Pro-inflammatory and adjuvant functions of Toll-like receptor 4 antagonists

Abstract

Methods and compositions are provided for specifically activating the inflammatory response of dendritic cells (DCs). [Solution] 1-palmitoyl-2-arachidonyl-sn-glycero-3-phosphorylcholine (PAPC) and its oxidized variant (oxPAPC) have been identified to promote DC-mediated immunity and are provided as adjuvants for immunostimulatory compositions, including vaccines.

Description

[Technical Field]

[0001] CROSS-REFERENCE TO RELATED APPLICATIONS This is an international patent application claiming the benefit of priority under 35 U.S.C. §119(e) to U.S. Provisional Patent Application No. 62 / 102,245, entitled "Pro-Inflammatory and Adjuvant Functions of Toll-Like Receptor 4 Antagonists," filed January 12, 2015, the contents of which are incorporated herein by reference in their entirety.

[0002] STATEMENT OF RIGHTS TO INVENTIONS MADE IN FEDERALLY SPONSORED RESEARCH This work was supported by Grant No. AI103082-01A1 awarded by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health. The federal government has certain rights in this invention.

[0003] The present invention relates generally to the fields of adjuvants, immune activation, and vaccines. [Background technology]

[0004] The ability to distinguish between self and non-self molecules is a fundamental feature of all living organisms, yet our understanding of this distinction remains incomplete. In mammals, pattern recognition receptors (PRRs) of the innate immune system are generally believed to function to distinguish between self and non-self molecules. This concept, first proposed by Charles Janeway Jr., has been widely validated through studies of various PRR families, including Toll-like receptors (TLRs), RIG-I-like receptors (RLRs), NOD-like receptors (NLRs), and C-type lectin receptors (CLRs) (Iwasaki, A., and Medzhitov, R. (2015) Nat Immunol 16, 343-353). PRRs either directly or indirectly detect molecules common to a wide variety of microorganisms. These molecules are traditionally called pathogen-associated molecular patterns (PAMPs) and include factors such as bacterial lipopolysaccharide (LPS), bacterial flagellin, or viral double-stranded RNA (Janeway, CA, Jr. (1989) Spring Harb Symp Quant Biol 54 Pt 1, 1-13). Detection of microbial products activates PRR-dependent cellular responses that are either proinflammatory or immunoregulatory. The best example of the latter is the promotion of adaptive immunity through the activation of antigen-specific T cells (Iwasaki, A., and Medzhitov, R. (2015) Nat Immunol 16, 343-353). While PRR-mediated proinflammatory responses can be considered to occur in multiple cell types, the activity to promote T cell activation often occurs specifically in dendritic cells (DCs).DC-specific activities induced by PRRs include the acidification of endosomes and phagosomes (Delamarre, L. et al., (2005) Science 307, 1630-1634, and Trombetta, E.S. et al., (2003) Science 299, 1400-1403), delivery of major histocompatibility complex (MHC) molecules to microbe-containing phagosomes (Nair-Gupta, P. et al. (2014) Cell 158, 506-521), loading of MHC with microbial peptides, and delivery of MHC molecules to the cell surface (Blander, J.M., and Medzhitov, R. (2006). Nature 440, 808-812; Inaba, K. et al., (2000) J Exp Med 191, 927-936; Pierre, P. et al. al., (1997) Nature 388, 787-792; Turley, SJ et al., (2000) Science 288, 522-527). All of these activities promote efficient antigen presentation to T cells and the initiation of adaptive immunity.

[0005] Adjuvants are substances that accelerate and / or enhance antigen-specific immune responses. The purpose of an adjuvant is to make antigens visible to the immune system's equivalent of eyes (macrophages / dendritic cells). Antigen recognition by antigen-presenting cells (APCs), such as macrophages and dendritic cells, essentially initiates a cascade of key events that lead to local inflammation, which in turn recruits APCs and ultimately initiates germ cell- and / or antibody-mediated immune responses. Currently, most human vaccines contain aluminum salts as adjuvants, and pharmaceutical companies are developing oil-based adjuvants for incorporation into vaccines. For the development of improved immunostimulatory compositions (e.g., vaccines), the identification and inclusion of adjuvants that selectively activate dendritic cells (DCs) while minimally activating macrophages would be beneficial in reducing adverse effects, such as discomfort and inflammation, associated with the administration of such compositions. Adjuvants that act as agonists for TLR-2, TLR-5, TLR7 / 8, and TLR-9 are currently under investigation, and one TLR-4 agonist, monophosphoryl lipid A, is FDA approved. [Prior art documents] [Non-patent literature]

[0006] [Non-Patent Document 1] Iwasaki, A., and Medzhitov, R. (2015) Nat Immunol 16, 343-353 [Non-patent document 2] Janeway, CA, Jr. (1989) Spring Harb Symp Quant Biol 54 Pt 1,1-13 [Non-patent document 3] Delamarre, L. et al., (2005) Science 307, 1630-1634 [Non-patent document 4] Trombetta,ESet al.,(2003)Science 299,1400-1403 [Non-patent document 5] Nair-Gupta,P.et al.(2014)Cell 158,506-521 [Non-patent document 6] Blander, J.M., and Medzhitov, R. (2006).Nature 440,808-812 [Non-Patent Document 7] Inaba,K.et al.,(2000)J Exp Med 191,927-936 [Non-patent document 8] Pierre, P. et al., (1997) Nature 388, 787-792 [Non-Patent Document 9] Turley,SJet al.,(2000)Science 288,522-527 Summary of the Invention

[0007] The present invention is based, at least in part, on the discovery that endogenous oxidized phospholipids, which are Toll-like receptor (TLR) antagonists found at sites of tissue injury, created a hyperinflammatory state in dendritic cells. In particular, we demonstrated that oxPAPC induced several responses in DCs in a context-dependent manner that promoted their ability to activate antigen-specific T cells. These findings indicated that oxPAPC (and related phospholipids capable of activating non-canonical inflammasomes, such as Rhodo LPS, which has also been found to activate non-canonical inflammasomes) could function as enhanced adjuvants for use in prophylactic and therapeutic immunostimulatory compositions.

[0008] We found that, in the presence of various TLR ligands, oxPAPC promoted DC survival and induced the release of the T cell-activating cytokine interleukin-1 beta (IL-1β). Mechanistically, oxPAPC was characterized by binding to the LPS receptor CD14 on the DC surface, which delivered oxPAPC into endosomes and subsequently provided access to the cytosolic protein caspase-11. Binding of oxPAPC to caspase-11 triggered inflammasome-mediated IL-1β release. These oxPAPC-induced responses did not occur in macrophages, indicating that this lipid acts as a unique mechanism for promoting the immunomodulatory activity of DCs, rather than a general (macrophage-mediated) inflammatory response. Consequently, we found that oxPAPC synergized with microbial products to induce more robust antigen-specific T cell activation than could be induced by PAMPs alone. We identified that these molecules (termed vita-DAMPs) functioned together with PAMPs to hyperactivate DCs and elicit maximal adaptive immune responses.

[0009] In one embodiment, the invention provides a composition for eliciting an immune response to an immunogen, comprising an immunogen and a non-canonical inflammasome-activating lipid.

[0010] In one embodiment, the non-canonical inflammasome-activating lipid is oxPAPC. In another embodiment, the non-canonical inflammasome-activating lipid is PAPC. Optionally, the non-canonical inflammasome-activating lipid is one or more species of oxPAPC. In a related embodiment, the non-canonical inflammasome-activating lipid is one or more of HOdiA-PC, KOdiA-PC, HOOA-PC, and KOOA-PC. In another embodiment, the non-canonical inflammasome-activating lipid is Rhodo LPS.

[0011] In further embodiments, the non-canonical inflammasome-activating lipid enhances the immune response of an immunogen when the composition is administered to a subject, compared to a composition lacking the non-canonical inflammasome-activating lipid.

[0012] In one embodiment, the immunogen and lipid are present in concentrations sufficient to induce dendritic cell (DC) activation when the composition is administered to a subject.

[0013] Optionally, the composition does not induce a macrophage inflammatory response when administered to a subject.

[0014] In one embodiment, the immunogen is a human papillomavirus antigen, a herpesvirus antigen such as a herpes simplex antigen or a varicella zoster antigen, a retrovirus antigen such as a human immunodeficiency virus type 1 antigen or a human immunodeficiency virus type 2 antigen, a hepatitis virus antigen, an influenza virus antigen, a rhinovirus antigen, a respiratory syncytial virus antigen, a cytomegalovirus antigen, an adenovirus antigen, a Mycoplasma pneumoniae antigen, or a genotype antigen. pneumoniae antigens, antigens from bacteria of the genera Salmonella, Staphylococcus, Streptococcus, Enterococcus, Clostridium, Escherichia, Klebsiella, Vibrio, and Mycobacterium, amoeba antigens, malaria parasite antigens, and / or Trypanosoma cruzi antigens.

[0015] Optionally, the composition is lyophilized.

[0016] In another embodiment, the composition consists essentially of an immunogen in combination with a non-canonical inflammasome-activating lipid.

[0017] Another aspect of the present invention provides a pharmaceutical composition comprising the immunogenic adjuvant composition of the present invention and a pharmaceutically acceptable carrier.

[0018] In one embodiment, the carrier is an aqueous carrier. In another embodiment, the carrier is a solid carrier.

[0019] A further aspect of the invention provides a method for inducing an inflammatory response in dendritic cells in a subject, the method comprising administering a composition according to claim 1 to the subject.

[0020] A further aspect of the present invention provides a method for enhancing a subject's protective immune response to an immunogen by administering to the subject an immunogen and a non-canonical inflammasome-activating lipid in amounts effective to enhance the subject's protective immune response, wherein the non-canonical inflammasome-activating lipid is administered in an adjuvant-effective amount.

[0021] In one embodiment, the immunogen and the non-canonical inflammasome-activating lipid are co-administered to the subject.

[0022] Another aspect of the invention provides a method for inducing an immune response in a subject, the method comprising co-administering to the subject an immunogen and a non-canonical inflammasome-activating lipid in amounts effective to generate an immune response in the subject.

[0023] In one embodiment, the subject is a human.

[0024] In another embodiment, the immunogen and non-canonical inflammasome-activating lipid are co-administered with a common pharmaceutical carrier.

[0025] Optionally, the immunogen and non-canonical inflammasome-activating lipid are administered parenterally.

[0026] In one embodiment, the immune response is a prophylactic immune response.

[0027] In another embodiment, the immune response is a therapeutic immune response.

[0028] In a further embodiment, the immune response comprises a humoral immune response.

[0029] Other aspects of the invention are described in or are obvious from the following disclosure and are within the scope of the invention. [Brief explanation of the drawings]

