Spectrum and Mechanisms of Inflammasome Activation by Chitosan

Chitosan, a β-(1, 4)-linked polymer of glucosamine, is the deacetylated derivative of chitin, a β-(1, 4)-linked polymer of N-acetylglucosamine. Chitosan is not as prevalent naturally as chitin, although chitin deacetylases, which catalyze conversion of chitin to chitosan, are present in some medically important fungi such as Cryptococcus neoformans and members of the Zygomycetes (1, 2). Chitin is an essential component of fungal cell walls, as well as a major component in crustacean shells, insect exoskeletons, and some parasites, including helminths and protozoa (3–9). Human exposure to these polysaccharides, particularly chitosan, may occur not only during fungal infection, but may arise as a result of their presence in pharmaceutical and commercial applications such as gene and drug delivery constructs, tissue scaffolds, and wound dressings (10–13).

We previously found that chitosan, but not chitin, activates the NOD-like receptor family, pyrin domain containing 3 (NLRP3) inflammasome of bone marrow–derived macrophages (BMMΦ) (14). The NLRP3 inflammasome is a cytosolic complex containing NLRP3, the adaptor molecule apoptosis-associated specklike protein containing a caspase recruitment domain, and caspase-1. Activation is a two-step process with the first step priming the system and resulting in an upregulation of both pro–IL-1β and NLRP3 (15), and the second step inducing caspase-1–dependent cleavage of pro–IL-1β to the active form of IL-1β. The NLRP3 inflammasome has been shown to be activated by a wide variety of stimuli such as ATP, amyloid-β, alum, silica, and nigericin, as well as a variety of fungi, bacteria, and viruses (16). Unlike other described inflammasomes with more specific stimuli, such as AIM2 with DNA (17), and IPAF with flagellin (18), the NLRP3 inflammasome is unlikely to be activated by direct interaction with each of its varied activators.

Although BMMΦ have been the most often studied cell type by inflammasome researchers, other proinflammatory cell types have also been investigated. Macrophages are polarized between classically activated macrophage (M1) and alternatively activated macrophage (M2) phenotypes. M1 macrophages are generally considered proinflammatory, whereas M2 macrophages are considered anti-inflammatory; however, there is reversible plasticity between the phenotypes, and some macrophages exhibit intermediate polarities (19). M1 macrophages have been shown to have a strong inflammasome response, which diminishes as macrophages become polarized toward intermediate and M2 phenotypes (20). Similar to cultured cells, primary cells such as peritoneal macrophages have also been shown to have strong inflammasome responses (21). Activation of the inflammasome in murine dendritic cells (DCs) may be an important intermediary between the innate immune response and the adaptive immune response. DC activation is crucial for vaccine adjuvants to stimulate protective adaptive immunity (22), and the IL-1β produced by DCs is required for the optimal priming of T cells (23). Many parallels exist between mouse and human cell inflammasome activation. However, one important difference is that human blood monocytes have constitutively active caspase-1 and can be stimulated by LPS alone to secrete IL-1β (24).

Three mechanisms for NLRP3 inflammasome activation have been proposed: K⁺ efflux, reactive oxygen species (ROS) generation, and lysosomal destabilization. K⁺ efflux has been shown to be required for NLRP3 inflammasome activation by many different stimuli. This model was first described for ATP, with ATP-mediated activation of the NLRP3 inflammasome being dependent upon activation of P2X7, the ATP-gated ion channel, which triggers rapid K⁺ efflux (25). This K⁺ efflux is then somehow sensed, thereby activating the NLRP3 inflammasome. The second proposed mechanism of activation involves NLRP3 functioning as a more general sensor of cellular stress by recognizing and being activated by ROS. ROS involvement in the activation of the inflammasome has been shown for all NLRP3 stimuli tested (16), and recent work has suggested that the important ROS source for NLRP3 inflammasome activation is mitochondrial in origin (26, 27). ROS generation is frequently accompanied by K⁺ efflux, and one may trigger the other, combining to activate the NLRP3 inflammasome (28). In the third model, particulate stimuli are phagocytosed, leading to phagosomal maturation and lysosomal fusion, ultimately resulting in phagosomal destabilization and lysosomal rupture. With the destabilization of the lysosome, cathepsins including cathepsin B are released into the cytoplasm, resulting in activation of the NLRP3 inflammasome (29). Additional work on how these mechanisms work in relation to each other is still needed.

