ABSTRACT
The influence of cell death on adaptive immunity has been studied for decades. Despite these efforts, the intricacies of how various cell death pathways shape immune responses in the context of infection remain unclear, particularly with regard to more recently discovered pathways such as pyroptosis. The emergence of Listeria monocytogenes as a promising immunotherapeutic platform demands a thorough understanding of how cell death induced in the context of infection influences the generation of CD8+ T-cell-mediated immune responses. To begin to address this question, we designed strains of L. monocytogenes that robustly activate necrosis, apoptosis, or pyroptosis. We hypothesized that proinflammatory cell death such as necrosis would be proimmunogenic while apoptosis would be detrimental, as has previously been reported in the context of sterile cell death. Surprisingly, we found that the activation of any host cell death in the context of L. monocytogenes infection inhibited the generation of protective immunity and specifically the activation of antigen-specific CD8+ T cells. Importantly, the mechanism of attenuation was unique for each type of cell death, ranging from deficits in costimulation in the context of necrosis to a suboptimal inflammatory milieu in the case of pyroptosis. Our results suggest that cell death in the context of infection is different from sterile-environment-induced cell death and that inhibition of cell death or its downstream consequences is necessary for developing effective cell-mediated immune responses using L. monocytogenes-based immunotherapeutic platforms.
INTRODUCTION
Listeria monocytogenes is a Gram-positive, genetically tractable pathogen that stimulates a robust CD8+ T-cell response capable of breaking self-tolerance. These properties combine to make L. monocytogenes a promising cancer immunotherapeutic platform (1). The use of attenuated L. monocytogenes has done well in clinical trials (2); however, the mechanism by which L. monocytogenes stimulates robust cell-mediated immunity remains largely unknown. Throughout the course of infection, L. monocytogenes triggers a variety of innate immune responses, including Toll-like receptor (TLR) signaling and type I interferons (IFNs), that are hypothesized to be required for the development of a host protective immune response (reviewed in reference 3). Further, cross-priming from CD8α dendritic cells (DCs) is important to induce L. monocytogenes-stimulated immunity (4, 5). Dying cells are one major source of antigens for cross-presentation, and L. monocytogenes induces a variety of host cell death pathways, including apoptosis, necrosis, and pyroptosis, both directly in infected cells and in uninfected bystanders (6–9). In other live attenuated vaccine platforms, such as Mycobacterium bovis BCG, modulation of host cell death has been proposed as a means to increase the efficacy of this vaccine (10). However, how activation of cell death by L. monocytogenes influences the development of CD8+ T-cell responses and ultimately antitumor responses remains largely unknown.
Historically, the influence of cell death on the development of adaptive immune responses has been examined in a “sterile environment” through the coinjection of a bolus of dead cells with an antigen (11; reviewed in reference 12). Furthermore, cell death was primitively divided into two distinct forms: programmed, physiological apoptosis, which was considered anti-inflammatory and tolerogenic, and lytic necrosis, which was considered inflammatory and immunogenic (13). This dichotomy stems from the observation that the injection of necrotic cells results in the recruitment of innate immune cells and the release of danger-associated molecular patterns (DAMPs), such as HMGB1 (high-mobility group box 1), and actively induces dendritic cell (DC) costimulatory molecule expression (11, 14, 15). Conversely, apoptosis results in the production of anti-inflammatory cytokines, such as transforming growth factor beta (TGF-β) and interleukin-10 (IL-10) (15, 16), and fails to stimulate dendritic cell costimulatory molecule expression (11). Recently, however, these strict categories of apoptosis and necrosis have been blurred as other forms of cell death have been described, including immunogenic apoptosis (17, 18) and the programmed, inflammasome-dependent form of cell death known as pyroptosis (19). Pyroptosis has characteristics of apoptosis, including exposure of phosphatidylserine and fragmentation of DNA (20), and characteristics of necrosis, including cellular swelling, pore formation, and the release of HMGB1 (21, 22). How pyroptosis polarizes the immune response in a sterile context remains unclear; however, recent reports have begun to characterize the role of inflammasome in adaptive immune responses (23–26).
While decades of work have focused on understanding how cell death and its associated DAMPs influence adaptive immunity in a sterile environment, simultaneously decades of work focused on how pathogens, through their pathogen-associated molecular patterns (PAMPs), can induce innate immune responses that inform adaptive immune responses (27, 28). Though cell death and the release of DAMPs can be immunogenic in some cases, in the context of normal physiology it can also induce tissue repair enzymes (29) that ultimately limit activation of adaptive immune responses (30). Thus, when both DAMPs and PAMPs engage the innate immune system, as in the case of L. monocytogenes-induced cell death, it is unclear how these competing environments for tissue repair and formation of a protective immune response, respectively, synergize or antagonize each other to ultimately affect the adaptive immune response against the invading pathogen.
We established a unique set of L. monocytogenes strains that robustly induce host cell apoptosis, necrosis, or pyroptosis in myeloid-derived antigen-presenting cells to characterize how cell death influences L. monocytogenes-stimulated cell-mediated immunity. This system allows us to skew host cell death responses while maintaining similar infection conditions and pathogen-associated responses such as PAMP expression, subcellular localization of the pathogen, and antigen expression. Contrary to our hypothesis and the sterile cell death literature, we found that robust activation of necrosis, apoptosis, or pyroptosis negatively influences the generation of cell-mediated immunity, both at the level of antigen-specific T-cell activation and for long-term protective immunity. Using the 3-signal hypothesis of antigen presentation, costimulation, and inflammation (31) as a framework, our data suggest that each form of cell death negatively affects different signals required for optimal CD8+ T-cell generation, ranging from poor activation of costimulatory molecule expression to induction of a suboptimal inflammatory state. Overall, this work describes unique tools and provides critical insight into how different forms of cell death influence cell-mediated immunity in the context of L. monocytogenes infection, potentially illuminating novel strategies for the improvement of pathogen-based vaccines and immunotherapies.