[0030] [Figure 1] FIG. 1 shows that PAPC (eg, oxPAPC) was identified to act as a TLR4-antagonist but not as a TLR4-agonist. [Figure 2] FIG. 2 shows that various doses of PAPC modulated TLR4 signaling. [Figure 3] FIG. 3 shows that various doses of PAPC modulated CD14 levels on the plasma membrane. [Figure 4] FIG. 4 shows that various doses of PAPC modulated LPS-dependent TLR4 internalization. [Figure 5] FIG. 5 shows that various doses of PAPC modulated LPS-dependent TLR4 dimerization. [Figure 6] FIG. 6 shows that PAPC activated the inflammasome in DCs. [Figure 7] FIG. 7 shows that KOdiA-PC activated inflammasomes. [Figure 8] FIG. 8 shows that CD14 regulated inflammasome activation in response to PAPC. [Figure 9] FIG. 9 shows that inflammasome activation in response to PAPC was CD14-specific, but PAPC was also able to induce CD36 internalization. [Figure 10] FIG. 10 shows that CD14 regulated PAPC-mediated inflammasome activation independently of type I IFN. [Figure 11] FIG. 11 shows that the regulation of caspase-1 and caspase-11 expression was similar in wtDCs and Cd14 − / − DCs. [Figure 12]FIG. 12 shows that PAPC induced inflammasome activation in a cell type-specific manner. [Figure 13] FIG. 13 shows that other PAMPs primed PAPC-induced inflammasome activation. [Figure 14] FIG. 14 shows further results showing that other PAMPs primed PAPC-induced inflammasome activation. [Figure 15] FIG. 15 shows that not all modified PCs induced inflammasome activation. [Figure 16] FIG. 16 shows that Nlrp3 was required for the induction of inflammasome activation. [Figure 17] FIG. 17 shows that Asc was required for the induction of inflammasome activation. [Figure 18] FIG. 18 shows that Casp1 / Casp11 was required for the induction of inflammasome activation. [Figure 19] FIG. 19 shows that oxPAPC-induced, but not ATP-induced, inflammasome activation was caspase-11 dependent. [Figure 20] FIG. 20 shows that caspase-11 was required for PAPC-induced inflammasome activation after pre-stimulation with both LPS and Pam3. [Figure 21] FIG. 21 shows that biotinylated PAPC potently induced CD14 internalization. [Figure 22] FIG. 22 shows that biotinylated PAPC did not induce TLR4 internalization. [Figure 23] FIG. 23 shows that biotinylated PAPC did not induce IL-1b secretion. [Figure 24]FIG. 24 shows an in vitro binding assay of biotinylated LPS, OxPac, and Pac to caspase-11 and MD-2, which was thought to identify complex formation in a caspase-11-dependent manner. [Figure 25] Figure 25 shows that PAPC acted as a dose-dependent competitor in Bio-LPS pull-down. Biotin-LPS was used at 5 μg per pull-down assay. PAPC was used at 5, 50, and 500 μg to compete with LPS binding to MD-2 and caspase-11, respectively. Competition began to be effective at a ratio of 1:100 (LPS:PAPC). [Figure 26] FIG. 26 shows that oxPAPC and LPS likely bound to the same domain of CD14. [Figure 27] FIG. 27 shows that LPS treatment affected DC survival. [Figure 28] FIG. 28 shows that PAPC treatment of pre-stimulated DCs promoted DC survival. [Figure 29] FIG. 29 shows that the observed PAPC-dependent pro-survival effect was not CD14-dependent. [Figure 30] FIG. 30 shows that only P2C and P3C supported DC survival. [Figure 31] FIG. 31 shows that DCs primed with P2C and P3C did not increase their survival in response to PAPC treatment. [Figure 32] FIG. 32 shows that inflammasomes were efficiently activated by priming and co-administration of PAPC, but not by ATP. [Figure 33] FIG. 33 shows that priming and co-administration of PAPC did not alter NF-κB activation of wtDCs. [Figure 34]FIG. 34 shows that in the absence of CD14, oxPAPC acted as an antagonist of TLR4 signaling. [Figure 35] FIG. 35 shows that co-administration of LPS and oxPAPC affected TLR4 internalization. [Figure 36] FIG. 36 shows that co-administration of LPS and oxPAPC affected CD14 internalization. [Figure 37] FIG. 37 shows that co-administration of LPS and oxPAPC partially affected TLR4 dimerization. [Figure 38] FIG. 38 shows that Rhodo LPS was a potent inducer of inflammasome activation. [Figure 39] FIG. 39 shows that LPS-induced inflammasome activation was Nlrp3 dependent. [Figure 40] FIG. 40 shows that LPS-induced inflammasome activation was Asc-dependent. [Figure 41] FIG. 41 shows that LPS-induced inflammasome activation was Casp1 / 11 dependent. [Figure 42] FIG. 42 shows that Rhodo LPS-induced inflammasome activation was CD14-independent. [Figure 43]Figures 43A-43D are images showing that oxPAPC did not bind to TLR4 or induce TLR4F signaling. Figure 43A shows a line graph depicting the extent of TLR4 dimerization in iMΦ treated with LPS (1 μg / ml) or oxPAPC (50 μM) for the indicated time periods. TLR4 dimerization was measured by flow cytometry. The line graphs represent the mean and standard deviation of two independent experiments. Figure 43B shows a line graph depicting the levels of IL-1β, IL-6, IFNβ, and Viperin in iMΦ treated with LPS (1 μg / ml) or oxPAPC (50 μM). Gene expression relative to GAPDH was analyzed by qPCR at the indicated times. Untreated cells were used as a negative control in all experiments. The line graphs represent the mean, and error bars represent the standard deviation of triplicate determinations in one representative experiment out of three. Figure 43C shows blots demonstrating myddosome formation in iMΦs at the indicated time points after treatment with LPS (1 μg / ml) or oxPAPC (50 μM) by co-immunoprecipitation (IP) of IRAK4 with MyD88 followed by Western analysis of the indicated proteins. Figure 43D shows blots depicting harvested whole cell lysates (WCLs) and DCs treated with LPS (1 μg / ml) or oxPAPC (50 μM) and monitored for STAT-1 phosphorylation and viperin expression. [Figure 44]Figures 44A-44F are images showing that oxPAPC acted as both a CD14 agonist and a TLR4 antagonist. Figure 44A shows line graphs depicting the surface levels of CD14 and TLR4 in iMΦ lines treated with LPS (1 μg / ml) or oxPAPC (50 μM) for the indicated times. Surface levels of CD14 and TLR4 were measured by flow cytometry. The line graphs represent the mean and standard deviation of two independent experiments. Figure 44B shows line graphs depicting the results of primary DCs and MΦs treated with oxPAPC (50 μM) for the indicated times. Surface levels of CD14 and TLR4 and TLR4 dimerization were measured by flow cytometry. The line graphs represent the mean and standard deviation of two independent experiments. Figure 44C shows line graphs depicting the results of iMΦ treated with LPS alone (1 μg / ml), oxPAPC alone (at the indicated concentrations), or 30 min pre-treated with oxPAPC followed by LPS treatment. TLR4 surface levels and TLR4 dimerization were measured by flow cytometry. Line graphs represent the mean and standard deviation of biological replicates from one representative experiment out of three. Figure 44D shows images depicting the results of iMΦ treated with LPS alone (at the indicated concentrations), oxPAPC alone (120 μM), or 30 min pre-treated with oxPAPC followed by LPS treatment. Figure 44D (left panel) shows TNFα secretion measured by ELISA 18 h after LPS stimulation. Line graphs represent the mean, and error bars represent the standard deviation of triplicate determinations from one representative experiment out of three. Figure 44D (right panel) shows STAT-1 phosphorylation measured by Western analysis 4 hours after LPS treatment. Figure 44E shows a blot showing the oxPAPC-binding ability of CD14 mutants determined by biotinylated oxPAPC pull-down assay. Lysates of 293T cells expressing the indicated CD14 mutants were incubated with biotinylated oxPAPC (10 μg). CD14-oxPAPC complexes were then captured using neutravidin beads. The amount of CD14 retained by oxPAPC was determined by Western analysis.The CD14 mutant in which 26DEES29 was mutated to 26AAAA29 was designated CD14 1R. The CD14 mutant in which 26DEES29 and 37PKPD40 were mutated to 26AAAA29 and 37AAAA40 was designated CD14 2R. The CD14 mutant in which 26DEES29, 37PKPD40, 52DVE54, and 74DLGQ77 were mutated to 26AAAA29, 37AAAA40, 52AAA54, and 74AAAA77 was designated CD14 4R. Figure 44F shows a graph demonstrating that the 4R CD14 mutant was not internalized in response to oxPAPC or LPS treatment. The indicated iMΦ lines were treated with LPS (1 μg / ml) or oxPAPC (50 μM) for the indicated times. Surface levels of CD14 were measured by flow cytometry. Line graphs represent the mean and error bars represent the standard deviation of biological replicates in one representative experiment out of three. [Figure 45]Figures 45A-G show images demonstrating that oxPAPC induced NLRP3 inflammasome activation in DCs. Figure 45A shows bar graphs depicting IL-1β secretion in DCs treated with LPS alone (1 μg / ml), three doses of oxPAPC (10, 50, and 120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC treatment. Commercially available oxPAPC and oxPAPC enriched in PEIPC were used in this experiment. Eighteen hours after LPS administration, secreted (left panel) and cell-bound (right panel) IL-1β were measured by ELISA. The mean and standard deviation of biological replicates from one representative experiment out of two are shown. Figures 45B–45D show bar graphs depicting the results of WT DCs or caspase-1 KO and caspase-1 / -11 dKO DCs (Figure 45B), ASC KO DCs (Figure 45C), and NLRP3 KO DCs (Figure 45D) treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC. Eighteen hours after LPS administration, IL-1β secretion (left panel) and TNFα secretion (right panel) were measured by ELISA. Means and standard deviations of two independent experiments are shown. Figure 45E shows bar graphs depicting the results of IL-1β secretion from MΦs treated with LPS alone (1 μg / ml), three doses of oxPAPC (10, 50, and 120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC. In this experiment, commercially available oxPAPC and oxPAPC enriched in PEIPC were used. Eighteen hours after LPS administration, secreted (left panel) and cell-bound (right panel) IL-1β were measured by ELISA. Mean and standard deviation of biological replicates from one representative experiment out of two are shown. Figure 45F shows a bar graph depicting IL-1β secretion from MΦs treated with Pam3CSK (P3C) alone (1 μg / ml), oxPAPC alone (120 μM), ATP alone (5 mM), or pre-stimulated with Pam3CSK for 3 hours followed by treatment with oxPAPC, ATP, DOTAP, LPS (5 μg), or oxPAPC encapsulated in DOTAP.IL-1β was measured by ELISA 18 hours after P3C administration. Means and standard deviations of three replicates from one of two experiments are shown. Figure 45G shows bar graphs demonstrating that LPS-prestimulated DCs and MΦs exhibited unique differences in their response to NLRP3 activation after ATP treatment. DCs (left panel) or MΦs (right panel) were prestimulated with LPS (1 μg / ml) for 3 hours and treated with ATP (3 mM). At the indicated time points, IL-1β was measured by ELISA, and cell death was measured by PI permeability assay. Means and standard deviations of four replicates from one of three experiments are shown. [Figure 46]Figures 46A-G show oxPAPC non-canonical inflammasome activation. Figure 46A shows bar graphs depicting IL-1β secretion from WT DCs and caspase-11 KO DCs treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC treatment. 18 hours after LPS administration, IL-1β secretion was measured by ELISA. The mean and standard deviation of two independent experiments are shown. Figure 46B shows bar graphs depicting TNFα secretion from WT DCs and caspase-11 KO DCs treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC treatment. 18 hours after LPS administration, TNFα secretion was measured by ELISA. The mean and standard deviation of two independent experiments are shown. Figure 46C shows images demonstrating that DCs primed with LPS, but not ATP, formed specks in a caspase-11-dependent manner in response to oxPAPC. DCs were either left untreated or primed with LPS (1 μg / ml) for 3 hours and then stimulated with ATP (1 mM) or oxPAPC (120 μM). Speck-containing ASCs (green) and caspase-1 (Casp1, red) were analyzed 18 hours after LPS stimulation. Nuclei are shown in blue. Each panel represents four independent experiments. Figure 46D shows blots demonstrating the in vitro binding of endogenous caspase-11 to oxPAPC. S100 fractions (0.5 mg) from untreated (nt) or P3C-primed (P3C) MΦs were incubated with biotinylated LPS (Bio-LPS) or biotinylated oxPAPC (Bio-oxPAPC). Endogenous proteins bound to biotinylated lipids were captured with streptavidin and resolved by Western analysis. A representative blot from three independent experiments is shown. Figure 46E shows a graph of SPR analysis of protein interactions with the indicated lipids. Figure 46F shows a graph showing gel filtration analysis of the size of caspase-11 complexes before and after contact with oxPAPC.Complex size was monitored by A280 or Western analysis, as indicated. Figure 46G shows bar graphs depicting secretion and viability results for bone marrow cells infected with the pMSCV2.2-IRES-GFP vector (empty), the pMSCV2.2-IRES-GFP vector encoding WT caspase-11 (WT caspase-11), or the same vector containing the catalytic mutant caspase-11 (C254A). After 7 days of differentiation in GM-CSF-containing medium, DCs were either pre-stimulated with LPS (1 μg / ml) for 3 hours or not, and then stimulated with oxPAPC (120 μM) or transfected with LPS-containing FuGENE (LPS, 5 μg). Eighteen hours after LPS pre-stimulation, supernatants were collected, and IL-1β and TNFα secretion were measured by ELISA. Cell viability was assessed by measuring LDH release. [Figure 47]Figures 47A-47E are bar graphs showing that CD14 promoted caspase-11-mediated IL-1β release from DCs. Figure 47A shows bar graphs depicting the secretion results of WT and CD14KO DCs treated with LPS alone (1 μg / ml), Pam3CSK (P3C) alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with LPS or Pam3CSK for 3 hours followed by oxPAPC treatment. IL-1β and TNFα secretion were measured by ELISA 18 hours after LPS or P3C administration. Means and standard deviations of two independent experiments are shown. Figure 47B shows bar graphs depicting IL-1β and TNFα secretion from spleen-derived WT DCs, CD14 KO DCs, and caspase-11 KO DCs treated with LPS alone (1 μg / ml), ATP alone (1.5 mM), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC or ATP treatment. IL-1β and TNFα secretion were measured by ELISA 18 hours after LPS administration. Means and standard deviations of two replicates from one representative experiment out of three are shown. Figure 47C shows bar graphs depicting gene expression from WT DCs and CD14 KO DCs treated with LPS or Pam3CSK. Gene expression in response to TBP was analyzed by qPCR at the indicated times. Results are shown as gene expression relative to untreated cells. Line graphs represent the mean of triplicate measurements from one representative experiment out of three. Figure 47D shows bar graphs depicting IL-1β and TNFα secretion results from WT and CD14 KO DCs treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by treatment with oxPAPC in the presence or absence of rIFNβ (100 U / ml). 18 hours after LPS administration, IL-1β and TNFα secretion were measured by ELISA. Means and standard deviations of two independent experiments are shown.Figure 47E shows a bar graph depicting IL-1β secretion from WT and CD14KO DCs treated with LPS alone (1 μg / ml), Pam3CSK (P3C) alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with Pam3CSK for 3 hours followed by oxPAPC treatment. As indicated, LPS or oxPAPC was complexed with DOTAP before addition to the cell culture. As indicated, cells were treated with the pan-caspase inhibitor zVAD for 30 minutes before addition of the DOTAP / oxPAPC complex to the culture. IL-1β secretion was measured by ELISA 18 hours after stimulation. The mean and standard deviation of two independent experiments are shown. [Figure 48]Figures 48A-G show images demonstrating that oxPPAC acted like a natural adjuvant, preventing DC death and enhancing adaptive immune responses. Figure 48A shows bar graphs depicting the viability results of DCs treated with LPS alone (1 μg / ml), ATP alone (1 mM), oxPAPC alone (120 μM), FuGENE-complexed LPS (5 μg) (Fugene(LPS)), or pre-stimulated with LPS (1 μg / ml) for 3 hours followed by the indicated stimuli. Cell death was measured by LDH release 4 and 18 hours after LPS pre-stimulation. Figure 48B shows a bar graph depicting IL-1β secretion from DCs treated with LPS alone (1 μg / ml), ATP alone (1 mM), oxPAPC alone (120 μM), or FuGENE-complexed LPS (5 μg) (Fugene(LPS)), or pre-stimulated with LPS (1 μg / ml) for 3 hours followed by the indicated stimuli. IL-1β secretion was measured by ELISA 4 and 18 hours after LPS pre-stimulation. Figures 48C-D show images depicting staining of DCs pre-treated with LPS (1 μg / ml) for 3 hours followed by activation with ATP (1 mM) or oxPAPC (120 μM). After 18 hours, cells were stained with ASC (green), nuclei (blue), Zombie dye (red) (Figure 48C), or active mitochondria (red) (Figure 48D). Panels represent three independent experiments. Figures 48E-48F show bar graphs depicting the viability results of DCs treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), ATP alone (1 mM), or pre-stimulated with LPS for 3 hours followed by treatment with oxPAPC or ATP. At the indicated time points, cell viability was measured by 7-AAD staining. The mean and standard deviation of two independent experiments are shown. Figure 48G shows a bar graph demonstrating that oxPAPC enhanced memory T cell responses in vivo.CD4+ T cells were isolated from draining lymph nodes of WT, caspase-1 / -11 dKO, or caspase-11 KO mice 40 days after immunization with OVA + LPS in IFA(LPS), OVA + LPS + oxPAPC in IFA(LPS) or OVA + oxPAPC in IFA(oxPAPC). CD4+ T cells were either restimulated with OVA in the presence of DCs or not. IFNγ secretion (left panel) and IL-17 secretion (right panel) were measured by ELISA 5 days later. Bar graphs represent the mean and standard error of two experiments with five animals per group. [Figure 49] Figures 49A-49B are blots showing that oxPAPC did not induce myofibroblast growth or type I IFN signaling (related to Figures 43A-43D). Figure 49A shows a bar graph demonstrating myofibroblast growth in iMΦs assessed at the indicated time points after treatment with LPS (1 μg / ml) or different doses of oxPAPC (10, 50, 120 μM) by coimmunoprecipitation of IRAK4 with MyD88 followed by Western analysis for the indicated proteins. Figure 49B shows blots showing whole-cell lysates (WCLs) collected from DCs and treated with LPS (1 μg / ml) or different doses of oxPAPC (10, 50, 120 μM) followed by Western analysis for the indicated proteins. [Figure 50]Figures 50A-50D are graphs showing that oxPAPC was an agonist of CD14 but not TLR4 (see also Figures 44A-44F). Figure 50A shows a bar graph depicting surface CD14 results for DCs treated with the indicated concentrations of oxPAPC. Surface levels of CD14 were measured by flow cytometry. Line graphs represent the mean and standard deviation of two independent experiments. Figures 50B-50D show bar graphs depicting surface CD14, surface TLR4, and TLR4 dimerization results for DCs treated with LPS (1 μg / ml) or oxPAPC (120 μM) in the presence or absence of cycloheximide (100 μg / ml) for the indicated times. Surface levels of CD14 (Figure 50B), TLR4 (Figure 50C), and TLR4 dimerization (Figure 50D) were measured by flow cytometry. Line graphs represent the mean and standard deviation of two independent experiments. [Figure 51]Figures 51A-51I are images showing that oxPAPC induced IL-1β release in a cell type-specific manner (see also Figures 45A-45G). Figure 51A shows bar graphs showing that DCs and iMΦs released IL-1β in response to ATP. Freshly induced DCs and iMΦs were treated with LPS alone (1 μg / ml), ATP alone (ATP low: 0.5 mM; ATP high: 5 mM), or pre-stimulated with LPS for 3 hours followed by ATP treatment. Eighteen hours after LPS administration, supernatants were collected, and the levels of secreted IL-1β were measured by ELISA. The mean and standard deviation of two independent experiments are shown. Figures 51B-51C show bar graphs depicting secreted and cell-bound IL-1β levels in DCs pre-stimulated with or without different doses of LPS (100 and 1000 ng / ml), and then activated with oxPAPC (120 μM) or ATP (0.5 mM) 3 hours later. Eighteen hours after LPS administration, supernatants were collected, and the levels of secreted IL-1β (Figure 51B) and cell-bound IL-1β (Figure 51C) were measured by ELISA. The mean and standard deviation of biological replicates from one representative experiment out of three are shown. Figure 51D shows bar graphs depicting TNFα results in DCs or MΦs treated with LPS alone (1 μg / ml), ATP alone (0.5 mM), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by treatment with ATP or oxPAPC. Eighteen hours after LPS administration, supernatants were collected, and TNFα levels were measured by ELISA. Means and standard deviations of two independent experiments are shown. Figure 51E shows bar graphs depicting the results of secreted and cell-bound IL-1β from DCs treated with LPS alone (1 μg / ml), ATP alone (0.5 mM), oxPAPC alone (120 μM), or co-administered with LPS and ATP or LPS and oxPAPC. Secreted and cell-bound IL-1β were measured by ELISA 18 hours later. Means and standard deviations of two independent experiments are shown.Figure 51F shows bar graphs depicting IL-1β and TNFα secretion from DCs treated with LPS alone (1 μg / ml), ATP alone (5 mM), the indicated phospholipids alone (120 μM), or pre-stimulated with LPS for 3 hours followed by treatment with ATP or the indicated phospholipids. IL-1β and TNFα secretion were measured by ELISA 18 hours after LPS administration. Means and standard deviations of two independent experiments are shown. Figure 51G shows bar graphs depicting IL-1β and TNFα secretion from DCs and MΦs (pre-treated or not with IFNγ) treated with LPS alone (1 μg / ml), ATP alone (1 mM for DCs and 5 mM for MΦs), oxPAPC alone (120 μM), or pre-treated with LPS for 3 hours followed by activation with ATP or oxPAPC. Secreted IL-1β and TNFα were measured by ELISA after 18 hours. Means and standard deviations of two replicates from one representative experiment are shown. Figure 51H shows blots demonstrating that DCs and MΦs expressed different levels of ASC protein. DCs and MΦs were treated with or without LPS for 4 hours. ASC protein content was assessed by Western analysis using total lysates. Figure 51I shows bar graphs depicting gene expression results for DCs and MΦs treated with or without LPS for 4 hours. Expression of ASC, Nlrp3, caspase-1 (Casp-1), and caspase-11 (Casp-11) was measured by qPCR. Gene expression levels relative to TBP are shown. Line graphs represent the mean of triplicate measurements from one representative experiment. [Figure 52]Figures 52A-52I show images demonstrating that oxPAPC binds to caspase-11 and regulates inflammasome activation in DCs (see also Figures 46A-46G). Figure 52A shows a line graph depicting the formation of ASC- and caspase-1-containing specks in DCs pre-stimulated with LPS (1 μg / ml) and then activated with ATP (1 mM) or oxPAPC (120 μM). ASC- and caspase-1-containing speck formation was assessed at the indicated times. Data represent the mean and standard deviation of three fields containing approximately 50 cells obtained in three independent experiments. Figure 52B shows a bar graph depicting the formation of ASC- and caspase-1-containing specks in DCs pre-stimulated with LPS (1 μg / ml) and then activated with ATP (1 mM) or oxPAPC (120 μM). ASC- and caspase-1-containing speck formation was assessed 18 hours later. Data represent the mean and standard deviation of three independent experiments. Figure 52C shows bar graphs depicting IL-1β and TNFα secretion results from DCs of the indicated genotypes treated with Pam3CSK (P3C) alone (1 μg / ml), ATP alone (0.5 mM), oxPAPC alone (120 μM), or pre-stimulated with Pam3CSK for 3 hours followed by treatment with ATP or oxPAPC. Eighteen hours after Pam3CSK administration, IL-1β and TNFα secretion were measured by ELISA. Means and standard deviations of two independent experiments are shown. Figure 52D shows bar graphs depicting IL-1β and TNFα secretion from DCs treated with CpG alone (1 μM), oxPAPC alone (120 μM), or pre-stimulated with CpG for 3 hours followed by treatment with oxPAPC. Eighteen hours after CpG administration, IL-1β and TNFα secretion were measured by ELISA. The mean and standard deviation of two independent experiments are shown.Figure 52E shows bar graphs depicting secreted and cell-bound IL-1β levels in DCs derived from C3H / HeSNJ (WT C3H) and C3H / HeJ (TLR4 mutant C3H) mice treated with Pam3CSK (P3C) alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with Pam3CSK for 3 hours followed by oxPAPC treatment. Eighteen hours after LPS administration, secreted (left panel) and cell-bound (right panel) IL-1β were measured by ELISA. The mean and standard deviation of one biological replicate from one representative experiment out of two are shown. Figure 52F shows line graphs depicting the infectious virus load in the eyes of WT C57BL / 6 (WT) or caspase-11 KO mice infected with HSV-1 in the eyes. At the indicated times, the infectious virus load in the eyes was measured by plaque assay. Figure 52G shows blots depicting caspase-11 levels in lysates from 293T cells expressing the indicated caspase-11 alleles incubated with biotinylated ligand and streptavidin beads. Caspase-11 retained by the biotinylated ligand and input were detected by Western analysis. A representative blot from three independent experiments is shown. The caspase-11 alleles used were as follows: caspase-11 (WT), CARD domain (1–92 a.a.), and delta CARD (ΔCARD) domain (93–373 a.a.). Figures 52H–I show graphs depicting the enzymatic activity of recombinant caspase-11 monomers or multimers mixed with the indicated lipids. Enzymatic activity was monitored over time by spectrofluorometry. [Figure 53]Figures 53A-53D are images showing that inflammasome component expression was similar in WT and CD14KO DCs (related to Figures 47A-47E). Figure 53A shows bar graphs depicting IL-18 levels in WT, CD14KO, and caspase-11 KO DCs treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), ATP alone (1 mM), or pre-stimulated with LPS for 3 hours followed by oxPAPC or ATP. IL-18 in the supernatants was measured by ELISA after 18 hours. Figure 53B shows flow cytometry data demonstrating MHC class II and CD40 levels in splenic DCs treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), or pre-stimulated with LPS for 3 hours followed by oxPAPC. After 18 hours, MHC class II and CD40 levels were measured by flow cytometry. Figure 53C shows a bar graph depicting gene expression results for WT and CD14 KO DCs treated with LPS or Pam3CSK (1 μg / ml). Gene expression in response to TBP was analyzed by qPCR at the indicated times. In all experiments, untreated cells were used as a negative control. Figure 53D shows a bar graph depicting secreted and cell-bound IL-1β from DCs pretreated or not with chloroquine (10 μM) for 30 minutes, and then treated with LPS alone (1 μg / ml), oxPAPC alone (120 μM), or LPS-primed followed by oxPAPC treatment. Secreted and cell-bound IL-1β were measured by ELISA after 18 hours. [Figure 54]Figures 54A-54C are images showing that oxPAPC did not induce pyroptosis in DCs but promoted enhanced T cell activation (see also Figures 48A-48G). Figure 54A shows a line graph depicting the pyroptotic cell levels in WT DCs treated with LPS alone (1 μg / ml) or pre-stimulated with LPS for 3 hours followed by treatment with oxPAPC (120 μM) or ATP (5 mM). Pyroptosis induction was assessed up to 18 hours after stimulus addition. Pyroptotic cells were identified by flow cytometry as Annexin V and 7-AAD double-positive cells. Data are representative of two independent experiments. Figure 54B shows a bar graph showing IL-2 secretion by CD4+ T cells isolated from draining lymph nodes of WT, caspase-1 / -11 dKO, or caspase-11 KO mice 40 days after immunization with OVA + LPS in IFA(LPS), OVA + LPS + oxPAPC in IFA(LPS) or OVA + oxPAPC in IFA(oxPAPC). CD4+ T cells were either restimulated with OVA in the presence of DCs as antigen-presenting cells or not. IL-2 secretion was measured by ELISA 5 days later. Bar graphs show the mean and standard error of two experiments with five animals per group. Figure 54C shows a bar graph demonstrating that oxPAPC enhanced effector T cell responses in vivo. CD4+ T cells were isolated from draining lymph nodes of wild-type, caspase-1 / -11 dKO, or caspase-11 KO mice 7 days after immunization with OVA + LPS in IFA (LPS), OVA + LPS + oxPAPC in IFA (LPS + oxPAPC), or OVA + oxPAPC in IFA (oxPAPC). CD4+ T cells were either restimulated with OVA or not in the presence of DCs as antigen-presenting cells. IFNγ (upper panel) and IL-17 (lower panel) secretion were measured by ELISA 5 days later. Bar graphs represent the mean and standard error of four experiments with three animals per group. DETAILED DESCRIPTION OF THE INVENTION

[0031] The present invention relates, at least in part, to the unexpected observation that PAPC lipids, particularly oxidized PAPC lipids (oxPAPC), function as specific activators of the inflammatory response in dendritic cells (DCs). In particular, we have identified 1-palmitoyl-2-arachidonyl-sn-glycero-3-phosphorylcholine (PAPC) and its oxidized variant (oxPAPC) as the first specific activators of the inflammatory response in dendritic cells (DCs). In the presence of TLR ligands, oxPAPC promoted DC survival and triggered the release of the T cell-activating cytokine interleukin-1 beta (IL-1β).