The immunostimulatory properties of chitin and chitosan remain poorly understood. These polysaccharides have been characterized as relatively inert, proinflammatory, and proallergenic in different reports (14, 30–34). Chitin has been shown to induce an allergic response consisting of an accumulation of eosinophils and basophils expressing IL-4 and alternatively activated macrophages (31). Conversely, in another report, chitin downregulated the allergic response to ragweed in mice (35). Possible explanations for the disparate findings include the sources of chitin (e.g., shrimp, crab, fungal), the manufacturing processes (which, in turn, could affect the degree of deacetylation and tertiary structure), and the presence of contaminants (36–38). Another possible explanation for the varied immunological response is particle size. The importance of size has been suggested by studies demonstrating differential stimulation of TNF-α and IL-10 by size-fractionated chitin. Particles of intermediate size (40–70 μm) induced just TNF-α, whereas smaller particles (<40 μm) induced both TNF-α and IL-10 (39).

In this study, we assayed chitosan-induced IL-1β release in mouse macrophages with an M1, intermediate, or M2 phenotype, DCs, and peritoneal cells, as well as human PBMCs. For all cell types tested, chitosan, but not chitin, induced IL-1β release. We then analyzed the role of each of the proposed NLRP3 inflammasome activation mechanisms described earlier. We found evidence that all three mechanisms—K⁺ efflux, ROS, and lysosomal destabilization—participate in the NLRP3 inflammasome activation by chitosan. Finally, with use of a multiplex assay, we determined that purified chitosan and chitin are relatively weak inducers of cytokines and chemokines from unprimed BMMΦ.

All materials were obtained from Sigma-Aldrich unless otherwise stated. Ultrapure LPS (free of TLR2-stimulating lipopeptides) was treated with deoxycholate twice followed by phenol extraction and ethanol precipitation (40) to further purify the original Sigma-Aldrich stock (catalog #L2630). Chitosan was obtained from Primex (ChitoClear, high m.w. shrimp chitosan, 76% deacetylated). Complete media are defined as RPMI 1640 media (Invitrogen Life Technologies) supplemented with 10% heat-inactivated FBS (Tissue Culture Biologicals), 2 mM l-glutamine (Invitrogen), 100 U/ml penicillin, and 100 μg/ml streptomycin. Cell culture was at 37°C in humidified air supplemented with 5% CO₂. All experiments were performed under conditions designed to minimize endotoxin contamination.

BMMΦ were generated as described previously (41). In brief, bone marrow was extracted from the femurs and tibiae of wild type C57BL/6 mice (The Jackson Laboratory) and NLRP3^−/− mice (originally obtained from Millennium Pharmaceuticals and kindly provided by Dr. Kate Fitzgerald, UMass Medical School). Cells were cultured in complete media supplemented with 10 ng/ml rM-CSF (eBioscience) and fed on days 4 and 7 with fresh media containing M-CSF. On day 8, nonadherent cells were washed away and the adherent macrophages were treated with 0.05% trypsin-EDTA, harvested, and washed once in complete media before use in experiments. For the experiments examining macrophage skewing shown in Fig. 1, BMMΦ were cultured with rGM-CSF (5 ng/ml; Miltenyi Biotec), IFN-γ (150 U/ml; eBioscience), M-CSF (10 ng/ml; eBioscience), and/or IL-4 (20 U/ml; eBioscience) (42). The cytokines were also in the media during the subsequent priming and stimulation steps. Bone marrow–derived DCs (BMDCs) were generated as described previously (43). In brief, bone marrow was extracted and cultured as described for M1-like macrophages, except on day 8 nonadherent cells were collected for use in experiments. Resident peritoneal cells were harvested by lavaging the peritoneal cavity of C57BL/6 mice or NLRP3^−/− mice with 10 ml PBS. Experimental protocols involving animals were approved by the University of Massachusetts Medical School Institutional Animal Care and Use Committee. Human PBMCs were isolated from the blood of adult, healthy donors under a protocol approved by the University of Massachusetts Medical School Institutional Review Board using Ficoll-Hypaque density centrifugation.

