intratracheal instillation of CLCs into the lungs of ASC-mCitrine Tg reporter mice (33). ASC-mCitrine Tg mice were instilled with either PBS or CLCs. Silica crystals were used as a positive control, as these crystals were reported to cause IL-1–dependent neutrophil infiltration in vivo (23, 34). Six hours after instillation, we assessed the infiltration of neutrophils, formation of mCitrine-fluorescent ASC specks, and production of proinflammatory cytokines in the BALF. CLCs caused a massive infiltration of neutrophils (CD11b+, Ly6G+, Ly6Cint) into the lungs, which could be detected in their BALF (Fig. 5A). Very few or no neutrophils (CD11b+, Ly6G+, Ly6Cint) were found in the BALF of mice instilled with PBS. Of note, imaging of cells recovered from the BALF revealed the presence of neutrophils and monocytes that had phagocytosed CLCs (Fig. 5B), confirming that CLCs are also phagocytosed in vivo. Importantly, ASC-mCitrine specks were assembled in BALF cells that had phagocytosed CLCs. The assembly of ASC-mCitrine specks was further validated with costaining of BALF cells with an anti-ASC Ab directly conjugated to Alexa Fluor 555 (Fig. 5B). In line with the activation of ASC inflammasomes, IL-1β levels were increased in the BALF of mice injected with silica crystals or CLCs compared with PBS-injected animals.

FIGURE 5.

CLCs cause inflammasome activation in vivo. (A) Flow cytometry quantification of live neutrophils (7AAD, CD11b+Ly6G+) and cytokine levels (IL-1β and TNF-α) in the BALF of female (10-wk-old) ASC-mCitrine Tg mice 6 h after intratracheal administration of 60 μl of PBS alone or 60 μl of PBS containing 1 mg of silica crystals or 3.5 × 106 CLCs. Each symbol represents an individual mouse (n = 3); small horizontal lines indicate the mean and SD. (B) Confocal imaging of cells present in the BALF of ASC-mCitrine Tg mice instilled intratracheally with CLCs or silica crystals showing the formation of ASC aggregates in their cytosol. Note that ASC remains mainly evenly distributed in the cytosol of the cells exposed to only PBS. ASC (green, mCitrine), neutrophils (magenta, Ly6G–Alexa Fluor 647 [A647]), anti-ASC (red, anti-ASC–Alexa Fluor 555 [A555]). Scale bars: 23.0 μm, inserts: 6.0 μm. Data are representative of two independent experiments. (C) Flow cytometry quantification of live neutrophils (7AAD, CD11b+Ly6G+) and live inflammatory monocytes (7AAD, CD11b+Ly6C+Ly6G) and IL-1β in the PELF from female (8-wk-old) C57BL/6 WT or age-matched female NLRP3-deficient (Nlrp3−/−) mice injected i.p. with PBS, silica crystals, or CLCs. Each symbol represents an individual mouse; small horizontal lines indicate the mean and SD. Data are pooled from two independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

FIGURE 5.

CLCs cause inflammasome activation in vivo. (A) Flow cytometry quantification of live neutrophils (7AAD, CD11b+Ly6G+) and cytokine levels (IL-1β and TNF-α) in the BALF of female (10-wk-old) ASC-mCitrine Tg mice 6 h after intratracheal administration of 60 μl of PBS alone or 60 μl of PBS containing 1 mg of silica crystals or 3.5 × 106 CLCs. Each symbol represents an individual mouse (n = 3); small horizontal lines indicate the mean and SD. (B) Confocal imaging of cells present in the BALF of ASC-mCitrine Tg mice instilled intratracheally with CLCs or silica crystals showing the formation of ASC aggregates in their cytosol. Note that ASC remains mainly evenly distributed in the cytosol of the cells exposed to only PBS. ASC (green, mCitrine), neutrophils (magenta, Ly6G–Alexa Fluor 647 [A647]), anti-ASC (red, anti-ASC–Alexa Fluor 555 [A555]). Scale bars: 23.0 μm, inserts: 6.0 μm. Data are representative of two independent experiments. (C) Flow cytometry quantification of live neutrophils (7AAD, CD11b+Ly6G+) and live inflammatory monocytes (7AAD, CD11b+Ly6C+Ly6G) and IL-1β in the PELF from female (8-wk-old) C57BL/6 WT or age-matched female NLRP3-deficient (Nlrp3−/−) mice injected i.p. with PBS, silica crystals, or CLCs. Each symbol represents an individual mouse; small horizontal lines indicate the mean and SD. Data are pooled from two independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Close modal

Differently from what was observed with silica crystals, increased levels of TNFα, an inflammasome-independent proinflammatory cytokine, were also observed in the BALF of mice treated with CLCs (Fig. 5A). As CLCs do not naturally form in mice, this observation suggests that injection of CLCs in vivo might activate additional inflammatory signaling pathways. To confirm that and to investigate whether NLRP3 inflammasome is required for CLC-induced inflammation in vivo, we used a well-established mouse peritonitis model (20, 23, 28). We injected PBS, CLCs, or silica crystals into the peritoneal cavity of wild-type (WT) C57BL/6 or NLRP3-deficient (Nlrp3−/−) mice. Similar to the lung instillation model, i.p. injection of CLCs caused peritonitis characterized by massive infiltration of neutrophils (CD11b+, Ly6G+, Ly6Cint) and inflammatory monocytes (CD11b+, Ly6G, Ly6C+) (Fig. 5C, Supplemental Fig. 3A) into the peritoneal cavity. Peritoneal cells were also able to phagocytose CLCs in both WT and Nlrp3−/− mice (Supplemental Fig. 3B). Although Nlrp3−/− mice displayed reduced infiltration of immune cells and production of IL-1β in the peritoneal lavage, these changes did not reach statistical significance between both groups. These data suggest that CLCs may activate other NLRP3-independent inflammatory mechanisms in vivo.