[0032] Without wishing to be bound by theory, mechanistically, oxPAPC binds to the LPS receptor CD14 on the surface of DCs, promoting their delivery into endosomes and subsequent access to the cytosolic protein caspase-11. Binding of oxPAPC to caspase-11 triggered inflammasome-mediated IL-1β release. Notably, these oxPAPC-induced responses did not occur in macrophages, indicating that this lipid acts uniquely to promote the immunomodulatory activity of DCs rather than a general (macrophage-mediated) inflammatory response. Consequently, oxPAPC synergized with microbial products to induce more robust antigen-specific T cell activation than could be elicited by PAMPs alone. Thus, oxPAPC has been identified as a member of a new class of immunomodulatory factors (termed "vita-DAMPs") that function together with PAMPs to promote DC survival and elicit maximal adaptive immune responses.

[0033] DCs are the most potent activators of protective (adaptive) immunity, and the design of vaccine adjuvants that selectively promote DC-mediated immunity is currently under intensive research. All current FDA-approved vaccine adjuvants are unable to specifically activate DCs. They all promote a general inflammatory response in various immune cells, including macrophages and DCs.

[0034] The present discovery that PAPC can specifically activate DCs identifies this molecule as a lead candidate for a next-generation vaccine adjuvant. While PAPC has been studied previously by other research groups, most of the work in this area has focused on PAPC's ability to act as an anti-inflammatory molecule. The present discovery that PAPC acts to promote immunity, rather than inhibit it, distinguishes the uses of PAPC described and exemplified herein from previous suggestions of the therapeutic value of such molecules.

[0035] A key finding in identifying this invention was the discovery that co-administration of PAPC with microbial products could promote exclusively DC-mediated immune responses. This co-administration generated a previously unobserved state of DCs. This novel cellular behavior is predicted to have significant therapeutic potential.

[0036] definition The term "oxPAPC" or "oxidized PAPC," as used herein, refers to lipids produced by oxidation of 1-palmitoyl-2-arachidonyl-sn-glycero-3-phosphorylcholine (PAPC). Oxidation results in a mixture of oxidized phospholipids containing either fragmented or full-length oxygenated sn-2 residues. Well-characterized oxidatively fragmented species contain five-carbon sn-2 residues bearing omega-aldehyde or omega-carboxyl groups. Oxidation of arachidonic acid residues also produces phospholipids containing esterified isoprostanes. oxPAPC includes HOdiA-PC, KOdiA-PC, HOOA-PC, and KOOA-PC species, among other oxidation products present in oxPAPC.

[0037] The term "non-canonical inflammasome-activating lipids," as used herein, refers to lipids capable of inducing a cellular caspase-11-dependent inflammasome inflammatory response. Exemplary "non-canonical inflammasome-activating lipids" include PAPC, oxPAPC, and species of oxPAPC (e.g., HOdiA-PC, KOdiA-PC, HOOA-PC, KOOA-PC), as well as Rhodo LPS (LPS-RS or LPS derived from Rhodobacter sphaeroides).

[0038] "Immunogen" and "antigen" are used interchangeably and refer to any compound that will elicit a cellular or humoral immune response. Non-living immunogens include, for example, killed immunogens, subunit vaccines, recombinant proteins, or peptides. The adjuvants of the present invention can be used with any suitable immunogen. Exemplary immunogens of interest include those that constitute or are derived from viruses, mycoplasmas, parasites, protozoans, or prions, etc. Thus, immunogens of interest may be derived from, but are not limited to, human papillomavirus, herpesviruses such as herpes simplex or varicella zoster, retroviruses such as human immunodeficiency virus type 1 or 2, hepatitis virus, influenza virus, rhinovirus, respiratory syncytial virus, cytomegalovirus, adenovirus, Mycoplasma pneumoniae, Salmonella, Staphylococcus, Streptococcus, Enterococcus, Clostridium, Escherichia, Klebsiella, Vibrio, Mycobacterium bacteria, amoeba, Plasmodium, and / or Trypanosoma cruzi. It is further contemplated that the adjuvant lipids of the present invention can be co-administered with tumor antigens or other cancer antigens, thereby providing an immunostimulatory cancer therapy / cancer vaccine.

[0039] "Co-administered," as used herein, means that two compounds are administered sufficiently close in time to achieve a combined immune effect. Thus, co-administration may be achieved by sequential administration or simultaneous administration (e.g., co-administration in a common carrier or the same carrier).

[0040] For example, "modulation" of a molecule's symptoms, level, or biological activity refers to, for example, a detectable increase or decrease in symptoms or activity. Such an increase or decrease can be observed in a subject treated with an adjuvant lipid (non-canonical inflammasome-activating lipid) of the present invention compared to a subject not treated. In this case, the untreated subject (e.g., a subject administered an immunogen in the absence of an adjuvant lipid) has or is prone to develop the same or similar disease or infection as the treated subject. Such an increase or decrease can be at least about 2%, 5%, 10%, 15%, 20%, 25%, 30%, 40%, 50%, 60%, 70%, 75%, 80%, 85%, 90%, 95%, 98%, 100%, 150%, 200%, 250%, 300%, 400%, 500%, or 1000% or more, or within any range between any two of these values. Modulation may be determined subjectively or objectively, e.g., by subject self-assessment, by a clinician's assessment, or by performing a suitable assay or measurement, including, e.g., assessing the degree and / or nature of a subject's immune activation achieved by an immunogen administered in the presence of an adjuvant lipid of the invention (a non-canonical inflammasome-activating lipid). Modulation may be temporary, long-term, or permanent, and may occur within a suitable time period during or after administration of an adjuvant lipid of the invention to a subject, or during or after use in an assay or other method described herein or in the cited references, e.g., within the time periods described below, i.e., from about 12 to 24 or 48 hours after administration or use of an adjuvant lipid of the invention, to about 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 21, 28 days, or 1, 3, 6, or 9 months or more after the subject has received such immunostimulatory composition / treatment.

[0041] As used herein, a "subject" includes animals with an adaptive immune system, such as humans (e.g., human subjects) and non-human animals. The term "non-human animal" includes all vertebrates, e.g., mammals, e.g., rodents, e.g., mice, and non-mammals, e.g., non-human primates, e.g., sheep, dogs, cows, chickens, amphibians, reptiles, etc.

[0042] A "suitable dose level" refers to a dose level that provides a therapeutically reasonable balance between pharmaceutical efficacy and adverse effects (e.g., between sufficient immunostimulatory activity conferred by the immunogen administered in the presence of an adjuvant lipid of the invention and a sufficiently low level of macrophage stimulation). For example, this dose level can be related to, for example, the peak or mean serum level in a subject of anti-immunogen antibodies produced after administration of a particular dose level of an immunogenic composition (comprising an adjuvant lipid of the invention).

[0043] It is understood that ranges provided herein are a shorthand for all values within that range. For example, a range of 1 to 50 is understood to include any number, combination of numbers, or subrange from the group consisting of 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47, 48, 49, or 50.

[0044] Unless otherwise stated or apparent from the context, the term "or" as used herein is understood to be inclusive.

[0045] Unless otherwise stated or apparent from the context, the terms "a," "an," and "the," as used herein, are understood to be singular or plural.

[0046] Any composition or method provided herein can be combined with one or more of any of the other compositions and methods provided herein.

[0047] Dendritic cells (DCs) and the regulation of pattern recognition receptors (PRRs) The innate immune system has classically been viewed as operating in an all-or-none fashion, with DCs either initiating or not initiating inflammatory responses that promote adaptive immunity. Therefore, TLRs expressed by DCs are thought to be crucial in determining the immunogenic potential of these cells. The mammalian immune system is involved in detecting microorganisms and activating defensive responses that contain infection. Central to this task are dendritic cells, which sense microorganisms and then promote T cell activation. It has been suggested that dendritic cells can assess any infectious threat and direct a proportionate response (Blander, JM (2014). Nat Rev Immunol 14, 601-618; Vance, RE et al., (2009) Cell Hostµbe 6, 10-21), but the mechanisms by which this immunomodulatory activity may occur remain unclear.

[0048] PRRs act either directly or indirectly to detect molecules common to a wide class of microorganisms, classically termed pathogen-associated molecular patterns (PAMPs), including factors such as bacterial lipopolysaccharide (LPS), bacterial flagellin, or viral double-stranded RNA, among others.

[0049] A key attribute of PRRs as immune regulators is their ability to recognize specific microbial products. Therefore, PRR-mediated signaling events should provide definitive indications of infection. It is hypothesized that "GO" signals are activated by PRRs expressed in DCs, which promote inflammation and T cell-mediated immunity. Interestingly, several groups have recently proposed that DCs do not simply operate in this all-or-none manner (Blander, JM, and Sander, LE (2012). Nat Rev Immunol 12, 215-225; Vance, RE et al. (2009) Cell Hostµbe 6, 10-21). Rather, DCs may have the ability to assess the threat (or virulence) of any possible infection and mount a proportionate response. The most commonly discussed means by which virulence can be assessed is based on the ability of virulent pathogens to activate a greater variety of PRRs than nonpathogens. However, not all microorganisms share a common set of PRR activators, and not all PRR activators exhibit equal potency. Therefore, the number of PRRs activated during infection may not be an ideal assessment of virulence. Furthermore, an increase in the number of PRRs activated during infection is generally associated with a greater inflammatory response, which may indirectly promote a greater T cell response. Conditions previously suggested to enhance DC activation (e.g., by using virulent pathogens as stimuli) are also expected to enhance MΦ activation (Vance, RE et al., (2009) Cell Host Microbiology 6, 10-21). Therefore, it remains unclear whether the immune system (i.e., DCs) truly possesses mechanisms for specifically assessing the threat of infection.

[0050] One possible means by which infection threat may be assessed would be the well-recognized process of coincidence detection, in which independent inputs result in responses distinct from those elicited by any single input. In the context of PRRs, one such input must be a microbial product indicative of infection, regardless of virulence threat. To assess virulence threat, a second input must be present. Without wishing to be bound by theory, we propose that this putative second input is a molecule produced at the site of tissue injury, since cellular damage is often a feature associated with highly pathogenic microorganisms. Potential candidate molecules that provide a second stimulus to DCs are a diverse family of molecules called damage-associated molecular patterns (DAMPs), also known as alarmins (Kono, H., and Rock, KL (2008) Nat Rev Immunol 8, 279-289; Pradue, T., and Cooper, EL (2012) Front Immunol 3, 287). DAMPs are found at sites of infectious and non-infectious tissue injury and have been proposed to regulate inflammatory responses, although their mechanism of action remains unclear. One such class of DAMPs is represented by oxidized phospholipids derived from 1-palmitoyl-2-arachidonyl-sn-glycero-3-phosphorylcholine (PAPC), collectively known as oxPAPC. These lipids are produced at sites of both infectious and non-infectious tissue injury (Berliner, JA, and Watson, AD (2005). N Engl J Med 353, 9-11; Imai, Y. et al. (2008) Cell 133, 235-249; Shirey, KA et al. (2013) Nature 497, 498-502) and are found at very high levels in the membranes of dying cells (Chang, MK et al., (2004) J Exp Med 200, 1359-1370).Additionally, oxPAPC is an active component of oxidized low-density lipoprotein (oxLDL) aggregates that promote inflammation in atherosclerotic tissues (Leitinger, N. (2003) Curr Opin Lipidol 14, 421-430), and local concentrations in atherosclerotic tissues can be as high as 10-100 μM (Oskolkova, OV et al. (2010) J Immunol 185, 7706-7712). The association between oxPAPC and dying cells raises the possibility that these lipids may serve as a global indicator of tissue health. Thus, in the presence of microbial products, oxPAPC may signal an increased threat of infection.

[0051] While not wishing to be bound by theory, mechanistically, the receptor CD14 is thought to capture lipids such as oxPAPC and PAPC and deliver them to intracellular locations where they activate the non-canonical inflammasome (caspase-11-dependent inflammasome). Inflammasome-mediated activity then synergizes with independently occurring TLR signaling events to promote more robust T cell responses than those induced by TLR ligands alone. Thus, oxidized lipids may alert dendritic cells that they have encountered highly infectious microorganisms, enabling them to promote adaptive responses commensurate with the threat of infection.

[0052] Toll-like receptors Toll-like receptors (TLRs) are type I transmembrane receptors that are evolutionarily conserved between insects and humans. Ten TLRs (TLRs 1-10) have been identified (Sabroe, I. et al., (2003) Journal of Immunology 171(4):1630-5). Members of the TLR family share similar extracellular and intracellular domains. The extracellular domain has been shown to contain leucine-rich repeats, and the intracellular domain is similar to the intracellular region of the interleukin-1 receptor (IL-1R). TLRs are differentially expressed in immune cells and other cells, including vascular epithelial cells, adipocytes, cardiac myocytes, and intestinal epithelial cells. The intracellular domain of TLRs interacts with the adaptor protein Myd88, which also contains an IL-1R domain in its cytoplasmic region, leading to cytokine activation of NF-κB. This Myd88 pathway is one pathway by which cytokine release is achieved through TLR activation. TLRs are primarily expressed on cell types such as antigen-presenting cells (e.g., dendritic cells, macrophages, etc.). One such TLR is TLR4, which is involved in the activation of the innate immune system and recognizes lipopolysaccharide (LPS), a component of Gram-negative bacteria. TLR4 has been shown to interact with lymphocyte antigen 96, Myd88 (myeloid differentiation primary response gene 88), and TOLLIP (toll-interacting protein).

[0053] Activation of dendritic cells by stimulation via TLRs leads to dendritic cell maturation and the production of proinflammatory cytokines such as IL-12. Previous studies have shown that TLRs recognize various types of agonists, but some agonists are common to several TLRs. TLR agonists are mainly derived from bacteria or viruses and include molecules such as flagellin or bacterial lipopolysaccharide (LPS).

[0054] Two states of DC activation Herein, we identified two states of DC activation: the first activation state was mediated by encounter with microbial products, such as TLR ligands, which activated TLRs to release cytokines, upregulate costimulatory molecules, and promote MHC-mediated antigen presentation. All of these were important for T cell activation. However, because these TLR ligands were common to both pathogens and nonpathogens, they could not be used to assess threat to the host. The second state of DCs was considered "hyperactive" and was mediated by simultaneous encounter with microbial products and oxidized phospholipids, which were abundant at sites of tissue damage. Concordant detection of TLR ligands and oxidized lipids (e.g., oxPAPC) promoted all activities induced by the classical activation state and induced inflammasome-mediated release of IL-1β, a potent T cell activator. Neither TLR ligands nor oxPAPC alone had the ability to induce IL-1β release. This observation provided formal experimental evidence that the innate immune system uses the principle of coincident detection to induce a hyperactive state in DCs.

[0055] An intriguing aspect of the hyperactive DC state is the mechanism by which it is induced. Classical activation was induced by microbial products, whereas the hyperactive state was induced by both microbial and autologous products. This self-referential aspect of immune activation was not entirely unprecedented, as T cell maturation and maintenance have previously been shown to depend on interactions with MHC molecules carrying microbial and self-peptides (Janeway, CA, Jr. (2002) Annu Rev Immunol 20, 1-28). Mechanistically, analysis of oxPAPC revealed that this molecule is a selective endogenous mimic of LPS in that it bound to and activated the LPS receptors CD14 and caspase-11. Interestingly, oxPAPC did not induce TLR4 dimerization, endocytosis, myosin formation, or gene expression. Indeed, when administered before microbial encounter, oxPAPC acted as a TLR4 antagonist (Bochkov, V. et al., (2002) Nature 419, 77-81; Erridge, C. et al., (2008) The Journal of biological chemistry 283, 24748-24759; Oskolkova, O. et al. (2010) J Immunol 185, 7706-7712). Thus, these data together reveal an intriguing cellular process in which CD14 functions to coordinate the activities of TLR4 and caspase-11 by delivering either PAMP (LPS) or DAMP (oxPAPC) to their respective receptors.

[0056] Several observations supported this proposed CD14-caspase-11 pathway as a central step leading to DC hyperactivation. First, oxPAPC formed a complex with CD14 and caspase-11 in vitro. Second, genetic deletion of CD14 and caspase-11 phenocopied each other, in that loss of either protein caused DC dysfunction and IL-1β release in response to oxPAPC treatment. In contrast, neither CD14 nor caspase-11 was required for ATP-mediated IL-1β release. Third, neither of these proteins was required for the priming step of inflammasome activation, as assessed by normal levels of expression of various TLR-dependent cytokines. Fourth, binding of oxPAPC to CD14 promoted endocytosis and delivery of this lipid to intracellular caspase-11. This statement was supported by the ability of cytosolic transfection of oxPAPC to rescue the defect in IL-1β release in CD14 KO, providing conclusive evidence that the trafficking function of CD14 was important for caspase-11 activation.