NO generation was quantified by measuring nitrite concentration in media removed from wells of cultured cells (44). Each sample (70 μl) was mixed with 70 μl water and 70 μl Griess Reagent in a 96-well plate and then incubated at 22°C for 30 min followed by measurement of absorbance at 540 nm. Standard curves were made using serial dilutions of sodium nitrite. After the removal of media, cells were lysed and arginase activity was measured (45). In brief, cells that had been cultured in 24-well plates were lysed by addition of 100 μl 0.1% Triton X-100, 1× Protease Inhibitor Cocktail (Roche), and incubation at 22°C for 30 min. To each well, 100 μl 25 mM Tris-HCl, pH 7.4, and 35 μl 10 mM MnCl₂ were added followed by incubation at 55°C for 10 min. Next, 12.5 μl of sample was mixed with 12.5 μl 0.5M l-arginine, pH 9.7, and incubated for 1 h at 37°C. The reaction was stopped with 200 μl H₂SO₄/H₃PO₄/H₂O (1:3:7) followed by addition of 12.5 μl 9% (w/v) α-isonitrosopropiophenone (dissolved in 100% ethanol) and heating at 99°C for 30 min. Standards were serial dilutions of urea, and absorbance readings were at 540 nm.

Chitosan was cleaved by pepsin to reduce the polymer length, purified, and converted to chitin as previously described (14). In brief, chitosan (250 mg) was dissolved in 25 ml 0.1 M sodium acetate, pH 4.5. Pepsin (P7000; Sigma) was added (100 U/ml) for 18 h at 37°C to partially digest the chitosan (46). This was followed by extraction with chloroform:isoamyl alcohol (24:1), then mixing the recovered aqueous layer with an equal volume of 12% potassium hydroxide and heating at 80°C for 90 min. Precipitated chitosan was collected by centrifugation and washed three times with water followed by PBS to neutralize. Half of the chitosan was converted to chitin by suspending in 20 ml 1.0 M sodium bicarbonate, followed by addition of 1 ml acetic anhydride (Acros) and incubation at 22°C for 20 min with periodic mixing. The acetylation reaction was repeated, then terminated with heating at 100°C for 10 min. Chitin was collected by centrifugation and washed three times with PBS. The degree of acetylation for chitin was ∼93% as previously reported (14). Chitin and chitosan suspensions were passed through a 100-μm filter (BD Falcon) to remove the largest particles, then treated in 0.1 M sodium hydroxide at 22°C for 30 min, followed by washing with PBS and storage at 4°C.

Murine cells were plated at 1 × 10⁵ cells/well in a 96-well plate. PBMCs were plated at 5 × 10⁶/well in 24-well plates and after 1 h, nonadherent cells were washed away. Mouse cells were primed with 100 ng/ml ultrapure LPS, whereas PBMCs were primed with 50 pg/ml ultrapure LPS for 3 h (control cells were left unprimed), followed by incubation with the stimuli for 6 h (18 h for PBMCs). Positive stimuli controls included silica (top size 15 μm, US Silica, MIN-U-SIL-15, used as described previously [29]), synthetic dsDNA:poly(dA:dT), ATP, and streptolysin O (SLO) or SLO + flagellin (FLA-ST; Invivogen). Supernatants were collected for cytokine measurement and assayed by ELISA for IL-1β (eBioscience) or IL-18 (MBL International). Western blotting was performed as described previously (14) to confirm that the IL-1β was processed. Only mature IL-1β was detected; no pro–IL-1β was identified (data not shown). For K⁺ efflux, ROS, and lysosomal inhibition assays, the inhibitors were added 1 h before secondary stimuli addition. K⁺ efflux was inhibited with glibenclamide, mitochondrial ROS was inhibited with Mito-TEMPO (Enzo Life Sciences), total ROS inhibited with diphenyleneiodonium chloride (DPI), lysosomal acidification was inhibited with bafilomycin A1 or chloroquine, and cathepsin B was selectively inhibited with CA-074-Me (Enzo Life Sciences). All inhibitors were assayed for cytotoxicity using the cytotoxicity detection kit (LDH) from Roche, and none of the inhibitors at the concentrations used induced significant cell death above background (data not shown).

BMMΦ were plated and primed as described earlier. Ten minutes after stimuli were added, cells given SLO or SLO + flagellin were washed three times with fresh media. Supernatants were collected for all stimuli after 2 h and analyzed by ELISA. For the K⁺ and Na⁺ buffer experiments, after the priming step the media were replaced with either K⁺ buffer (150 mM KCl, 5 mM NaH₂PO₄, 10 mM HEPES, 1 mM MgCl₂, 1 mM CaCl₂, 1% BSA) or Na⁺ buffer (150 mM NaCl, 5 mM KH₂PO₄, 10 mM HEPES, 1 mM MgCl₂, 1 mM CaCl₂, 1% BSA). SLO- and SLO + flagellin–treated cells were again washed three times after 10-min incubation and fresh buffer added; then supernatants for all stimuli were collected after 2 h and analyzed by ELISA for IL-1β.