Altogether, our data show that CLCs induce inflammation in vivo and suggest that their accumulation in tissues might be an important driver for chronic tissue inflammation following eosinophil infiltration.

Our study identifies a previously unappreciated inflammatory capacity of CLCs and opens new avenues to consider the immunogenicity and potential relevance of these crystals for common chronic immunological diseases, such as asthma.

CLCs are found in the sputum of asthma patients with eosinophilia (10, 35). Although asthma patients can display heterogeneous phenotypes, an important molecular mechanism of asthma is type 2 inflammation, which is characterized by an increase in IgE titers, the recruitment of eosinophils, and the predominance of Th2 cells, which secrete IL-4, IL-5, and IL-13 (36). Type 2 immunity can also influence the expression of classical proinflammatory cytokines, such as IL-1β, which can assist in the development of chronic inflammation. In line with these observations, endogenous danger-associated molecular patterns, including uric acid crystals, traditionally known as the cause of inflammation in gout, are strong adjuvants for Th2 immunity (37, 38). Indeed, increased IL-1β levels have been reported in asthma, and the number of macrophages expressing IL-1β in the bronchial epithelium of asthma patients is higher in comparison with control subjects (39).

Our findings show that CLCs, which are commonly found in fluids and tissues of patients with eosinophilia, induce the release of the highly proinflammatory cytokine IL-1β. This finding, together with the report of CLCs found in the phagosomes of macrophages in vivo (40, 41), suggests that the uptake of CLCs by macrophages and the subsequent release of IL-1β can be one of the signals that perpetuate and exacerbate inflammation in the tissues of patients with eosinophilic disorders. On that note, IL-1β has been reported as a critical cytokine for the development of a Th2/Th17-predominant subtype in asthma patients (42). Th2/Th17-predominant asthma patients present dual-positive Th2/Th17 cells in their BALF and exhibit a severe form of asthma. Moreover, these patients show increased eosinophil counts in their BALF. It has recently been proposed that signaling through the IL-1R in CD4+ T cells drives the induction of pathogenic Th17 cells involved in the development of autoimmunity (43). Whether CLCs, which are a hallmark of eosinophilic diseases, are present in the BALF of Th2/Th17-predominant asthma patients, and whether their recognition by alveolar macrophages are the cause of the increased levels of IL-1β, is a matter of further research. In this scenario, inhibition of CLC formation, engulfment, or their effects on IL-1β release—potentially by blocking NLRP3 activity—could benefit patients with asthma.

The activation of the NLRP3 inflammasome requires two steps: a priming signal promotes transcription of NLRP3 and IL-1β in myeloid cells, whereas an activation signal engages NLRP3 to form an inflammasome that processes IL-1β into its mature secreted form. Our study reveals that CLCs can act as an activation signal for NLRP3 inflammasome assembly in vitro and in vivo. Whereas in our in vitro system human macrophages needed the priming signal given by LPS treatment, in vivo treatment with CLCs promoted IL-1β release and neutrophil recruitment without a priming step. On the contrary, pulmonary injections with CLCs also induced the release of TNF-α, a cytokine usually secreted upon the priming step. This could be a consequence of other effects of the CLCs in the body. Because of their shape, CLCs could disrupt the tissue upon injection, and this could lead to the release of danger-associated molecular patterns that could provide a priming signal. On this note, it has been recently reported that neutrophil extracellular traps can license macrophages for IL-1β secretion in atherosclerosis (44). Importantly, CLCs have been associated with the formation of extracellular traps in human tissues (45). Furthermore, our experiments in vivo revealed that CLC injection brought about a massive recruitment of neutrophils to the sites of injection. Hence, although not experimentally tested, CLCs could cause inflammasome activation in vivo through both mechanisms, priming NLRP3 and pro–IL-1β through NETosis and further NLRP3 activation through lysosomal damage after their phagocytosis.

As Galectin-10 is not expressed in mouse cells and CLCs do not naturally form in mice, it is not possible at this point to speculate how the results of the injection of CLCs in the murine models used in our study can be extrapolated to the human disease. As CLC buildup in the body might be a slow process, it is not possible to predict how the artificial injection of crystals can be compared, both in quantity and tissue localization.

The results presented in this study expand our current understanding of how protein crystallization contributes to immunogenicity and highlight the need to consider novel therapeutic strategies that neutralize protein crystallization lingering from the activity of granulocytes.

We thank Meghan Sheehan, Cassandra C. Paul, and Michael A. Baumann, from Wright State University for the AML14.3D10 cell line. We thank Maximillian Rothe for technical assistance and Matthew Mangan for fruitful discussions of data and hypothesis. We thank Feng Shao (National Institute of Biological Sciences, Beijing, China) for the plasmid encoding the PrgI-protective Ag conjugate protein, as well as Matthias Geyer, Matthew Mangan, and David Fußhöller for the purified PrgI protein used in this study to activate the NLRC4 inflammasome.

The online version of this article contains supplemental material.

Abbreviations used in this article:

     
  • BALF

    bronchoalveolar lavage fluid

  •  
  • CLC

    Charcot–Leyden crystal

  •  
  • hMDM

    human monocyte-derived macrophage

  •  
  • HTRF

    homogeneous time resolved fluorescence

  •  
  • PELF

    peritoneal lavage fluid

  •  
  • PRR

    pattern recognition signaling receptor

  •  
  • siRNA

    small interfering RNA

  •  
  • TBS-T

    TBS with 0.1% Tween 20

  •  
  • Tg

    transgenic

  •  
  • WGA

    wheat germ agglutinin

  •  
  • WT

    wild-type.

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The authors have no financial conflicts of interest.