[0057] Caspase-11 has attracted much attention in recent years due to its ability to promote IL-1β release and pyroptosis in response to Gram-negative cytosolic bacteria (Hagar, JA et al., (2013) Science 341, 1250-1253; Kayagaki, N. et al. (2013) Science 341, 1246-1249). This selective promotion of immune responses to Gram-negative bacteria by caspase-11 is explained by its newly recognized ability to act as a bona fide LPS receptor (Shi, J. et al., (2014a) Nature 514, 187-192). oxPAPC binds to caspase-11, extending the role of caspase-11 beyond its function as an LPS receptor. Indeed, caspase-11 was required for oxPAPC-mediated IL-1β release in the absence of TLR4 ligands, for example, when cells were stimulated with ligands that often bind to Gram-positive bacteria (i.e., Pam3CSK) or viruses (CpG DNA).

[0058] Based on these data, caspase-11 had a fundamental function as an indicator of toxic threat to DCs. Threat was assessed in two ways. First, caspase-11, an autologous indicator of damage, could be activated during encounters with any pathogen that causes tissue damage and cell death due to its ability to bind oxPAPC. Therefore, this activity would provide DCs with a fundamental mechanism for hyperactivation during infection with virulent microorganisms. Second, in the case of bacteria encoding type III and type IV secretion systems that deliver LPS directly to the cytosol (Hagar, JA, and Miao, EA (2014) Curr Opin Microbiol 17, 61-66), caspase-11 likely hyperactivated DCs even before tissue damage occurred. Under these latter conditions, delivery of LPS to the cytosol by the virulence-associated secretion system would not have been involved in the transport function of CD14. Indeed, we found herein that the genetic requirement for CD14 for inflammasome activation could be bypassed by direct transfection of LPS or oxPAPC into the cytosol. In contrast, spontaneous delivery of oxPAPC from the extracellular medium to caspase-11 was dependent on CD14. This critical role of CD14 in mediating caspase-11 activation suggested that this protein has a broader function in inducing adaptive immunity than would be expected from its role as an LPS receptor. Rather, CD14 and caspase-11 are general regulators of immunity against a wide range of pathogens. This model was supported in vivo by the finding that HSV-1 proliferation in the eye was suppressed by the action of caspase-11.

[0059] Furthermore, mechanistic studies revealed that oxPAPC differed from LPS in its effects on caspase-11 in several fundamental respects. First, both lipids bound to caspase-11 and induced its multimerization, but LPS bound to the CARD, whereas oxPAPC bound to the catalytic domain. The functional consequences of these different binding mechanisms were that binding to the CARD promoted caspase-11 enzymatic activity, whereas binding to the catalytic domain inhibited it. Because caspase-11 enzymatic activity is required for pyroptosis, oxPAPC should not kill cells. Indeed, population- and single-cell-based analyses determined that oxPAPC did not kill cells and identified the presence of inflammasomes in viable DCs exposed to oxPAPC. In contrast, in DCs exposed to ATP, inflammasomes were present only in dead cells. Indeed, oxPAPC promoted the viability of DCs also exposed to LPS. Although there were several examples of endogenous molecules that bound PRRs, recent information suggested that the mode of interaction was similar to that mediating microbial interactions (or was unknown). Thus, caspase-11 was a unique PRR in that it contained separate domains that interacted with PAMPs (LPS) and endogenous molecules (oxPAPC). These different mechanisms of interaction resulted in distinct cellular responses, suggesting that, like DCs, PRRs also had distinct states of activation.

[0060] The dual activity of oxPAPC in promoting inflammasome activation and DC survival indicated that such activities played a role in enhancing adaptive immune responses. Indeed, we identified LPS / oxPAPC as a superior adjuvant to LPS alone in eliciting antigen-specific effector and memory T cells in vivo. In other cases, inflammasome activation occurring independently of cell death has also been observed (Broz, P. et al., (2010) Cell Host Microbe 8, 471-483; Ceballos-Olvera, I. et al., (2011) PLoS Pathog 7, e1002452; Schmidt, RL, and Lenz, LL (2012) PLoS One 7, e45186). Ongoing studies are investigating the mechanism by which death and IL-1β release are coupled. Based on its unique ability to function as both an inflammasome-promoting and pro-survival stimulus, we conclude herein that oxPAPC can be considered a vita-DAMP that functions to promote DC survival and the initiation of adaptive immunity. Because vita-DAMPs promote cell survival rather than pyroptotic cell death, they can be operationally distinguished from traditionally defined DAMPs, such as ATP. We also contemplate herein that other known TLR4 antagonists may be selective LPS mimics with similar activity as oxPAPC. Note that oxPAPC was also released under non-infectious conditions. Under these conditions, the ability of oxPAPC to promote CD14 endocytosis likely helped limit TLR4-dependent inflammatory responses that could be misactivated by other DAMPs present at the site of injury (Mancek-Keber, M., et al. (2015) Science Signaling 8, ra60). These context-dependent activities of oxPAPC, either anti-inflammatory or pro-inflammatory, identified their importance in helping DC assess the source of injury in a given tissue.

[0061] In summary, we herein identified a means by which endogenous self-molecules can create a hyperactive state in DCs through their ability to bind caspase-11 in an atypical manner. The existence of this hyperactive state revealed that the innate immune system acted by a mechanism in which the threat of infection was assessed by coincident detection of PAMPs and vita-DAMPs.

[0062] Adjuvants and Vaccines Immunogenic compositions containing the adjuvants of the present invention can be administered to a subject using any known vaccine format, e.g., attenuated virus, protein, nucleic acid, etc., such that the subject produces an effective amount of the selected immunogen to induce a therapeutic or prophylactic immune response against the target antigen. The subject may be a human or non-human subject. Animal subjects include, but are not limited to, non-human primates, dogs, cats, horses, ruminants (e.g., sheep, goats, cattle, camels, alpacas, llamas, deer), pigs, birds (e.g., chickens, turkeys, quails), rodents, and chirodoptera. Subjects can be treated for any purpose, including, but not limited to, eliciting a protective immune response or producing antibodies (or B cells) for collection and use for other purposes.

[0063] In one embodiment, the invention features adjuvanted microbial vaccines. Microbial vaccines are often composed of cell wall components that allow the immune system to recognize the whole organism, or, in the case of bacteria that cause disease through toxicity, such as diphtheria, toxins or derived toxoids may be used. Antitoxins are under development for several disease organisms, primarily for therapeutic use. Bacteria can be grown in liquid or solid media cultures, harvested, purified, and used directly as killed or attenuated vaccines.

[0064] Optionally, the immunogen of interest is expressed in disease target cells (e.g., tumor cells, infected cells) but less expressed or not expressed at all in other tissues. Examples of target cells include cells derived from neoplastic diseases, including, but not limited to, sarcoma, lymphoma, leukemia, carcinoma, melanoma, breast cancer, prostate cancer, ovarian cancer, cervical cancer, colon cancer, lung cancer, glioblastoma, and astrocytoma. Alternatively, the target cells may be infected with, for example, viruses, mycoplasmas, parasites, protozoans, and prions. Thus, immunogens of interest may be derived from, but are not limited to, human papillomavirus (see below), herpesviruses such as herpes simplex or varicella zoster, retroviruses such as human immunodeficiency virus type 1 or 2, hepatitis virus, influenza virus, rhinovirus, respiratory syncytial virus, cytomegalovirus, adenovirus, Mycoplasma pneumoniae, Salmonella, Staphylococcus, Streptococcus, Enterococcus, Clostridium, Escherichia, Klebsiella, Vibrio, Mycobacterium bacteria, amoeba, malaria parasites, and Trypanosoma cruzi.

[0065] In addition to tumor antigens and antigens of infectious agents, tumor suppressor gene products, including but not limited to p53, BRCA1, BRCA2, retinoblastoma, and TSG101, or mutant oncogene products, such as but not limited to RAS, WT, MYC, ERK, and TRK, can also provide target antigens for use in accordance with the present invention. Target antigens may also be autoantigens, such as those associated with cancer or neoplastic disease. In one embodiment of the present invention, the immunogen is a peptide derived from a heat shock protein (hsp)-peptide complex of a diseased cell, or the hsp-peptide complex itself.

[0066] In some embodiments, immunogens may be purified from natural sources, obtained by recombinant expression, or directly synthesized. In some embodiments, immunogens may be provided by whole cells, microorganisms, or virus particles, which may be live, attenuated, or killed. In other embodiments, immunogens may comprise protein fragments that contain one or more immunogenic regions in the molecule.

[0067] Immunogens include those that have been modified or derivatized, such as by attaching or coupling one or more groups to enhance the subject's immune response. Examples of immunogenic carrier proteins are KLH and BSA. Immunogenic carriers also include polypeptides that are broad class II activators (see, e.g., Panina-Bordignon et al., Cold Spring Harb Symp Quant Biol 1989). Linkages for conjugation are prepared by methods well known to those skilled in the art.

[0068] The immunogenic compositions of the present invention comprise an immunogen and an adjuvant lipid and can be administered for therapeutic and / or prophylactic purposes. In therapeutic applications, the immunogenic compositions of the present invention are administered in an amount sufficient to induce an immune response effective in treating or halting the progression and / or symptoms of disease. The dosage of the adjuvant of the present invention will vary depending on the nature of the immunogen and the condition of the subject. However, it should be sufficient to enhance the efficacy of the immunogen in eliciting an immunogenic response. For therapeutic or prophylactic treatments, the amount of adjuvant administered may range from 0.05, 0.1, 0.5, or 1 mg per kg of body weight to about 10, 50, or 100 mg per kg of body weight or more. The adjuvants of the present invention are generally non-toxic and can generally be administered in relatively large amounts without causing life-threatening side effects.

[0069] The term "therapeutic immune response" as used herein refers to an increase in humoral and / or cellular immunity against a target antigen, as measured by standard techniques. Preferably, the level of induced immunity against the target antigen is at least 4-fold, preferably at least 16-fold, as compared to the level before administration of the immunogen. The immune response can also be measured qualitatively. In this case, the arrest or reduction of the progression of a tumor or infectious disease in a subject by a suitable in vitro or in vivo assay is considered to indicate the induction of a therapeutic immune response.

[0070] In the methods of the present invention, a composition comprising an immunogen and adjuvant of the present invention combined in a therapeutically effective amount is administered to a mammal in need thereof. The term "administering," as used herein, means delivering the immunogen and adjuvant of the present invention to a mammal by any method capable of achieving the desired result. The immunogen and adjuvant of the present invention can be administered, for example, intravenously or intramuscularly. The term "mammal," as used herein, is intended to include, but is not limited to, humans, laboratory animals, domestic pets, and livestock. A "therapeutically effective amount" refers to an amount of immunogen and adjuvant effective to produce a desired therapeutic effect when administered to a mammal.

[0071] Compositions comprising the immunogens and adjuvants of the invention can be administered dermally, subcutaneously, intravenously, intramuscularly, parenterally, pulmonary, vaginally, rectally, nasally, or topically. Compositions may also be delivered by injection, orally, by spray, or by particle bombardment.

[0072] The composition to be administered may further contain various additional substances, such as pharmaceutically acceptable carriers. Suitable carriers include any of the standard pharmaceutically acceptable carriers, such as phosphate-buffered saline solution, water, emulsions such as oil / water emulsions or triglyceride emulsions, various types of wetting agents, tablets, coated tablets, and capsules. Typically, such carriers contain excipients such as starch, milk, sugar, certain types of clay, gelatin, stearic acid, talc, vegetable oils and fats, gums, glycols, or other known excipients. Such carriers may also contain flavorings, coloring agents, or other ingredients. The compositions of the present invention may also contain suitable diluents, preservatives, solubilizers, emulsifiers, adjuvants, and / or carriers. Such compositions may be in liquid form or may be lyophilized or otherwise dry formulations, and may contain diluents varying in buffer content (e.g., Tris-HCl, acetate, phosphate), pH, and ionic strength, additives such as albumin or gelatin to prevent absorption to surfaces, surfactants (e.g., Tween 20, Tween 80, Pluronic F68, bile salts), solubilizers (e.g., glycerol, polyethyleneglycerol), antioxidants (e.g., ascorbic acid, sodium metabisulfite), and the like. These compositions may include preservatives (e.g., thimerosal, benzyl alcohol, parabens), bulking agents or tonicity modifiers (e.g., lactose, mannitol), covalent attachment of polymers such as polyethylene glycol to proteins, complexation with metal ions, or incorporation of substances into or onto particulate preparations of polymeric compounds such as polylactic acid, polyglycolic acid, hydrogels, or onto liposomes, microemulsions, micelles, unilamellar or multilamellar vesicles, erythrocyte ghosts, or spheroplasts. Such compositions will influence the physical state, solubility, stability, rate of in vivo release, and rate of in vivo clearance.

[0073] Pharmaceutical Composition In one embodiment, the present invention provides a pharmaceutical composition comprising an immunogen and an adjuvant lipid as identified herein. The immunostimulatory composition may be suitably formulated and introduced into the environment of a subject or cell by any means recognized for such delivery.

[0074] Such compositions typically include an active agent and a pharmaceutically acceptable carrier. As used herein, the term "pharmaceutically acceptable carrier" includes saline, solvents, dispersion media, coatings, antibacterial and antifungal agents, isotonic and absorption delaying agents, and the like, that are compatible with pharmaceutical administration. Complementary active compounds can also be incorporated into the compositions.

[0075] Pharmaceutical compositions are formulated to be compatible with their intended route of administration. Examples of routes of administration include parenteral, e.g., intravenous, intradermal, subcutaneous, oral (e.g., inhalation), transdermal (topical), transmucosal, and rectal administration. Solutions or suspensions used for parenteral, intradermal, or subcutaneous administration may contain the following components: a sterile diluent such as water for injection, saline solution, fixed oils, polyethylene glycol, glycerin, propylene glycol, or other synthetic solvents; an antibacterial agent such as benzyl alcohol or methylparaben; an antioxidant such as ascorbic acid or sodium bisulfite; a chelating agent such as ethylenediaminetetraacetic acid; a buffer such as acetate, citrate, or phosphate, and an agent for adjusting tonicity such as sodium chloride or dextrose. pH can also be adjusted with acids or bases such as hydrochloric acid or sodium hydroxide. Parenteral preparations can be enclosed in glass or plastic ampoules, disposable syringes, or multiple-dose vials.

[0076] Pharmaceutical compositions suitable for injectable use include sterile aqueous solutions (where water soluble) or dispersions and sterile powders for the extemporaneous preparation of sterile injectable solutions or dispersions. For intravenous administration, suitable carriers include physiological saline, bacteriostatic water, Cremophor EL™ (BASF, Parsippany, NJ), or phosphate-buffered saline (PBS). In all cases, the composition must be sterile and should be fluid to the extent that easy syringability exists. The composition should be stable under the conditions of manufacture and storage and preserved against the contaminating action of microorganisms such as bacteria and fungi. The carrier can be a solvent or dispersion medium containing, for example, water, ethanol, polyol (e.g., glycerol, propylene glycol, and liquid polyethylene glycol, etc.), and suitable mixtures thereof. Proper fluidity can be maintained, for example, by the use of a coating such as lecithin, by the maintenance of the required particle size in the case of dispersions, and by the use of surfactants. Prevention of the action of microorganisms can be achieved by various antibacterial and antifungal agents, for example, parabens, chlorobutanol, phenol, ascorbic acid, and thimerosal. In many cases, it will be preferable to include isotonic agents, for example, sugars; polyalcohols such as manitol, sorbitol, sodium chloride, and the like, in the composition. Prolonged absorption of injectable compositions can be brought about by including in the composition an agent which delays absorption, for example, aluminum monostearate and gelatin.

[0077] Sterile injectable solution can be prepared by incorporating the required amount of active compound into the selected solvent with one or combination of the above-mentioned components, and then optionally sterilize by filtration.Generally, dispersion system is prepared by incorporating active compound into a sterile medium that contains basic dispersion medium and other necessary components from above-mentioned.For the sterile powder that is used to prepare sterile injectable solution, the preferred method of preparation is vacuum drying and freeze-drying, thereby obtaining the powder of active compound and any additional desired components from the solution that has been previously sterilized and filtered.

[0078] Oral compositions generally include an inert diluent or an edible carrier. For oral therapeutic administration, the active compound may be incorporated with excipients and used in the form of tablets, troches, or capsules, such as gelatin capsules. Oral compositions can also be prepared using flowable carriers for use as mouthwashes. Pharmaceutically compatible binders and / or adjuvant substances may be included as part of the composition. Tablets, pills, capsules, and lozenges may contain any of the following ingredients or compounds of a similar nature: binders such as microcrystalline cellulose, tragacanth, or gelatin; excipients such as starch or lactose; disintegrants such as alginic acid, Primogel, or cornstarch; lubricants such as magnesium stearate or SteroTe; glidants such as colloidal silicon dioxide; sweeteners such as sucrose or saccharin; or flavorings such as peppermint, methyl salicylate, or orange flavoring.

[0079] The compositions of the present invention can also be formulated as nanoparticle formulations.

[0080] The compounds of the invention can be administered in immediate, delayed, modified, sustained, pulsed, or controlled release dosage forms.

[0081] Pharmaceutical compositions of the invention may contain from 0.01 to 99% by weight / volume of the active material.

[0082] For administration by inhalation, the compounds are delivered in the form of an aerosol spray from pressured container or dispenser which contains a suitable propellant, e.g., a gas such as carbon dioxide, or a nebulizer, methods including those described in U.S. Patent No. 6,468,798.

[0083] Systemic administration can also be via transmucosal or transdermal means.For transmucosal or transdermal administration, a penetrant appropriate to the barrier to be permeated is used in the formulation.Such penetrants are generally known in the art, and for example, for transmucosal administration, surfactants, bile salts, and fusidic acid derivatives are included.Transmucosal administration can be achieved by using nasal sprays or suppositories.For transdermal administration, the active compound is formulated into ointments, salves, gels, or creams as generally known in the art.

[0084] The compounds can also be prepared in the form of suppositories (eg, with conventional suppository bases such as cocoa butter and other glycerides) or retention enemas for rectal delivery.

[0085] In one embodiment, the active compounds are prepared with carriers that will protect the compound from rapid elimination from the body, such as controlled-release formulations, including implants and microencapsulated delivery systems. Biodegradable, biocompatible polymers can be used, such as ethylene vinyl acetate, polyanhydrides, polyglycolic acid, collagen, polyorthoesters, and polylactic acid. Such formulations can be prepared using standard techniques. These materials are also commercially available from Alza Corporation and Nova Pharmaceuticals, Inc. Liposomal suspensions (including liposomes targeted to infected cells using monoclonal antibodies against viral antigens) can also be used as pharmaceutically acceptable carriers. These can be prepared by methods known to those skilled in the art, for example, as described in U.S. Pat. No. 4,522,811.

[0086] The toxicity and therapeutic efficacy of such compounds can be determined in cell cultures or experimental animals by standard pharmaceutical procedures, for example, to determine LD50 (the dose lethal to 50% of a population) and ED50 (the dose therapeutically effective in 50% of a population). The dose ratio between toxic and therapeutic effects is the therapeutic index, which can be expressed as the ratio LD50 / ED50. Compounds that exhibit high therapeutic indices are preferred. Compounds that exhibit toxic side effects can also be used, but care should be taken to design a delivery system that targets such compounds to the site of affected tissues in order to minimize the possibility of damage to non-infected cells, thereby reducing side effects.