BMMΦ were plated as described earlier and stimulated for 6 h with 0.1 mg/ml chitin or chitosan. Supernatants were collected and analyzed by Bio-Plex Pro Assays (Bio-Rad).

Data were analyzed and figures were prepared using GraphPad Prism. Significance was assessed by two-way ANOVA with Bonferroni post hoc test, one-way ANOVA with Dunnett post hoc test, or Kruskal–Wallis one-way ANOVA by ranks with Dunn post hoc tests, as indicated. The p values <0.05 after correction for multiple comparisons were considered significant.

We previously demonstrated that chitosan elicited a robust NLRP3 inflammasome-dependent IL-1β response in BMMΦ, whereas little IL-1β was elicited by chitin (14). To characterize the spectrum of cells that release IL-1β in response to chitosan and chitin, we studied a variety of cultured and primary cell types from mice and humans. First, to skew macrophages along a spectrum ranging from classically activated (M1) to alternatively activated (M2) phenotypes, mouse bone marrow cells were cultured in M-CSF, M-CSF + IL-4, GM-CSF, or GM-CSF + IFN-γ (42, 47). M1/M2 skewing was assessed by assaying for NO and arginase, respectively (Fig. 1A and 1B). Culture with M-CSF resulted in an intermediate phenotype with little NO or arginase activity observed. The addition of IL-4 to M-CSF strongly biased the cells toward an M2 phenotype. In contrast, GM-CSF induced an M1 type phenotype, which was more pronounced with the addition of IFN-γ. An IL-1β response was induced by chitosan in all four cell types (Fig. 1C). The response to chitosan was more pronounced in the GM-CSF–promoted M1 phenotype, which is consistent with the response to other stimuli (20). No significant differences were observed between the responses with M-CSF alone and M-CSF plus IL-4 after chitosan stimulation. Interestingly, though, the addition of IL-4 to M-CSF did result in a diminished IL-1β response to silica stimulation. We also observed a robust IL-1β response to chitosan in the primary peritoneal macrophages (Fig. 2A) and BMDCs (Fig. 2B). Finally, we tested whether human PBMCs respond to chitosan similarly as the mouse cells tested, and once again we saw a strong IL-1β response (Fig. 2C). Significant amounts of IL-1β above background levels were not released in response to chitin for any of the cell types tested (Figs. 1C and 2).

Our previous study demonstrated the inflammasome response induced by chitosan in BMMΦ was NLRP3 dependent (14). NLRP3^−/− mice were used to generate peritoneal macrophages and DCs, which were then challenged with chitosan to assess whether inflammasome activation in these cell types was NLRP3 dependent. For both peritoneal macrophages and DCs, IL-1β induced by chitosan was abolished in the NLRP3^−/− cells (Fig. 3).

To further analyze the spectrum of the inflammasome response to chitosan, we assayed release of the inflammasome-related cytokine, IL-18, in BMMΦ. As was observed when measuring IL-1β release, IL-18 release was stimulated by chitosan and silica, but not chitin (Fig. 4).

To understand how chitosan stimulates the NLRP3 inflammasome, we analyzed each of the three proposed mechanisms mentioned earlier in murine BMMΦ. To examine the contribution of cellular K⁺ efflux, we blocked K⁺ efflux using the K⁺ ion channel inhibitor glibenclamide (Fig. 5A). The release of IL-1β in response to both chitosan and silica was significantly inhibited in a dose-dependent manner by glibenclamide. However, the IPAF inflammasome activator flagellin (delivered to the cytosol by SLO) still induced IL-1β release even at the highest dose of inhibitor tested. To further confirm a role for K⁺ efflux, we destroyed the gradient required for K⁺ efflux by replacing the extracellular media with a buffer containing 150 mM K⁺, which is approximately equal to the intracellular K⁺ concentration. As a control, we used a buffer containing 150 mM Na⁺, which preserves the K⁺ gradient. Although all three stimuli induced IL-1β in the Na⁺ buffer, neither chitosan nor silica induced significant quantities of IL-1β in the high K⁺ buffer (Fig. 5B). Once again, flagellin delivered with SLO induced IL-1β even when the K⁺ gradient was collapsed. Thus, K⁺ efflux appears to be required for chitosan inflammasome activation.