[0087] Data obtained from cell culture assays and animal studies can be used to formulate a dosage range for use in humans. The dosage of such compounds is preferably within a circulating concentration range that includes the ED50 with little or no toxicity. Dosages may vary within this range depending on the dosage form and route of administration used. For compounds used in the methods of the present invention, therapeutically effective doses can be initially estimated using cell culture assays. Doses can be formulated in animal models to achieve a circulating plasma concentration range that includes the IC50 (i.e., the concentration of the test compound that achieves half-maximal inhibition of symptoms) determined in cell culture. Such information can be used to more accurately determine useful doses for humans. Plasma levels can be measured, for example, by high-performance liquid chromatography.

[0088] As defined herein, a therapeutically effective amount (i.e., effective dose) of an adjuvant-containing compound of the invention targeted to a disease or disorder depends on the immunogen selected and the target disease or disorder. For example, the amount of immunogen in a single dose of an immunogen-adjuvant composition of the invention targeted to a disease or disorder can be administered in the range of approximately 1 pg to 1000 mg. In some embodiments, 10, 30, 100, or 1000 pg, or 10, 30, 100, or 1000 ng, or 10, 30, 100, or 1000 μg, or 10, 30, 100, or 1000 mg may be administered. In some embodiments, 1 to 5 g of the composition can be administered.

[0089] The therapeutically effective amount of the compounds of the present invention can be determined by methods known in the art. In addition to depending on the immunogen used, the therapeutically effective amount of the pharmaceutical composition of the present invention will depend on the age and overall physiological condition of the patient and the route of administration. In one embodiment, the therapeutic dose will generally be about 10-2000 mg / day, and preferably about 30-1500 mg / day. Other ranges may also be used, including, for example, 50-500 mg / day, 50-300 mg / day, and 100-200 mg / day.

[0090] Administration may be a single dose, multiple doses spaced to generate an immunogenic response, once daily, twice daily, or more frequently, and may be reduced during the maintenance phase of the disease or disorder, for example, to once every two or three days instead of daily or twice daily. The dose and frequency of administration will depend on clinical signs, as evidenced by maintenance of remission, in which at least one or more, preferably multiple, clinical signs of the acute phase are reduced or absent, as known to those of skill in the art. Those skilled in the art will recognize that certain factors, including, but not limited to, the severity of the disease or disorder, previous treatments, the subject's overall health and / or age, and other diseases present, may influence the dose and timing required to effectively treat a subject. Furthermore, treatment of a subject with a therapeutically effective amount of an immunogenic adjuvant-containing composition targeting a disease, disorder, or infectious agent may include a single treatment or, optionally, a series of treatments.

[0091] The pharmaceutical compositions may be included in a kit, container, pack, or dispenser together with instructions for administration.

[0092] The practice of the present invention will employ, unless otherwise indicated, conventional techniques of chemistry, molecular biology, microbiology, recombinant DNA, genetics, immunology, cell biology, cell culture, and transgenic biology, which are within the skill of the art. For example, see: Maniatis et al., 1982, Molecular Cloning (Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY); Sambrook et al., 1989, Molecular Cloning, 2nd Ed. (Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY); Sambrook and Russell, 2001, Molecular Cloning, 3rd Ed. (Cold Spring Harbor Laboratory Press, Cold Spring Harlow and Lane,1988, Antibodies, (Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY); Jakoby and Pastan,1979;Nucleic Acid Hybridization(BDHames&S.J.Higgins eds.1984);Transcription And Translation(BDHames&S.J.Higgins eds.1984);Culture Of Animal Cells(RIFreshney,Alan R.Liss,Inc.,1987);Immobilized Cells And Enzymes(IRL Press,1986);B.Perbal,A Practical Guide To Molecular Cloning(1984);the treatise,Methods In Enzymology(Academic Press,Inc.,N.Y.);Gene Transfer Vectors For Mammalian Cells(J.H.Miller and M.P.Calos eds.,1987,Cold Spring Harbor Laboratory);Methods In Enzymology,Vols.154 and 155(Wu et al.eds.),Immunochemical Methods In Cell And Molecular Biology(Mayer and Walker,eds.,Academic Press,London,1987);Handbook Of Experimental Immunology,Volumes I-IV(D.M.Weir and C.C.Blackwell,eds.,1986);Riott,Essential Immunology,6th Edition,Blackwell Scientific Publications,Oxford,1988;Hogan et al.,Manipulating the Mouse Embryo,(Cold Spring Harbor Laboratory Press,Cold Spring Harbor,N.Y.,1986);Westerfield,M.,The zebrafish book.A guide for the laboratory use of zebrafish(Danio rerio),(4th Ed.,Univ.of Oregon Press,Eugene,2000)。.

[0093] Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs. Although methods and materials similar or equivalent to those described herein can be used to practice or test the present invention, suitable methods and materials are described below. It should be understood and expected that those skilled in the art can make variations to the principles of the present invention disclosed herein, and such variations are intended to be included within the scope of the present invention.

[0094] Incorporation by Reference Each of the applications and patents cited herein, and each document or document cited in each of such applications and patents (including those in prosecution of each granted patent; "application cited documents"), and each of the PCT and foreign applications or patents corresponding to and / or claiming priority from any of such applications and patents, and each of the documents cited or referred to in each of the application cited documents, are expressly incorporated herein by reference. More generally, if a document or document is listed within this specification, either in the document list preceding the claims or in the specification itself, each of these documents or documents ("references herein"), and each document or document cited in each of the references herein (including all manufacturer specifications, instructions, etc.), are expressly incorporated herein by reference. In the case of conflict, the present specification, including definitions, will control. Additionally, the materials, methods, and examples are illustrative only and are not intended to be limiting. [Example]

[0095] Example 1: Materials and Methods Mouse strains and cell culture C57BL / 6J (Jax 000664), C57BL / 6NJ (Jax 005304), CD14 KO (Jax 003726), caspase-1 / -11 dKO mice (Jax 016621), TLR4 mutant (C3H / HeJ, Jax 000659), and wild-type control TLR4 mutant (C3H / HeSNJ, Jax 000661) were purchased from Jackson Labs. NLRP3 KO and ASC KO mice were kindly provided by Dr. T. Horng of the Harvard School of Public Health. Caspase-11 KO mice were kindly provided by Dr. Junying Yuan of the Harvard Medical School. Caspase-1 single KO mice were kindly provided by Thirumala-Devi Kanneganti of St. Jude's Hospital. DCs were differentiated from bone marrow in IMDM (Gibco), 10% B16-GM-CSF-derived supernatant, 2 μM 2-mercaptoethanol, and 10% FBS and used after 6 days of culture. Purity of DCs was assessed by flow cytometry and was typically greater than 90%. MΦs were differentiated from bone marrow in DMEM (Gibco), 30% L929 supernatant, and 10% FBS. Immortalized MΦs were cultured in DMEM supplemented with 10% L929 supernatant and 10% FBS. Splenic DCs were purified as previously described (Zanoni et al., 2012). Prior to stimulation, cultured cells were washed and replated in DMEM supplemented with 10% FBS at a concentration of 1 × 106 cells / ml in a final volume of 100 μl. For experiments using pan-caspase inhibitors, cells were treated with zVADfmk (20 μM) for 30 minutes before the addition of the inflammasome-activating stimulus. Cycloheximide (50 ng / ml) was added at the time of stimulus administration. DOTAP transfection was performed according to the manufacturer's instructions. Briefly, 375 ng of DOTAP was added to 5 μg of LPS or 10 μg of oxPAPC in a final volume of 10 μl of DMEM without FBS. After 30 minutes, the DOTAP / LPS complex or DOTAP / oxPAPC complex was added to the culture.The indicated concentrations of LPS and oxPAPC were transfected using FuGENE as previously described (Kayagaki, N. et al. (2013) Science 341, 1246-1249).

[0096] Gene expression analysis and ELISA RNA was isolated from cell cultures using a Qiashedder (Qiagen) and a GeneJET RNA purification kit (Life Technologies). Gene expression of purified RNA was analyzed using a TaqMan RNA-to-CT 1-step kit (Applied Biosystems) on a CFX384 real-time cycler (Bio-Rad). Probes specific for viperin (Mm00491265_m1), IFNb1 (Mm00439552_s1), IL6 (Mm00446190_m1), caspase-1 (Mm00438023_m1), caspase-11 (Mm00432307_m1), Nlrp3 (Mm00840904_m1), Asc (Mm00445747_g1), TBP (Mm00446971_m1), or GAPDH (Mm99999915_g1) were purchased from Life Technologies. ELISAs for IL-1β, IL-2, IL-17, IL-18, TNFα, and IFNγ were performed using Mouse Ready-SET-Go ELISA kits (eBioscience). To measure secreted cytokines, the supernatants were collected, clarified by centrifugation, and stored at -20°C. Cell-bound cytokines were measured as follows: the 96-well plates were centrifuged and the supernatants were discarded. 250 μl of PBS was added to each well. Cells were frozen and thawed twice at -80°C and then stored at -20°C for further analysis.

[0097] Antibodies and reagents E. coli LPS (serotype O55:B5-TLRgrade™) was purchased from Enzo. OxPAPC and Pam3CSK4 were purchased from Invivogen. Oxidized PAPE-N-biotin (biotin-oxPAPC) and oxPAPC enriched in PEIPC were produced as previously described (Springstead, JR et al., (2012) J Lipid Res 53, 1304-1315). KOdiA-PC and DMPC were from Cayman Chemical and Avanti Polar Lipids, respectively. The following antibodies were used: HA (Roche; 3F10), MyD88 (R&D; AF3109), actin (Sigma; 5441), ASC (Millipore, clone 2EI-7), caspase-11 (Biolegend, clone Cas11.17D9), caspase-3 (Santa Cruz, H-277), viperin (Biolegend), and phospho-Stat-1 (Cell Signaling, clone 58D6). IRAK4 antibody was a gift from Shizuo Akira (Osaka University). For flow cytometry-based assays, fluorophore-conjugated antibodies were used as follows: PE anti-TLR4 (Biolegend; clone Sa15-21), PE / Cy7 anti-TLR4 / MD2 (Biolegend; clone MTS510), FITC anti-CD14 (eBioscience; clone Sa2-8), and APC anti-CD14 (ebioscience; clone Sa-28). PE anti-MHC class II and APC anti-CD40 antibodies were from eBioscience. Annexin V and 7-AAD viability stain were purchased from BioLegend. Incomplete Freund's adjuvant (F5506) and cycloheximide (C1988) were purchased from Sigma. DOTAP was purchased from Roche. FuGENE 2000 was from Promega. Endotoxin-free OVA was purchased from Hyglos / Biovendor. Recombinant IFNβ was from R&D Systems, Inc. Pierce LDH Cytotoxicity Assay Kit was purchased from Life Technologies.

[0098] Protein purification and in vitro protein-lipid interactions For studies measuring direct binding of oxPAPC to caspase-11, protein and SPR analyses were performed as described (Shi, J., et al., (2014b) Nature). Briefly, full-length recombinant catalytic mutants caspase-11(C254A) and caspase-11ΔN59(C254A) were purified from P3 baculovirus-infected SF-21 insect cells cultured in Sf-900™ II SFM for 72 hours at 28°C. Cells were lysed with a lysis buffer containing 1% Triton X-100, 50 mM Tris-HCl (pH 7.6), 300 mM NaCl, 50 mM imidazole, and 5 mM 2-mercaptoethanol. His-tagged proteins were purified from the lysates using Ni-NTA beads (Qiagen). The protein was released from the beads with an elution buffer containing 50 mM Tris-HCl (pH 7.6), 250 mM imidazole, and 300 mM NaCl. The imidazole was removed by dialysis. The protein was further purified on a HiTrap Q column and a Superdex G200 column (GE Healthcare Life Sciences).

[0099] Surface plasmon resonance (SPR) analysis measured ligand binding kinetics using a BIAcore T100 SPR instrument (GE Healthcare). Assays were performed at 25°C in a buffer containing 150 mM NaCl, 3 mM EDTA, 50 mM HEPES (pH 7.5), and 0.005% Tween-20. A CM5 sensor chip was first activated with a 1:1 mixture of 0.1 M N-ethyl-N'-(3-diethylaminopropyl)-carbodiimide and 0.1 M N-hydroxysuccinimide at a flow rate of 10 μL / min for 7 min. Full-length catalytic mutants caspase-11(C254A) and caspase-11ΔN59(C254A) were diluted to a concentration of 20 μg / mL in 10 mM sodium acetate (pH 5.0) and stabilized at approximately 3100 and 3300 response units, respectively. Rabbit IgG protein (10 μg / ml) diluted with 10 mM sodium acetate was immobilized to 3400 response units and served as a negative control. 1 M ethanolamine (pH 8.5) was flowed through the CM5 chip to block all remaining protein binding sites for 7 minutes (flow rate 10 μL / min). Ligand was flowed through the flow cell and an adjacent control flow cell (activated and blocked in the same way as the target flow cell, but without immobilized protein) for 1 minute at a flow rate of 30 μL / min. The dissociation process was carried out for 2 minutes at a flow rate of 30 μL / min. Bound ligand was removed by washing with 20 mM NaOH for 20 seconds. KD values were calculated using BIAcore T100 evaluation software by curve fitting to a 1:1 Langmuir binding model (subtracting the control flow cell value).

[0100] For caspase-11 multimerization and enzyme activity assays, full-length murine caspase-11 was cloned into the pFastBac™ HTA vector (Invitrogen) incorporating a TEV-cleavable N-terminal 6xHis tag using the EcoRI and XhoI restriction sites. Protein was expressed using the Bac-to-Bac baculovirus-insect cell system. Forty-eight hours after infection, Sf9 cells expressing His-caspase-11 protein were harvested by centrifugation at 2,000 rpm for 20 minutes. The cell pellet was resuspended in lysis buffer containing 20 mM HEPES, 150 mM NaCl, 5 mM tris(2-carboxyethyl)phosphine (TCEP), 20 mM imidazole, and a protease inhibitor cocktail at pH 7.5 and homogenized by sonication. The cell lysate was clarified by ultracentrifugation at 42,000 rpm for 2 hours at 4°C. The supernatant containing the target protein was incubated with Ni-NTA resin (Qiagen) pre-equilibrated with lysis buffer for 1 hour at 4°C. After incubation, the resin-supernatant mixture was poured into a column, and the resin was washed with lysis buffer. The protein was eluted with lysis buffer supplemented with 500 mM imidazole and further purified by size-exclusion chromatography.

[0101] To measure the ability of oxPAPC to multimerize caspase-11, monomeric or multimeric fractions of His-caspase-11 were incubated with oxPAPC on ice for 2 h and then analyzed with Superdex 200 (10 / 300).

[0102] To characterize the binding of biotinylated oxPAPC to HA-tagged caspase-11 in cell lysates, 293T cells were transiently transfected with pcDNA vectors expressing the indicated caspase-11 alleles (WT, K19E, and 3K (K62E K63E K64E)) to which the HA epitope was C-terminally fused. Forty-eight hours after transfection, cells were harvested with chilled PBS and lysed in 1 ml of lysis buffer containing 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 10% glycerol, and 1% NP-40 supplemented with Complete protease inhibitors (Roche). Cells were lysed on ice for 30 min, and the whole-cell extract was collected into a new tube after centrifugation at 14,000 × g for 15 min in a cold-room benchtop centrifuge. A 100 μL aliquot of whole-cell lysate containing the indicated caspase-11 allele was removed and saved as input. The remaining 900 μL of lysate was aliquoted into three tubes, each containing 300 μL. 1 μg of biotinylated LPS and 10 μg of biotinylated oxPAPC were added to the first and second tubes, respectively. The third tube served as a mock control (to monitor the degree of nonspecific binding of the indicated caspase-11 allele to streptavidin beads). The biotinylated ligand and lysate were mixed and incubated on a nutator at 4°C for either 6 hours or overnight. Streptavidin beads (20 μL bed volume) were then added to all tubes (including the mock control, which was not treated with any biotinylated ligand) to capture the ligand-caspase-11 complex. This capture step continued for an additional 2–3 hours at 4°C. The beads were then washed three times with lysis buffer, and finally, 50 μL of SDS loading buffer was added. The protein complexes were further eluted by heating at 65°C for 15 min. 25 μL of the eluted protein complexes were separated by SDS-PAGE, and proteins retained by the biotinylated ligand were detected by Western analysis.

[0103] Capture of endogenous caspases by protein-lipid interactions The S100 fraction of iMΦs was prepared as follows. Confluent iBMDMs cultured in complete DMEM medium were harvested with ice-cold PBS containing EDTA (0.4 mM). Cells were then washed once with homogenization buffer (HB) (20 mM HEPES / KOH, pH 7.9, 250 mM sucrose, 0.5 mM EGTA) supplemented with complete protease inhibitor tablets (Roche). Cells were mechanically lysed by 20 compression strokes in a Wheaton™ Dounce Dura-Grind™ tissue grinder. The extent of cell lysis was monitored by trypan blue staining, confirming that >80% of cells were lysed. The crude lysate was then centrifuged at 800 × g for 10 minutes at 4°C to remove unbroken cells and nuclear components. The post-nuclear supernatant was collected and centrifuged at 13,000 × g for 10 min at 4 °C to remove large organelles and membranes. Finally, the clarified lysate was transferred to a Beckman polycarbonate ultracentrifuge tube (343778) and centrifuged at 100,000 × g for 1 h at 4 °C to remove residual membrane components. The resulting S100 supernatant (containing soluble cytosolic proteins) was either stored at -80 °C at a protein concentration of 2 mg / mL or used as a source of endogenous caspases for capture with biotinylated lipids. 1 mg of S100 supernatant was incubated with 15 μg of biotin-oxPAPC for 12–16 h at 4 °C on a nutator. Endogenous protein complexes bound to biotinylated oxPAPC (using 20 μL bed volume resin per reaction) were captured using streptavidin agarose resin (Pierre, P. et al., (1997) Nature 388, 787-792) on a nutator for 1-2 hours at 4°C. The captured protein complexes were then washed four times with detergent-containing wash buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 10% glycerol, 1% NP-40) and further eluted by incubation with 60 μL of SDS loading buffer at 65°C for 20 minutes. One-third of the eluate was resolved by SDS-PAGE, and endogenous caspases retained by biotin-oxPAPC were detected by Western blotting using the indicated antibodies.

[0104] Caspase-11 activity assay 5 μM His-caspase-11 with or without lipids (LPS, oxPAPC, and DMPC) was used for caspase activity assays in a reaction buffer containing 50 mM HEPES (pH 7.5); 10% (v / v) glycerol; 10 mM DTT; 1.0 mM EDTA; and 0.2% (w / v) BSA in Corning® 96-well, half-surface, black, flat-bottom microplates. The reaction was initiated by the addition of the substrate YEVD-AMC to a final concentration of 10 μM. Data were collected on a SpectraMax M5e multimode microplate reader (Molecular Devices) using excitation at 385 nm and emission at 460 nm with an automatic cutoff filter at 455 nm.

[0105] Flow cytometry iMΦ, primary bone marrow-derived MΦ, and DCs or splenic DCs (0.5 × 10 6) were treated with E. coli LPS, oxPAPC, or chemical inhibitors for the indicated times at 37°C. Cells were then washed with 1 mL of chilled PBS and stained with the appropriate antibodies on ice for 20–30 minutes. To reduce nonspecific antibody binding, 2% mouse or rat serum was used as a blocking reagent. Stained cells were then washed with 1 mL of chilled PBS and resuspended in 200 μL of PBS. Surface receptor staining was analyzed using a BD FACSCanto II. The mean fluorescence intensity (MFI) of CD14 and TLR4 from unstimulated or stimulated cells was recorded. The percentage of surface receptor staining (the ratio of the MFI measured in stimulated cells to the MFI measured in unstimulated cells) at the indicated time points was plotted to reflect the efficiency of receptor endocytosis. To measure the degree of TLR4 / MD-2 dimerization, the percentage of TLR4 / MD-2 dimers was calculated, with the percentage of TLR4 / MD-2 monomers set at 100%. The percentage of TLR4 / MD-2 monomer was determined by the ratio of the MFI value of stimulated cells (obtained by MTS510 antibody staining) to the MFI value of unstimulated cells. Cells were stained with anti-MHC class II or anti-CD40 antibodies.

[0106] Western blotting and midsome formation For Western blotting, iMΦ (5 × 10 6 ) were stimulated with ligand for the indicated time periods and then lysed in 700 μL of lysis buffer containing 1% NP-40, 50 mM Tris-HCl (pH 7.4), 150 mM NaCl. Protease and phosphatase inhibitors were added immediately before cell lysis. Immunoblotting was performed using standard molecular biology techniques.

[0107] For myosinosome formation, iMΦ (3 × 10 6) were stimulated with ligand for the indicated periods and then lysed in 700 μL of lysis buffer containing 1% NP-40, 50 mM Tris-HCl (pH 7.4), and 150 mM NaCl. Protease and phosphatase inhibitors were added immediately before cell lysis. The lysates were centrifuged at maximum speed in a tabletop centrifuge in a cold room (4°C) for 15 minutes. The clarified supernatant was collected, and 80 μL of the supernatant was saved as the total extract. 1 μg of anti-MyD88 antibody and 15 μL (bed volume) of Protein G Sepharose were added to the remaining supernatant, and incubation continued overnight at 4°C on a nutator. The beads were then washed three times with lysis buffer, and 60 μL of SDS loading buffer was added. Protein complexes were further eluted by heating at 65°C for 15 minutes. Aliquots (20 μL) of the eluted protein complexes were separated by SDS-PAGE and visualized by Western blotting using the indicated antibodies.

[0108] Immunofluorescence BMDC cells were treated with LPS (1 μg / mL) for 3 hours before inflammasome-inducing stimulus challenge. For viability or permeability experiments, MitotTacker CMX-ROS (Life Technologies) or Zombie Red (BioLegend) dye was used according to the manufacturer's instructions, followed by fixation with 4% paraformaldehyde. After a permeabilization step using 0.1% Triton X-100 in 0.2% BSA-PBS, cells were blocked with 2% BSA-PBS and incubated with rabbit anti-ASC pAb (AL177, Adipogen) and mouse anti-caspase-1 mAb (Caspase-1, Adipogen), followed by incubation with Alexa Fluor 488-conjugated chicken anti-rabbit IgG (Life Technologies) and Alexa Fluor 568-conjugated goat anti-mouse IgG (Life Technologies) diluted in blocking buffer. Nuclei were counterstained with DAPI (Life Technologies) or DRAQ5 (BioLegend). Images were obtained using a Zeiss Axiovert 200M confocal microscope or an Olympus BX41 fluorescence microscope.

[0109] PI permeabilization assay BMDCs or BMMs were seeded into black, clear-bottom 96-well tissue culture plates and treated with pre-stimulation for 3 hours. After gentle washing with PBS, 100 μl of pre-warmed staining solution (5 μM PI, 5% FBS, 20 mM HEPES, phenol red-free MgCl2 and CaCl2 in HBSS) was added to each well and incubated for 5 minutes at 37°C and 5% CO2. Immediately before measurement, 100 μL of staining solution containing 2× the inflammasome-inducing stimulus without PI was added to the appropriate wells. 0.1% Triton X-100 was used as a maximal permeability positive control. The increase in fluorescence intensity was continuously recorded over 3 hours at 37°C using a FLUOstar Omega microplate reader (BMG Labtech) with an excitation of 544 nm and an emission filter of 620-10 nm.

[0110] In vivo immunization and in vitro restimulation WT C57BL / 6NJ and caspase-1 / -11 dKO C57BL / 6NJ mice were immunized in the upper back (injected through each shoulder) with either 150 μg / mouse of endotoxin-free OVA and 7 μg / mouse of LPS emulsified in incomplete Freund's adjuvant, or 150 μg / mouse of endotoxin-free OVA, 65 μg / mouse of oxPAPC, and 7 μg / mouse of LPS emulsified in incomplete Freund's adjuvant. CD4+ T cells were isolated from draining lymph nodes 7 or 40 days after immunization by magnetic cell sorting using anti-CD4 beads (Miltenyi Biotech). Cells were seeded into 96-well plates at a concentration of 100,000 cells per well in the presence of 100,000 DCs and serial dilutions of OVA starting at 1 mg / ml. Secretion of IFNγ, IL-17, and IL-2 was measured by ELISA after 5 days.

[0111] Viral replication assay for HSV infection Mice were maintained in accordance with institutional and NIH animal experimentation guidelines, and all procedures were approved by the Institutional Animal Care and Use Committee of Harvard Medical School. The indicated mouse strains were anesthetized in an isoflurane chamber and then intraperitoneally injected with ketamine (3.7 mg / mouse) and xylazine hydrochloride (0.5 mg / mouse). Corneal scratches were made, and infections were performed as previously described (Cliffe, AR et al., (2009) Journal of Virology 83, 8182-8190). To measure viral amplification in the eyes, tear film swabs were collected using a sterile polyester applicator (Puritan) for the first 5 days after infection, and virus in tears from the eyes was titrated using Vero cells as previously described (Coen, DM et al., (1989) Proc Natl Acad Sci USA 86, 4736-4740).

[0112] statistical analysis Hypotheses were tested with a single pairwise comparison two-tailed t-test. p-values calculated in Excel (Microsoft Corporation) are coded with an asterisk: <0.05 ( * ), <0.01( ** ), <0.001( *** ).

[0113] Example 2. Identification of oxPAPC as a TLR4 antagonist and a CD14 agonist Oxidized phospholipids such as oxPAPC have a complex history and have been reported to act as both activators and inhibitors of inflammation. Some studies have shown that oxPAPC can inhibit LPS-induced TLR4-dependent proinflammatory cytokine expression in a concentration-dependent manner (Bochkov, V. et al., (2002) Nature 419, 77-81; Erridge, C. et al., (2008) The Journal of biological chemistry 283, 24748-24759; Oskolkova, O. V. et al. (2010) J Immunol 185, 7706-7712), while other studies have reported that oxPAPC is an activator of TLR4-dependent inflammatory responses (Imai, Y. et al. (2008) Cell 133, 235-249; Shirey, K. A. et al. (2013) Nature 497, 498-502).

[0114] To determine the activity of oxPAPC and PAPC, the ability of these lipids to bind to the LPS receptors TLR4 and CD14 was examined in immortalized murine bone marrow-derived macrophages (BMDMs; or iMΦ). Side-by-side comparisons of LPS- and oxPAPC-stimulated cells were performed to assess the ability of these molecules to induce the expression of known TLR4-dependent genes (Figures 1 and 2). Compared to LPS, which induced robust expression of the cytokines IL-1β and interferon beta (IFNβ) (Figure 2), and the IFN-stimulated gene viperin, oxPAPC failed to upregulate these genes (Figure 4B). It is possible that TLR4-dependent genes other than those assayed were activated by oxPAPC.

[0115] These studies evaluated several concentrations of oxPAPC, all of which were similar to those reported to be present in inflamed or injured tissues in vivo (Oskolkova, OV et al. (2010) J Immunol 185, 7706-7712). TLR4 dimerization was assessed by flow cytometry using an antibody that detects only TLR4 monomers. Dimerization was determined to be induced by LPS but not by oxPAPC treatment (Figure 43A). To complement these analyses, we also investigated the inducible interaction between the receptor-proximal protein MyD88 and IRAK4.

[0116] These proteins form a supramolecular organizing center (SMOC) called the midsome (Kagan, JC et al., (2014) Nat Rev Immunol 14, 821-826; Lin, SC et al., (2010) Nature 465, 885-890; Motshwene, PG et al., (2009) J Biol Chem 284, 25404-25411). Midsomes were assembled exclusively in response to TLR activation (Bonham et al., 2014). Therefore, detection of midsomes can be used as a general readout of TLR activation. LPS induced the formation of MyD88-IRAK4-containing midsomes within 30 min of treatment, whereas oxPAPC failed to induce any detectable binding between these proteins (Figure 43C and Figure 49A). Furthermore, cells treated with oxPAPC did not contain detectable amounts of phosphorylated STAT1 or the IFN-stimulated gene viperin (Figure 1, Figure 4D, and Figure 4B), both of which were abundant upon treatment with LPS. These data indicated that oxPAPC is not a mimic of LPS and had little or no ability to directly activate TLR4 in BMDMs.

[0117] In cell-free overexpression systems, oxPAPC acted as an inhibitor of TLR4 signaling events by competing with LPS for access to either CD14 or the LPS-binding protein MD-2 (Bochkov, V. et al., (2002) Nature 419, 77-81; Erridge, C. et al., (2008) The Journal of Biological Chemistry 283, 24748-24759). The LPS-binding protein MD-2 was involved in cross-linking and activation of TLR4. However, the ability of oxPAPC to bind to TLR4 regulators has been investigated primarily in cell-free systems or epithelial cells. The extent of oxPAPC-mediated inhibition was affected by varying the ratio of LPS and oxPAPC administration (Figures 1-5), indicating that these two factors likely compete for the same binding site as CD14. Consistent with competition for such a single binding site, mutant CD14 alleles that were unable to bind LPS were not endocytosed, even in the presence of oxPAPC or LPS.

[0118] To determine whether oxPAPC bound to CD14 in iMΦ or MD-2 in BMDM, we used several assays to monitor the inducible dimerization or endocytosis of candidate receptors by flow cytometry. As previously reported, LPS treatment caused endocytosis of CD14 and TLR4, resulting in the loss of surface staining for these proteins (Zanoni, I. et al., (2011) Cell 147, 868-880). Interestingly, oxPAPC was unable to induce endocytosis of TLR4 (Figure 4), but was able to promote rapid endocytosis of CD14 (Figures 3, 44A, and 50A). Thus, this rapid internalization of CD14 induced by oxPAPC created a depletion of CD14 at the cell surface. Without wishing to be bound by theory, we believe that this depletion of CD14 at the cell surface may explain the ability of this lipid to block TLR4 signaling. Indeed, oxPAPC-treated cells that were subsequently treated with LPS showed defects in TLR4 endocytosis and TLR4-induced gene expression.

[0119] CD14 surface enrichment was attributed to the antagonism of CD14 endocytosis and resynthesis (Tan, Y. et al., (2015). Immunity 43, 909-922), and was most clearly observed under conditions that prevented the latter. Consequently, the extent of oxPAPC- or LPS-induced CD14 endocytosis was enhanced under conditions in which protein synthesis was blocked with cycloheximide (Figure 50B). Cycloheximide treatment did not affect either TLR4 internalization or dimerization (Figures 50C and 50D). Primary bone marrow-derived MΦs and bone marrow-derived DCs behaved similarly to iMΦs in that oxPAPC promoted CD14 endocytosis but not TLR4 dimerization or endocytosis (Figure 44B). Thus, oxPAPC-induced endocytosis of CD14 (but not TLR4) created a lack of CD14 at the cell surface, likely explaining the ability of this lipid to block TLR4 signaling. Indeed, oxPAPC-treated cells subsequently treated with LPS exhibited defects in TLR4 dimerization, endocytosis, TNFα secretion, and STAT1 phosphorylation (Figures 44A and 44F), the latter two being classical readouts of TLR4 signaling.

[0120] To explore the possibility that CD14 interacts with PAMP (LPS) and DAMP (oxPAPC) using a similar mechanism, we investigated the amino acids in CD14 required for interaction with these lipids. The LPS-binding domain of CD14 was previously identified as a large hydrophobic pocket composed of four distinct regions in the primary amino acid sequence (Kim, J. et al., (2005) J Biol Chem 280, 11347-11351). CD14 alleles containing mutations in either one region (1R) or two regions (2R) retained the ability to form complexes with biotinylated LPS, whereas mutations in all four regions (4R) of CD14 abolished LPS-binding activity (Tan, Y. et al., (2015). Immunity 43, 909-922). Each of these mutant CD14 alleles encoded a full-length, folded protein that was transported to the cell surface (Tan, Y. et al., (2015). Immunity 43, 909-922). Notably, the 4R mutant was also defective in interacting with biotinylated oxPAPC (Figure 4E). Furthermore, when stably introduced into CD14 knockout (KO) iMΦ, the 4R mutant CD14 was not internalized in response to LPS or oxPAPC treatment (Figure 4F). Thus, these data indicate that the same amino acids within CD14 promoted the interaction with both the DAMP (oxPAPC) and the PAMP (LPS), providing molecular support for the conclusion that oxPAPC can be considered a selective LPS mimetic (i.e., for CD14-dependent activity). Overall, these data indicated that oxPAPC was an activator of CD14 but not TLR4. This ability to shedding CD14 and TLR4 endocytosis likely explains how oxPAPC functions as a TLR4 antagonist.

[0121] Example 3. oxPAPC promoted the activation of NLRP3 inflammasome in dendritic cells (DCs) Although the above examples demonstrated that oxPAPC was not an inflammatory activator, several studies have demonstrated the proinflammatory function of these lipids (Imai, Y. et al. (2008) Cell 133, 235-249; Shirey, K. A. et al. (2013) Nature 497, 498-502). Some DAMPs failed to induce a proinflammatory response in naive cells but were able to induce cytokine release from cells previously exposed to microbial products. For example, extracellular ATP has been reported to activate inflammasome-dependent IL-1β release from cells pre-stimulated with TLR ligands (Petrilli, V. et al. (2007) Current Opinion in Immunology 19, 615-622).

[0122] As shown in Figures 6 and 7, PAPC containing the oxPAPC component lipid KOdiA-PC (1-(palmitoyl)-2-(5-keto-6-octenedioyl)phosphatidylcholine) activated inflammasomes in DCs.

[0123] As shown in Figure 8, CD14 regulated inflammasome activation in response to PAPC. Although inflammasome activation in response to PAPC was CD14-specific, PAPC could also induce CD36 internalization (Figure 9). CD14 regulated PAPC-mediated inflammasome activation independently of type I IFN (Figure 10).

[0124] The regulation of caspase-1 and caspase-11 expression was similar in wt DCs and Cd14- / DCs (Fig. 11). PAPCs induced inflammasome activation in a cell type-specific manner (Fig. 12).

[0125] Other PAMPs were also identified that stimulated PAPC-induced inflammasome activation (FIGS. 13 and 14).

[0126] Notably, not all modified PCs induced inflammasome activation (Figure 15).

[0127] To determine whether oxPAPC had proinflammatory function in a context-dependent manner, we investigated IL-1β release from primary BMDMs or BMDCs pretreated with or without LPS. Consistent with previous observations (Petrilli, V. et al., (2007) Current Opinion in Immunology 19, 615-622), LPS pretreatment enabled ATP to induce IL-1β release from DCs in a dose-dependent manner (Figure 51A). Notably, oxPAPC exhibited similar activity, but not in a cell type-dependent manner. Interestingly, oxPAPC was also able to induce IL-1β secretion, but only in LPS-primed DCs (Figure 45A). Although oxPAPC did not induce IL-1β release from naive cells, LPS pretreatment of DCs enabled oxPAPC to promote IL-1β release in a dose-dependent manner (Figures 45A and 51B).

[0128] Without wishing to be bound by theory, IL-1β release is typically mediated by inflammasomes, cytoplasmic protein complexes that cause the processing and atypical secretion of IL-1 family members (Petrilli, V. et al., (2007) Current opinion in immunology 19, 615-622).

[0129] To determine whether the oxPAPC-mediated IL-1β release identified above was an inflammasome-dependent event, the activity of this lipid was examined in BMDCs derived from either caspase-1 / caspase-11 double-knockout (KO) mice or ASC-deficient mice (also known as Pycard). ASC is a common adaptor protein involved in inflammasome assembly (Martinon, F. et al., (2002) Molecular Cell 10, 417-426). oxPAPC (or ATP)-mediated IL-1β release was completely abolished in BMDCs lacking caspase-1 / -11 (Figure 18) or ASC (Figure 17, and Figures 45B-C). This observation provided conclusive genetic evidence demonstrating the requirement of the inflammasome for oxPAPC-induced cellular responses. Because NLRP3 is one of the most common upstream activators of the inflammasome (Ye, Z., and Ting, JP (2008) Current Opinion in Immunology 20, 3-9), we also investigated oxPAPC-mediated IL-1β release in NLRP3-deficient BMDCs. Because oxPAPC failed to induce IL-1β release from NLRP3-deficient BMDCs (Figure 16) or from NLRP3 KO DCs (Figure 45D), we identified oxPAPC-mediated IL-1β release as an NLRP3-dependent process. Furthermore, ATP-mediated IL-1β was NLRP3-dependent, as expected. Importantly, inflammasome regulators were not required for TNFα secretion (Figures 45B-45D), indicating that TLR4-induced gene expression occurred independently of inflammasome activation.

[0130] Commercially available (and natural) oxPAPC contains a mixture of various oxidized species. To determine whether alternative sources of oxPAPC exhibit similar activity, we used custom-made oxPAPC (Springstead et al., 2012) that was identified as being enriched in PEIPC (1-palmitoyl-2-(5,6 epoxyisoprostanoyl)-sn-glycero-3-phosphocholine), the most active component of oxPAPC. Control analysis of the two different oxPAPCs showed similar results (Figure 45A). This confirmed that oxPAPC induced IL-1β release in LPS-primed DCs, regardless of source. In contrast to the observed effects on IL-1β release, cell-bound IL-1β levels were similar compared to cells stimulated with LPS alone, LPS / oxPAPC, or LPS / ATP (Figure 45A, Figure 51B, and Figure 51C). This latter observation was consistent with the finding that oxPAPC could only act as an inhibitor of TLR4 signaling when cells were pretreated with this DAMP.