To analyze the role of mitochondrial ROS, we used the mitochondrial ROS inhibitor, Mito-TEMPO. Using an inhibitor concentration of 100 μM, there was a significant reduction in IL-1β release induced by both chitosan and silica (Fig. 6A). To demonstrate specificity, we examined the effect of Mito-TEMPO on IL-1β release stimulated by the ROS-independent AIM2 inflammasome activator poly(dA:dT) (48). No significant inhibition of poly(dA:dT)-stimulated IL-1β was observed in the presence of Mito-TEMPO (Fig. 6A). A less specific ROS inhibitor, DPI, blocked IL-1β release in response to both NLRP3 and AIM2 stimuli (Fig. 6B). These data suggest that mitochondrial ROS is required for optimal NLRP3 inflammasome stimulation by chitosan.

Upon phagolysosomal fusion, acidification occurs that has been shown to be necessary for NLRP3 inflammasome activation through the lysosomal destabilization pathway (29). Lysosomal acidification was inhibited with either bafilomycin A1 (Fig. 7A) or chloroquine (Fig. 7B). IL-1β release from both chitosan and silica was significantly inhibited by bafilomycin A concentrations as low as 10 nM. Chloroquine concentrations as low as 1 μM significantly blocked IL-1β release induced by silica. A trend, albeit not statistically significant, toward a reduction in IL-1β release induced by chitosan in the presence of chloroquine was noted. To further explore the lysosomal destabilization pathway, we inhibited cathepsin B using CA-074-me (Fig. 7C). Upon lysosomal destabilization, cathepsin B is likely released into the cytosol where it activates the NLRP3 inflammasome (29). IL-1β release by the particulate stimuli chitosan and silica was significantly inhibited at CA-074-me concentrations of 10 μM. IL-1β release induced by the soluble inflammasome stimulator ATP was unaffected by these inhibitors, as expected for a phagocytosis-independent stimulus.

The earlier studies examined IL-1β release from primed BMMΦ. To examine the spectrum of cytokines and chemokines stimulated by chitosan and chitin, a multiplex assay was run on the supernatants from unprimed BMMΦ stimulated with these polysaccharides (Table I). None of the 22 cytokines and chemokines assayed was significantly induced by either chitosan or chitin. Trends of higher responses for chitosan compared with chitin were observed for some of the cytokines and chemokines, but these did not achieve statistical significance after corrections for multiple comparisons. The positive control, LPS, stimulated significant amounts of all the cytokines and chemokines tested except MCP-1 and RANTES.

We previously demonstrated that phagocytosable particles of chitosan, but not chitin, elicited a strong NLRP3 inflammasome response in BMMΦ (14). In this study, we examined the capacity of these particulate glycans to stimulate responses in other cell populations, characterized the mechanisms responsible for inflammasome activation by chitosan, and more fully elucidated the profile of cytokines and chemokines stimulated by chitosan and chitin. Chitosan induced significant IL-18 and IL-1β responses, activating the inflammasome of a broad range of cell populations and across species, whereas chitin failed to induce a significant response from the cell types tested. Although macrophages polarized toward M1, M2, and intermediate phenotypes were all stimulated by chitosan, the M1 response was greater, consistent with their more proinflammatory nature. The similar chitosan-stimulated IL-1β responses of intermediate phenotype and M2 macrophages suggest that the M1 phenotype is a primary driver of inflammasome activation. Indeed, the IL-1β response seen from the M2 macrophages may be partly explained by the plasticity macrophages and a switch toward a more M1 phenotype caused by the LPS priming step (49). The finding that chitosan stimulates IL-1β secretion in human cells has implications for the translational use of this polysaccharide. For example, the use of chitosan as a drug or vaccine delivery system could result in a potent, targeted inflammatory response (38).

In analyzing the mechanistic basis for NLRP3 inflammasome activation, we found evidence that K⁺ efflux, mitochondrial ROS, and lysosomal destabilization each contribute to chitosan activation of the NLRP3 inflammasome. K⁺ efflux from the cytosol has been shown to be required for NLRP3 inflammasome activation by many different stimuli. Some of the most potent NLRP3 inflammasome inducers are, in fact, K⁺ channels inducers, including nigericin, gramicidin, maitotoxin, and α-toxin (50). Originally, these were thought to act by creating pores that allowed the NLRP3 stimuli access to the cytoplasm, whereby they could potentially interact directly with NRLP3. However, direct interaction with NLRP3 is unlikely given the variety of stimuli; therefore, NLRP3 activation may be through sensing the K⁺ efflux by an unknown mechanism. Inhibition of chitosan-induced IL-1β release through blocking K⁺ efflux is consistent with previous published reports for other NLRP3 inflammasome stimuli (16, 50), showing that chitosan also requires K⁺ efflux to activate the NLRP3 inflammasome.