[0131] To determine the specificity of the effect of oxPAPC on inflammasome-mediated events (e.g., IL-1β release), we investigated the effect of this lipid on the release of TNFα, a classical TLR-dependent cytokine. oxPAPC neither promoted nor inhibited TNFα release from DCs (Figure 51D). In addition, when DCs were cotreated with LPS / ATP or LPS / oxPAPC (i.e., without prestimulation), IL-1β was released only by oxPAPC-treated DCs (Figure 51E). This indicated that these two DAMPs differ in their ability to regulate IL-1β secretion. When a different phosphocholine variant, 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), was used, this variant was unable to induce IL-1β release (Figure 51F). In contrast, 1-(palmitoyl)-2-(5-keto-6-octenedioyl)phosphatidylcholine, a purified component of oxPAPC, or KOdiA-PC, was able to induce IL-1β secretion (Figure 51F). In all cases, TNFα secretion was unaffected by phosphocholine treatment (Figure 51F). These data established the specific ability of oxPAPC to promote IL-1β release without affecting TLR4 signaling in LPS-primed DCs.

[0132] oxPAPC-induced inflammasome activation was caspase-11 dependent, whereas ATP-induced inflammasome activation was not (Figure 19). Indeed, PAPC-induced inflammasome activation after prestimulation with both LPS and Pam3 required caspase-11 (Figure 20). Therefore, a biotinylated form of PAPC was contemplated as a tool for studying caspase-11 activation.

[0133] We identified biotinylated PAPC as a potent inducer of CD14 internalization (Figure 21), but not TLR4 internalization (Figure 22), or IL-1β secretion (Figure 23). In vitro binding assays of biotinylated LPS, OxPac, and Pac to caspase-11 and MD-2 indicated that they formed complexes in a caspase-11-dependent manner (Figure 24). Indeed, in Bio-LPS pull-down assays, PAPC acted as a dose-dependent competitor (Figure 25). In these assays, biotin-LPS was used at 5 μg per pull-down assay, while PAPC was used at 5, 50, and 500 μg to compete with LPS binding to MD-2 and caspase-11, respectively. Competition was effective at a ratio of 1:100 (LPS:PAPC).

[0134] It is likely that oxPAPC and LPS bound to the same domain of CD14 (Figure 26). However, LPS treatment affected DC survival (Figure 27), and PAPC treatment of pre-stimulated DCs promoted DC survival (Figure 28). This pro-survival effect of PAPC was not CD14-dependent (Figure 29). Inflammasome activation is typically associated with the release of IL-1β and subsequent death of activated cells. The lack of oxPAPC killing BMDCs was a positive result that further promoted its use as an adjuvant.

[0135] As shown in Figure 30, only P2C and P3C supported DC survival. DCs pre-stimulated with P2C and P3C did not show an increase in their survival in response to PAPC treatment (Figure 31).

[0136] Inflammasomes were efficiently activated by priming and coadministration of PAPC, but not ATP (Figure 32). Priming and coadministration of PAPC did not alter NF-κB activation in wild-type DCs (Figure 33).

[0137] In the absence of CD14, oxPAPC acted as an antagonist of TLR4 signaling (FIG. 34).

[0138] Given that the oxPAPC effect was observed to be CD14 dependent, it was possible that CD14 acted as a chaperone to remove (“clear”) PAPC from the extracellular space.

[0139] Co-administration of LPS and oxPAPC affected TLR4 internalization (FIG. 35), CD14 internalization (FIG. 36), and partially affected TLR4 dimerization (FIG. 37).

[0140] Some of the above results are summarized as follows: (1) A specific modified PC (PAPC but not DMPC) induced inflammasome activation. (2) PAPC-dependent, but not ATP-dependent, inflammasome activation was cell type specific (DCs but not Macs). (3) Inflammasome activation by PAPC required CD14, but activation by ATP did not require CD14. (4) Inflammasome activation by PAPC required caspases, whereas activation by ATP did not require caspase-11. (5) ATP induced pyroptosis in DCs, but PAPC did not. (6) PAPC promoted DC survival in a CD14-independent manner. (7) PAPC, but not ATP, was identified to induce inflammasome activation when coadministered with priming.

[0141] Thus, PAPC has been identified as a potent, natural adjuvant capable of augmenting adaptive immune responses.

[0142] We also evaluated LPS and Rhodo LPS for inflammasome activation. As shown in Figure 38, Rhodo LPS was a potent inducer of inflammasome activation. LPS-induced inflammasome activation was Nlrp3-dependent (Figure 39), Asc-dependent (Figure 40), and Casp1 / 11-dependent (Figure 41). On the other hand, Rhodo LPS-induced inflammasome activation was CD14-independent (Figure 42). Thus, Rhodo LPS was also identified as a non-canonical inflammasome-activating lipid, but its effect appeared to be CD14-independent (which distinguishes the apparent mechanism of Rhodo LPS from that seen in oxPAPC).

[0143] Example 4: oxPAPC did not promote IL-1β release from macrophages All well-defined inflammasome activators tested promoted IL-1β release from MΦs. To investigate whether oxPAPC had this ability, we performed similar experiments to those described above with primary bone marrow-derived MΦs. Interestingly, oxPAPC failed to induce IL-1β release in MΦs under any of the conditions tested (Figure 45E), whereas ATP promoted efficient IL-1β release from these cells in a dose-dependent manner (Figure 51A). These data identified oxPAPC as a cell-type-specific activator of inflammasome activity.

[0144] To better understand how DCs specifically respond to oxPAPC, we assessed their response to the priming step of inflammasome activation. In contrast, DCs produced more TNFα than MΦs in response to LPS (Fig. 51D). These results indicated that DCs were better "primed" than MΦs. However, IFNγ-treated MΦs and DCs primed with IFNγ still failed to release IL-1β in response to oxPAPC (Fig. 51G). It is likely that the differential responsiveness of DCs and MΦs to oxPAPC emerges after a priming step occurs during inflammasome activation.

[0145] We hypothesized that DCs might have intrinsic factors that transport oxPAPC into the cytosol and subsequently activate inflammasome-mediated IL-1β release. This possibility was investigated by directly transfecting oxPAPC into the cytosol of MΦs. This method promoted IL-1β release from DCs pre-stimulated with the TLR2 ligand Pam3CSK, but the pre-stimulated MΦs still failed to induce such a response (Figure 45F). Cytoplasmic LPS transfection was used as a positive control (Figure 45F) (Hagar, JA et al., (2013) Science 341, 1250-1253; Kayagaki, N. et al. (2013) Science 341, 1246-1249). These findings indicated the presence of a factor(s) in the cytosol of MΦs (or DCs) that enables oxPAPC to activate the latter.

[0146] To understand inflammasome activity present in MΦs and DCs more generally, we investigated ATP, another inflammasome activator that promoted IL-1β release from both cell types (Figure 51A). Interestingly, DCs and MΦs died with similar kinetics in response to LPS+ATP treatment. However, these cells released very different amounts of IL-1β (Figure 55G), expressed very different levels of ASC (Figures 51H–51I), and did not release or express other components of the canonical or noncanonical inflammasome (Figure 51I). In MΦs, there was a perfect correlation between the extent of cell death and the extent of IL-1β release. This observation is consistent with dying cells releasing this cytokine (Figure 55G). In contrast, the greatest amount of IL-1β was released from DCs when observed death was minimal. This observation is consistent with live cells releasing this cytokine (Figure 55G). Together, these data highlight fundamental differences in inflammasome activity in MΦs and DCs and demonstrate that oxPAPC is an activator of the inflammasome that is specific to DCs.

[0147] Example 5: oxPAPC promoted IL-1β release via non-canonical inflammasomes independently of TLR4 Caspase-11 is a known protease that binds to cytoplasmic LPS and promotes non-canonical inflammasome assembly and IL-1β release (Hagar, J.A. et al., (2013) Science 341, 1250-1253; Kayagaki, N. et al., (2013) Science 341, 1246-1249; Shi, J. et al., (2014a) Nature 514, 187-192). Because oxPAPC can mimic LPS and activate CD14 endocytosis, we also evaluated its ability to activate caspase-11-dependent responses. Notably, oxPAPC-mediated IL-1β release was largely abolished in caspase-11 KO DCs (Figure 46A). As expected, ATP-mediated IL-1β release remained intact in caspase-11 KO cells (Figure 46A). In all cases, TNFα secretion was unaffected (Figure 46B). This difference between oxPAPC and ATP in caspase-11-dependent IL-1β release ruled out the possibility that the activity of oxPAPC was mediated by indirect release of ATP from cells.

[0148] To complement these functional analyses, microscopic examination of individual DCs revealed that both oxPAPC and ATP induced the formation of "specks" containing ASC and caspase-1 in LPS-pretreated DCs (Figure 46C). These experiments were performed using doses of ATP (1 mM) and oxPAPC (120 μM), which induced similar levels of IL-1β release (Figure 51C). The kinetics of speck formation in response to oxPAPC was slower than that of ATP, but similar amounts of cells formed specks (Figures 52A-52B). These structures formed only under conditions in which IL-1β was released and individual inflammasomes were recognized (Stutz, A. et al., (2013) Methods in molecular biology 1040, 91-101). Interestingly, caspase-11 was required for the formation of ASC / caspase-1-containing specks in response to oxPAPC, but not ATP (Figures 46C and 52B). Without wishing to be bound by theory, because this protein was required for non-inflammasome assembly, caspase-11 was likely required for oxPAPC-induced IL-1β release.

[0149] Consistent with the notion that oxPAPC did not require TLR4 to exert its function, the ability of oxPAPC to activate IL-1β release was independent of TLR4 signaling. Indeed, cells pre-stimulated with the TLR2 ligand Pam3CSK or the TLR9 ligand CpG elicited responses similar to those pre-stimulated with LPS (Figures 52C-52D). As observed with LPS-stimulated cells, IL-1β release from Pam3CSK-pre-stimulated DCs required NLRP3, ASC, and caspase-11 (Figure 52C). ATP-mediated IL-1β release after Pam3CSK pre-stimulation remained intact in caspase-11 KO cells, but not in caspase-1 / -11 dKO cells (Figure 52C). All DC genotypes were capable of comparable levels of TNFα secretion (Figure 52C). To further exclude any possible activity of oxPAPC against TLR4, we pre-stimulated C3H / HeJ DCs (naturally unresponsive to LPS due to a mutation in the TLR4 TIR domain) (Poltorak, A. et al. (1998) Science 282, 2085-2088) with Pam3CSK and measured IL-1β secretion in response to oxPAPC. The absence of functional TLR4 did not alter the ability of oxPAPC to induce IL-1β release (Figure 52E). These data further confirmed that oxPAPC was not required for TLR4-mediated signaling and that oxPAPC activated DCs upon contact with TLR ligands characteristic of either bacterial or viral infection. Thus, caspase-11 could be classified as a receptor that regulates immune responses to multiple types of pathogens, not just Gram-negative bacteria.

[0150] To further explore this possibility in an infectious setting, wild-type (WT) or caspase-11 KO mice were infected with herpes simplex virus type 1 (HSV-1). HSV-1 infection activates the NLPR3 inflammasome in ocular infection models (Gimenez, F. et al., (2015). Journal of Leukocyte Biology 2015 Oct 29. p.i.: jlb.3HI0715-321R). However, because this virus does not encode LPS, HSV-1 seemed like a good pathogen to investigate. Whether caspase-11 was involved in HSV-1 infection was previously unknown.

[0151] We found that caspase-11 KO mice were more susceptible to HSV-1 than WT mice at day 2 post-ocular infection. Indeed, increased infectious virus abundance was detected in eye swabs from caspase-11 mice compared to WT mice at this time point (Figure 5F). This difference in viral growth at day 2 was consistent with a previous study showing that NLRP3 KO mice produced higher viral titers at this time point (Gimenez, F. et al., (2015). Journal of Leukocyte Biology 2015 Oct 29. pii:jlb.3HI0715-321R). At subsequent time points, virus disappeared from the eyes of all mice examined, likely due to natural migration of the virus into the nervous system. These findings indicated that caspase-11 contributed to the protection of mice against nonbacterial pathogens. Without wishing to be bound by theory, the simplest model to explain these findings was that oxPAPC production at the site of infection (the eye) contributed to caspase-11 activation and subsequent restriction of viral replication. Direct testing of this model entailed the development of reagents that specifically ablate oxPAPC activity in vivo.

[0152] Example 6: Caspase-11 was identified as a receptor for oxPAPC oxPAPC has been shown to have the ability to activate caspase-11-dependent responses, indicating an interaction between these molecules. As previously described (Shi, J., et al., (2014b) Nature), endogenous caspase-11 can be isolated from cell lysates by its interaction with biotinylated LPS (Figure 46D). Interestingly, biotin-oxPAPC also formed a complex with endogenous caspase-11 (Figure 46D). In contrast, neither lipid captured endogenous caspase-3 (Figure 46D). To determine whether oxPAPC directly bound to caspase-11, in vitro protein-lipid interaction studies were performed. As shown in Figure 46E, oxPAPC exhibited a dose-dependent resonance signal with immobilized catalytically inactive caspase-11 (C254A) in surface plasmon resonance (SPR). In contrast, DMPC, which did not promote IL-1β release from DCs (Figure 46E), showed no detectable binding to caspase-11, and oxPAPC showed no binding to IgG by SPR (Figure 46E). The dissociation constant (Kd) between caspase-11 and oxPAPC was 1.3 × 10 -6 These SPR data showed that caspase-11 formed a complex with its own encoded lipid (oxPAPC) in addition to LPS and promoted IL-1β release in response to both.

[0153] Example 7: LPS and oxPAPC interact with caspase-11 through distinct domains and induce distinct activation mechanisms Because the same residues within CD14 were required for binding to LPS and oxPAPC, we investigated whether the LPS-binding CARD was required for interaction with oxPAPC. As expected (Shi et al., 2014b), mutation of specific lysine residues within the caspase-11 CARD prevented its interaction with LPS, as assessed by the ability of biotin-LPS to capture caspase-11 protein produced in 293T cells (Figure 52G). Interestingly, these lysine residues did not prevent its interaction with biotin-oxPAPC (Figure 52G). Furthermore, a mutant caspase-11 lacking its entire CARD and containing only its C-terminal catalytic domain retained the ability to form a complex with biotin-oxPAPC (Figure 52G). These results were verified by SPR analysis. The Kd for the interaction of oxPAPC with the catalytic domain of caspase-11 (designated ΔN59) was nearly identical to that calculated for the interaction with full-length caspase-11 (Figure 46E). As expected, LPS did not exhibit the ability to bind to the catalytic domain of caspase-11. Thus, these data established that, unlike CD14, separate domains within caspase-11 form contacts with LPS and oxPAPC.

[0154] In addition to complex formation with caspase-11, oxPAPC induced the multimerization of this protein, as demonstrated by gel filtration chromatography. As shown in Figure 46F, elution of caspase-11 monomer occurred at 15.03 mL, whereas caspase-11 contacted with oxPAPC eluted at an earlier volume, indicating an increase in the size of the protein complex. Caspase-11 dimers were predicted to elute at 13.82 mL, with higher-order multimers predicted to elute earlier. Therefore, the ability of oxPAPC to induce early elution of caspase-11 indicated that oxPAPC can induce the dimerization and / or multimerization of this protein. The extent of oxPAPC-induced caspase-11 multimerization was less than that reported for the same activity in response to LPS (Shi, J., et al., (2014b) Nature).

[0155] LPS-induced multimerization has previously been shown to promote the intrinsic protease activity of caspase-11 (Shi, J., et al., (2014b) Nature). Because LPS and oxPAPC multimerize caspase-11 through interactions with different domains, we investigated the enzymatic activity of caspase-11 in response to each of these lipids. As shown in Figure 52H, the intrinsic enzymatic activity of the caspase-11 monomer was low but increased upon contact with LPS or oxPAPC, identifying LPS as an even more robust activator.

[0156] Without wishing to be bound by theory, there are two possible explanations for the minimal ability of oxPAPC to activate caspase-11 enzymatic activity. First, the affinity of oxPAPC for caspase-11 and its ability to multimerize caspase-11 were weaker than those of LPS, resulting in minimal caspase-11 activation. In this respect, oxPAPC is merely a less potent version of LPS. However, the different mechanisms by which oxPAPC and LPS bind caspase-11 suggest that these lipids bind caspase-11 in fundamentally different ways, and that the interaction of oxPAPC with the catalytic domain is likely designed to block (rather than activate) enzymatic activity. The intrinsic enzymatic activity of pre-existing caspase-11 multimers was high (Figure 52H). This activity was further increased upon contact with LPS, but, notably, this activity was reduced upon contact with oxPAPC (Figure 52H). Furthermore, the ability of LPS to enhance the enzymatic activity of caspase-11 was blocked by oxPAPC in a dose-dependent manner (Figure 52I). These data supported the idea that two distinct biochemical interactions occurred between caspase-11 and proinflammatory lipids. Upon binding to the CARD of caspase-11, LPS induced potent multimerization and enzymatic activity. In contrast, oxPAPC bound to the catalytic domain of caspase-11, which promoted multimerization but limited enzymatic activity. Despite these two distinct interaction mechanisms, both LPS and oxPAPC assembled inflammasomes in DCs, and both promoted IL-1β release.

[0157] These findings raised the question of whether the catalytic activity of caspase-11 was required for the induction of IL-1β release by oxPAPC. To address this question, caspase-11-deficient DCs were reconstituted with a WT caspase-11 expression vector, a catalytic mutant (C254A) caspase-11 expression vector, or an empty vector (as a control). Cells expressing WT caspase-11 regained the ability to release IL-1β in response to either LPS or oxPAPC, whereas cells expressing mutant caspase-11 failed to release IL-1β in response to LPS (Figure 46G). Interestingly, DCs reconstituted with the mutant produced IL-1β in response to oxPAPC to the same extent as cells expressing WT caspase-11 (Figure 46G). TNFα release was used as a control (data not shown). These data establish that the requirement for caspase-11 activity for IL-1β release in response to LPS is distinct from oxPAPC.

[0158] We also assessed the ability of mutant and WT caspase-11 to induce pyroptosis, another function regulated by the non-canonical inflammasome. Caspase-11 enzymatic activity was required for transfected LPS to induce pyroptosis (Figure 46G), confirming that cells were correctly reconstituted. Surprisingly, no cell death was measured in response to oxPAPC (Figure 46G). Thus, these data supported the existence of two mechanisms for caspase-11-mediated IL-1β release, with only catalytic activity required for the response to LPS.

[0159] Example 8: CD14 captures oxPAPC and delivers it to caspase-11, promoting IL-1β release The above studies demonstrated that oxPAPC possessed the following two activities: 1) oxPAPC promoted CD14 endocytosis; and 2) oxPAPC promoted caspase-11-dependent non-canonical inflammasome activation. To determine the relationship between these activities, we investigated the requirement of CD14 for oxPAPC-induced IL-1β release. Cells were pre-stimulated with LPS for 3 hours, which was sufficient to allow repopulation of the plasma membrane with newly synthesized CD14 (Tan, Y. et al., (2015). Immunity 43, 909-922). Cells were stimulated with either ATP or oxPAPC. Interestingly, CD14 was required for oxPAPC-induced IL-1β release, as CD14 KO DCs did not release IL-1β in response to LPS / oxPAPC or Pam3CSK / oxPAPC treatment (Figure 47A). Secretion of IL-18, another cytokine released by inflammasome action, followed a similar pattern (Figure 53A). Similar results were obtained when we examined stimulated DCs initially isolated from the spleens of WT, CD14, or caspase-11 KO mice. These cells exhibited CD14- and caspase-11-dependent IL-1β release in response to LPS / oxPAPC, whereas TNFα secretion and upregulation of MHC-II and costimulatory molecules were unaffected by CD14 or caspase-11 deficiency (Figure 47B and Figure 53B). Furthermore, the responses of splenic DCs to LPS / ATP treatment were all unaffected by CD14 or caspase-11 deficiency (Figure 47B).