The second proposed model for NLRP3 inflammasome activation involves ROS-induced activation of NRLP3. Some debate exists on what type of ROS is important for NLRP3 activation because ROS scavengers and inhibitors strongly impede NLRP3 activation (51, 52). In addition, it has been shown that knockdown of the p22phox subunit of the NOX 1–4 subfamily of NADPH oxidases also impairs NLRP3 activation (51). However, multiple studies have shown that cells from patients with chronic granulomatous disease, a disease with defects in NADPH oxidase, exhibit no NLRP3 deficiencies (53–55). Recent work has strongly suggested that the important ROS for NLRP3 inflammasome activation is mitochondrial in origin and that mitochondria are central to NLRP3 activation (26, 27). Oxidized mitochondrial DNA has also been proposed to act as an agonist of NLRP3 (56). In this article, we demonstrated that blocking mitochondrial ROS inhibited chitosan-induced IL-1β release. The targeted mitochondrial ROS inhibitor had no significant effect on the AIM2 inflammasome activator poly(dA:dT), whereas the less specific ROS inhibitor, DPI, blocked IL-1β for both NLRP3 and AIM2 stimuli. ROS may not only be involved in the NLRP3 activation, but may also be important for the priming step, the upregulation of pro–IL-1β and NLRP3 (48). This could explain why DPI has more far-reaching effects than the more targeted Mito-TEMPO.

Acidification-dependent lysosomal destabilization resulting in release of cathepsin B and subsequent activation of the NLRP3 inflammasome has been demonstrated after phagocytosis of particulate stimuli including silica and alum (29). We hypothesized that the polycationic properties of chitosan would enable its escape from lysosomes by sponging protons delivered by the vacuolar-ATPase. This, in turn, would lead to the retention of Cl⁻ ions and water molecules resulting in lysosome swelling, leakage, and eventual rupture (57, 58). Indeed, we found that inhibition of lysosomal acidification (which is required for the proton sponge effect) prevented chitosan from activating the inflammasome. The neutral charge of chitin due to acetylation of the polymer is a possible reason for its inability to activate the inflammasome. We were also able to block chitosan-induced NLRP3 inflammasome activation by blocking the downstream proposed activator of the lysosomal destabilization mechanism, cathepsin B, with the cathepsin B–specific inhibitor Ca-074-me. Inhibition was incomplete, which may be because of stimulation of the NLRP3 inflammasome by other lysosomal contents, such as cathepsin L or cathepsin D (59–61).

The preparations of chitosan and chitin that were used in our studies were extensively purified to remove potential contaminants including proteins, nucleic acids, and LPSs. The purification steps included solubilization, chloroform:isoamyl alcohol extraction, reprecipitation, hot-alkali treatment, and extensive washing. Using the resulting ultrapure preparations, chitin and chitosan did not stimulate unprimed macrophages to release statistically significant quantities of any of the 22 cytokines and chemokines analyzed by multiplex assay. However, because of our use of stringent criteria to correct for the large number of comparisons, the possibility of a Type II error causing us to miss significant associations cannot be ruled out. Indeed, some of the cytokines and chemokines trended higher, with chitosan being a more potent stimulus compared with chitin. It should also be noted that the concentrations of cytokines and chemokines stimulated by chitosan and chitin were relatively low compared with LPS.

Others have reported that chitin can elicit IL-10, TNF-α, IL-17A, IL-12, or IL-18 (35, 39, 62). The explanation for discrepancies between these published data and ours is speculative but may be because of differences in chitin source, contaminants, tertiary structures, and/or particle size (38). Less work has been done on the immunological activity of chitosan, although our inflammasome and multiplex assays suggest that chitosan is the more immunostimulatory of the two polymers. Defining the conditions under which chitin and chitosan trigger or fail to trigger immune responses has translational relevance given frequent natural exposure and the increasing use of these polymers in biomedical applications.

We thank Dr. Kate Fitzgerald for helpful discussions and providing the NLRP3^−/− mice, Dr. Vijay Rathinam for help with the Western blots, and Dr. Sumanth Gandra for statistical assistance.

The authors have no financial conflicts of interest.