[0160] Without wishing to be bound by theory, two factors suggested that the requirement for CD14 for IL-1β release was not due to a defect in the priming step (i.e., TLR signaling). First, the dose of LPS used (1 μg / ml) bypassed the requirement for CD14 for TLR4-induced cytokine expression, as assessed by analysis of IL-1β transcripts and TNFα secretion (Figures 47A-C). Second, DCs primed with Pam3CSK also required CD14 for oxPAPC-induced IL-1β release, even when Pam3CSK primed the cells via TLR2 but not CD14 (Figures 47A and 47C).

[0161] In other experimental settings, type I IFN promoted caspase-11 expression and / or activation (Broz, P. et al., (2012). Nature 490, 288-291; Case, C. L. et al., (2013). Proc Natl Acad Sci USA 110, 1851-1856; Rathinam, V. A. et al., (2012) Cell 150, 606-619). Because CD14 promoted IFN expression in response to LPS treatment, as observed by its requirement for viperin expression, the role of type I IFN was investigated (Figure 47C). DCs treated with Pam3CSK / oxPAPC secreted IL-1β (Figure 47A) without inducing the expression of functional type I IFN, as indicated by the lack of viperin expression (Figure 47C). These data indicated that type I IFN was not required for regulating IL-1β secretion in response to oxPAPC stimulation. Furthermore, the defect in IL-1β secretion in CD14 KO cells could not be rescued by exposure to recombinant IFNβ (Figure 47D). Thus, IFN expression was neither necessary nor sufficient for oxPAPC-mediated caspase-11-dependent inflammasome activation in DCs. Furthermore, caspase-1, caspase-11, NLRP3, and ASC were all expressed at similar levels in stimulated and unstimulated WT and CD14 KO DCs (Figure 53C). This suggested that CD14 was not required for the expression of inflammasome regulators. Taken together, these data indicated that the requirement for CD14 for oxPAPC-mediated IL-1β release was not due to the need for cell pre-stimulation. Thus, CD14 could exert a direct role in promoting inflammasome-mediated IL-1β release.

[0162] To determine the means by which CD14 promoted inflammasome activation, we assessed the endocytosis-promoting activity of this LPS receptor. CD14 promoted oxPAPC endocytosis, transporting oxPAPC into cells and promoting IL-1β release. This bypassed the need for CD14 by delivering oxPAPC into cells via alternative means. The transfection reagent DOTAP has previously been shown to be a useful tool for delivering proinflammatory stimuli directly into endosomes and the cytosol (Honda, K., et al., (2005) Nature 434, 1035-1040). Therefore, WT and CD14 KO DCs were pre-stimulated with Pam3CSK and then exposed to DOTAP in complex with either LPS or oxPAPC. Consistent with previous results, LPS failed to induce IL-1β when administered in the extracellular medium, whereas DOTAP-mediated LPS delivery promoted IL-1β release from WT and CD14 KO DCs (Figure 47E). Interestingly, oxPAPC treatment produced similar results. Extracellular oxPAPC did not induce IL-1β release from pre-stimulated CD14 KO DCs, whereas oxPAPC complexed with DOTAP induced IL-1β release from CD14 KO cells in a caspase-dependent manner (Figure 47E). These data indicated that an alternative delivery mechanism could bypass the requirement for CD14 for oxPAPC-induced IL-1β release. Therefore, the primary function of CD14 in inflammasome activation was likely to be to deliver oxPAPC into cells.

[0163] These data suggested that CD14 functioned to deliver LPS to TLR4 on the cell surface and oxPAPC to caspase-11 in the cytosol via several endosomal intermediates. Because endolysosomes are highly degradative organelles, delivery of oxPAPC to endosomes might consume this lipid and limit its proinflammatory activity. Consistent with this idea, treatment of DCs with the acidification inhibitor chloroquine, which blocks endolysosomal activity, slightly enhanced IL-1β release from DCs (Figure 5D).

[0164] Furthermore, while LPS promoted IL-1β release from naive cells when delivered directly to the cytoplasm, only DOTAP enabled oxPAPC to induce IL-1β release from cells primed with TLR ligands. This difference in pre-stimulation dependency was likely due to the fact that LPS had the ability to prime cells via TLR4 and activate IL-1β release via caspase-11. In contrast, oxPAPC did not have the ability to prime cells directly and therefore was dependent on TLR stimulation. These data reinforced the idea that the principle of coincidence detection operates to govern two types of DC activation states. The first activation state is achieved when DCs encounter PAMPs, leading to classical TLR-dependent cytokine release via conventional protein secretion. The second, hyperactive state was achieved when DCs encountered DAMPs in the presence of PAMPs (i.e., coincident detection) or when virulent bacteria delivered LPS directly into the cytosol (Aachoui, Y., et al. (2013a). Science 339, 975-978; Casson, CN, et al. (2013). PLoS Pathog 9, e1003400; Hagar, JA et al. (2013) Science 341, 1250-1253).

[0165] Example 9: oxPAPC, unlike other inflammasome activators, did not kill cells In addition to promoting IL-β release, inflammasome activation is typically associated with the induction of cell death (Aachoui, Y., (2013b). Current opinion in microbiology 16, 319-326). Without wishing to be bound by theory, cell death by non-canonical inflammasomes is thought to depend on the enzymatic activity of caspase-11, which cleaves at least two proteins, gasdermin d and pannexin-1 (Kayagaki, N. et al., (2015) Nature 526, 666-671; Shi, J. et al., (2015) Nature 526, 660-665; Yang, D. et al., (2015) Immunity 43, 923-932). Because oxPAPC did not require the catalytic activity of caspase-11 to promote IL-11 release, we hypothesized that oxPAPC did not kill cells. To directly assess this possibility, we measured pyroptosis induction after oxPAPC administration or LPS transfection in LPS-primed DCs. Pyroptosis is characterized by a rapid loss of plasma membrane integrity, which should result in the release of cytoplasmic proteins (and organelles) from the cell body. We assessed membrane permeability of cell populations undergoing pyroptosis using LDH release in the supernatant. Cells treated with LPS / ATP began to release LDH 4 hours after treatment (Figure 48A). This combination was a well-documented activator of inflammasome-mediated cell death (Aachoui, Y., (2013b). Current opinion in microbiology 16, 319-326). LPS-transfected cells died at a later time point than ATP-treated cells, regardless of LPS prestimulation (Fig. 48A). Interestingly, all conditions that activated caspase-11 (e.g., LPS transfection or oxPAPC treatment) resulted in similar amounts of IL-11 in the supernatant (Fig. 48B), but only LPS transfection caused LDH release (Fig. 48A). These data indicated that oxPAPC promoted IL-11 release from viable cells.

[0166] To explain these observations, we developed a single-cell assay to investigate the viability of cells that contained assembled inflammasomes, as evidenced by the presence of ASC-containing aggregates. It was assumed that viable cells would be resistant to Zombie dye, a dye that labels the cytosol of cells with disrupted plasma membranes. Furthermore, cells with intact plasma membranes should retain functional organelles. In contrast, pyroptotic cells would have lost organelles and would stain strongly with Zombie dye. As shown in Figures 48C and 48D, cells treated with LPS / ATP contained ASC specks. These cells lost mitochondria and stained positive for Zombie dye. In striking contrast, cells treated with LPS / oxPAPC contained ASC specks but retained functional mitochondria and showed minimal staining with Zombie dye (Figures 48C-D). These observations, taken together, demonstrated that oxPAPC did not have the ability to kill cells and strongly suggested that oxPAPC induced IL-1β release from viable cells. Furthermore, not only did oxPAPC not induce pyroptosis, but this lipid also interfered with the LPS-activated delayed death pathway of DCs (Zanoni, I. et al. (2009) Nature 460, 264-268). In this experiment, the health of individual cells within the population was assessed up to 72 hours after treatment by flow cytometry using the viability dye 7-AAD, which detects genomic DNA in membrane-permeabilized cells (Paterson, AM et al. (2011) J Immunol 187, 1097-1105). Using a concentration of ATP (1 mM) that induced IL-1β release equivalent to that elicited by oxPAPC (Figure 48B), LPS / ATP treatment reduced DC viability immediately after treatment (Figure 54A). Notably, LPS treatment alone reduced cell viability over time (Figures 48E-F), whereas LPS / oxPAPC treatment actually increased the viability of the cell population (Figure 48F). These data indicated that oxPAPC treatment prevented LPS-induced DC apoptosis and promoted survival.

[0167] Example 10: oxPAPC was a potent adjuvant complement that promoted T cell-mediated adaptive immunity Although caspase-11 contributed to the control of acute viral infection (Figure 52F), the dual ability of oxPAPC to promote DC survival and IL-1β release suggested that oxPAPC may also promote DC-mediated adaptive immune responses. Indeed, IL-1β, a product of caspase-11 activation, has been characterized as possessing several activities that promote T cell activation, including conferring resistance to regulatory T cell suppression (Schenten, D. et al. (2014). Immunity 40, 78-90) (Sims, JE, and Smith, DE (2010) Nat Rev Immunol 10, 89-102). The oxPAPC / LPS mixture was investigated for its potential in vivo adjuvant activity.

[0168] To address this possibility, we subcutaneously injected WT, caspase-11, and caspase-1 / -11 dKO mice with LPS, ovalbumin (OVA), and / or oxPAPC emulsified in incomplete Freund's adjuvant (IFA). This route of inoculation was precisely what we used to establish the ability of TLR ligands to promote T cell differentiation (Pasare, C., and Medzhitov, R. (2004) Immunity 21, 733-741; Schnare, M. et al. (2001) Nat Immunol 2, 947-950). Forty days after injection, CD4+ T cells were isolated from draining lymph nodes and contacted ex vivo with DCs pulsed (or not) with OVA. T cell activation was then assessed by measuring IL-2, IL-17, and IFNγ levels by ELISA. Restimulation performed with DCs alone (without OVA) did not induce IL-2, IL-17, or IFNγ, indicating that cytokines released during restimulation were due to antigen-specific T cell responses (Figure 48G and Figure 54B).

[0169] Interestingly, T cells isolated from mice immunized with the LPS / oxPAPC mixture released significantly higher levels of IFNγ and IL-17 than T cells isolated from mice immunized with LPS (Figure 48G and Figure 54B). The ability of oxPAPC to enhance T cell activation was lost in caspase-11 or caspase-1 / -11 dKO mice (Figure 48G and Figure 54B). This observation was consistent with all in vitro data presented herein. Similar results were obtained when T cell activation was measured 7 days after immunization, during the effector phase of T cell activation (Figure 54C). Thus, oxPAPC had the ability to enhance LPS-mediated T cell activation in a caspase-11-dependent manner.

[0170] equivalent Those skilled in the art will recognize, or be able to ascertain using no more than routine experimentation, many equivalents to the specific embodiments of the invention described herein. As such, the following embodiments are is intended to be inclusive. [1] A composition for inducing an immune response to an immunogen, the composition comprising the immunogen and a non-canonical inflammasome-activating lipid. [2] The composition described in [1], wherein the non-canonical inflammasome-activating lipid is oxPAPC. [3] The composition described in [1] above, wherein the non-canonical inflammasome-activating lipid is PAPC. [4] The composition described in [1], wherein the non-canonical inflammasome-activating lipid is selected from the group consisting of one or more species of oxPAPC. [5] The composition described in [1], wherein the non-canonical inflammasome-activating lipid is selected from the group consisting of one or more of HOdiA-PC, KOdiA-PC, HOOA-PC, and KOOA-PC. [6] The composition described in [1] above, wherein the non-canonical inflammasome-activating lipid is Rhodo LPS. [7] The composition described in [1], wherein the non-standard inflammasome-activating lipid enhances the immune response to the immunogen when the composition is administered to a subject, compared to a composition lacking the non-standard inflammasome-activating lipid. [8] The composition according to [1], wherein the immunogen and lipid are present at concentrations sufficient to induce dendritic cell (DC) activation when the composition is administered to a subject. [9] The composition according to [1] above, wherein the composition does not induce a macrophage inflammatory response when administered to a subject.

[10] The composition according to [1] above, wherein the immunogen comprises an antigen selected from the group consisting of a human papillomavirus antigen, a herpes virus antigen such as a herpes simplex antigen or a varicella zoster antigen, a retrovirus antigen such as a human immunodeficiency virus type 1 antigen or a human immunodeficiency virus type 2 antigen, a hepatitis virus antigen, an influenza virus antigen, a rhinovirus antigen, a respiratory syncytial virus antigen, a cytomegalovirus antigen, an adenovirus antigen, a Mycoplasma pneumoniae antigen, an antigen of bacteria of the genus Salmonella, the genus Staphylococcus, the genus Streptococcus, the genus Enterococcus, the genus Clostridium, the genus Escherichia, the genus Klebsiella, the genus Vibrio, and the genus Mycobacterium, an amoeba antigen, a malaria parasite antigen, and a Trypanosoma cruzi antigen.

[11] The composition according to [1] above, wherein the composition is freeze-dried.

[12] The composition described in [1], wherein the composition consists essentially of the immunogen in combination with the non-canonical inflammasome-activating lipid.

[13] A pharmaceutical composition comprising the composition according to [1] above and a pharmaceutically acceptable carrier.

[14] The pharmaceutical composition according to

[13] above, wherein the carrier is an aqueous carrier.

[15] The pharmaceutical composition according to

[13] above, wherein the carrier is a solid carrier.

[16] A method for inducing an inflammatory response in dendritic cells of a subject, the method comprising administering to the subject the composition described in [1] above.

[17] A method for enhancing a subject's protective immune response to an immunogen, comprising administering to the subject the immunogen and a non-standard inflammasome-activating lipid in amounts effective to enhance the protective immune response, wherein the non-standard inflammasome-activating lipid is administered in an adjuvant-effective amount.

[18] The method of

[17] , wherein the immunogen and the non-canonical inflammasome-activating lipid are administered simultaneously to the subject.

[19] A method for inducing an immune response in a subject, comprising co-administering to said subject an immunogen and a non-canonical inflammasome-activating lipid in amounts effective to generate said immune response.

[20] The method according to any one of

[16] to

[19] above, wherein the subject is a human.

[21] The method according to any one of

[16] to

[20] above, wherein the immunogen and the non-canonical inflammasome-activating lipid are co-administered with a common pharmaceutical carrier.

[22] The method according to any one of

[16] to

[21] above, wherein the immunogen and the non-canonical inflammasome-activating lipid are administered parenterally.

[23] The method according to any one of

[17] to

[22] above, wherein the immune response is a preventive immune response.

[24] The method according to any one of

[17] to

[22] above, wherein the immune response is a therapeutic immune response.

[25] The method according to any one of

[17] to

[24] above, wherein the immune response comprises a humoral immune response.

Claims

1. A composition for inducing an immune response to an immunogen derived from an infectious agent, the composition comprising the immunogen together with a TLR ligand and oxPAPC; a composition for preventing and / or treating an infectious disease, wherein the composition is administered to a subject to induce or enhance an adaptive immune response in the subject against immunogens derived from an infectious agent; Here, the TLR ligand is a TLR4 ligand, a TLR2 ligand, and / or a TLR9 ligand, the TLR4 ligand is LPS or monophosphoryl lipid A (MPLA), the TLR2 ligand is Pam3CSK or Pam2CSK, and the TLR9 ligand is CpG.

2. The composition of claim 1 , wherein the TLR ligand is a TLR agonist.

3. The oxPAPC is HOdiA-PC (2-[[(2R)-2-[(E)-7-carboxy-5-hydroxyhept-6-enoyl]oxy-3-hexadecanoyloxypropoxy]-hydroxyphosph oryl]oxyethyl-trimethylazanium), KOdiA-PC ([(2R)-2-[(E)-7-carboxy-5-oxohept-6-enoyl]oxy-3-hexadecanoyloxypropyl] 2-(trimethylazaniumyl)ethyl 3. The composition according to claim 1, wherein the hydroxypropyl methyl acrylate (HOOA-PC) is selected from the group consisting of 1-palmitoyl-2-(5-hydroxy-8-oxo-octenoyl)-sn-glycero-3-phosphorylcholine, ...,6epoxyisoprostanoyl)-sn-glycero-3-phosphocholine, and 1-palmitoyl-2-(5,6epoxyisoprostanoyl)-sn-glycero-3-phosphocholine.

4. The composition of claim 3, wherein the oxPAPC is KOdiA-PC.

5. The composition of any one of claims 1 to 4, wherein the composition is administered in an amount effective to induce hyperactivity of dendritic cells in a subject.

6. The composition according to any one of claims 1 to 5, wherein the subject is a mammal.

7. The composition of any one of claims 1 to 6, wherein the subject is a human.

8. The composition of any one of claims 1 to 7, wherein the composition is administered as part of a pharmaceutical composition.

9. 9. The composition of any one of claims 1 to 8, wherein the composition is administered dermally, subcutaneously, intravenously, intramuscularly, parenterally, pulmonary, vaginally, rectally, nasally, or topically.

10. The composition of any one of claims 1 to 9, wherein the adaptive immune response is a preventative immune response.

11. The composition of any one of claims 1 to 9, wherein the adaptive immune response is a therapeutic immune response.

12. The composition of any one of claims 1 to 9, wherein the adaptive immune response comprises T cell activation.

13. The composition according to any one of claims 1 to 12, wherein the immunogen is selected from the group consisting of a viral antigen, a bacterial antigen, an amoebic antigen, and a protozoan antigen.

14. 14. The composition of claim 13, wherein the viral antigen is selected from the group consisting of a human papillomavirus antigen, a herpesvirus antigen, a retrovirus antigen, a hepatitis virus antigen, an influenza virus antigen, a rhinovirus antigen, a respiratory syncytial virus antigen, a cytomegalovirus antigen, and an adenovirus antigen.

15. 15. The composition of claim 14, wherein the herpes virus antigen is selected from the group consisting of a herpes simplex antigen and a varicella zoster antigen.

16. 15. The composition of claim 14, wherein the retroviral antigen is selected from the group consisting of a human immunodeficiency virus type 1 antigen and a human immunodeficiency virus type 2 antigen.

17. 14. The composition of claim 13, wherein the bacterial antigen is selected from the group consisting of a Mycoplasma pneumoniae antigen, a Salmonella antigen, a Staphylococcus antigen, a Streptococcus antigen, an Enterococcus antigen, a Clostridium antigen, an Escherichia antigen, a Klebsiella antigen, a Vibrio antigen, and a Mycobacterium antigen.

18. 14. The composition of claim 13, wherein the protozoan antigen is selected from a Plasmodium antigen and a Trypanosoma cruzi antigen.

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