RESULTS
Engineered strains of L. monocytogenes activate different cell death pathways.Many pathogens manipulate host cell death for their benefit, while hosts use cell death as a defense mechanism (32). Furthermore, modulation of activation of cell death in live attenuated vaccine platforms has recently been proposed as a mechanism to improve the efficacy of the BCG vaccine (10). To test the hypothesis that activation of cell death during infection influences the generation of adaptive immune responses, we engineered strains of L. monocytogenes to preferentially activate pyroptosis, necrosis, or apoptosis, all forms of cell death that L. monocytogenes induces throughout infection (Fig. 1A) (6–8). To understand how L. monocytogenes-induced cell death impacts its efficacy as an immunotherapy, we engineered all our strains, including the parent L. monocytogenes strain (strain Lm), in the attenuated ΔactA ΔinlB background that is currently being developed as a cancer immunotherapeutic platform (33). This platform attenuates L. monocytogenes by preventing cell-to-cell spread (ΔactA) and prevents c-met-mediated internalization by hepatocytes (ΔinlB), thus limiting internalization to primarily professional antigen-presenting cells such as macrophages and dendritic cells (33). Thus, we are able to study how preferential activation of cell death pathways in myeloid-derived antigen-presenting cells influences the development of immunity.
Characterization of L. monocytogenes strains that trigger cell death. (A) Unique strains of L. monocytogenes that hyperinduce the indicated form of cell death were developed. Bone marrow-derived macrophages (B, D) or bone marrow-derived dendritic cells derived from GM-CSF stimulation (C, E) were infected with the indicated strains of L. monocytogenes at an MOI of 2 and assayed for lactate dehydrogenase release (B, C) at 4 h postinfection or caspase 3/7 activation (D, E) at 2 h postinfection. Data shown are the means ± standard deviations (SD) from 3 independent experiments and are normalized to uninfected controls; ****, P < 0.0001.
To activate pyroptosis, we utilized a previously described strain of L. monocytogenes (Lm-pyro) that triggers the Nlrc4 inflammasome via the ectopic secretion of flagellin (34). Upon infection of wild-type bone marrow-derived macrophages (BMDMs) or granulocyte-macrophage colony-stimulating factor (GM-CSF)-stimulated cells (referred to as bone marrow-derived dendritic cells [BMDCs]), strain Lm-pyro triggered robust lytic host cell death as measured by lactate dehydrogenase (LDH) release (Fig. 1B and C). Importantly, the majority of lytic host cell death was dependent on the inflammasome, as caspase-1−/− BMDMs or BMDCs underwent only minimal lytic host cell death following infection with Lm-pyro (Fig. 1B and C). To induce necrosis, we utilized a previously described strain of L. monocytogenes (Lm-necro) that produces increased levels of cytotoxic listeriolysin O (LLO) (35). Infection of wild-type BMDMs or BMDCs with Lm-necro resulted in robust activation of lytic host cell death, similar to what we observed with Lm-pyro (Fig. 1B and C). Importantly, unlike what occurred with Lm-pyro, infection of caspase-1−/− BMDMs or BMDCs with Lm-necro also resulted in high levels of lytic host cell death (Fig. 1B and C), indicating that death induced by Lm-necro infection was not inflammasome dependent. Finally, to activate robust apoptosis, we engineered a strain of L. monocytogenes (Lm-apo) to ectopically secrete the proapoptotic BCL2 family member Bims to initiate the intrinsic apoptotic cascade (36). Infection of either wild-type or caspase-1−/− BMDMs or BMDCs with Lm-apo did not increase LDH release, indicating that this strain triggers minimal lytic host cell death, similar to what is seen with strain Lm infection (Fig. 1B and C). However, analysis of caspase 3/7 activation revealed increased activation of apoptosis following infection of wild-type BMDMs or BMDCs with Lm-apo relative to strain Lm, Lm-pyro, or Lm-necro (Fig. 1D and E). To verify our caspase 3/7 results, we performed annexin V staining on infected BMDMs. Infection with Lm-pyro or Lm-necro resulted in a significant population of annexin V+ PI+ cells suggestive of lytic death. In contrast, infection with Lm-apo resulted in a large annexin V+ PI− population that is indicative of early apoptosis, thus confirming our caspase 3/7 results (see Fig. S1A and B in the supplemental material). Importantly, consistent with the literature, the parent vaccine strain Lm induced each pathway of cell death to some degree and cell death increased over time (Fig. 1 and Fig. S1). Additionally, the kinetics and magnitude of activation of each pathway of cell death by the respective strains were similar, reaching peak activation of cell death by ∼4 h (Fig. S1). Taken together, these results suggest that L. monocytogenes can be engineered to preferentially trigger specific host cell death pathways and that L. monocytogenes likely does not possess intrinsic mechanisms to inhibit host cell death. Furthermore, these results suggest that we could utilize these strains to characterize how host modulation of cell death in primary infected antigen-presenting cells influences the generation of cell-mediated immunity in the context of L. monocytogenes immunization.
Activation of host cell death inhibits L. monocytogenes-stimulated protective immunity.To test the hypothesis that lytic, inflammatory death would be proimmunogenic, we immunized mice with a low dose (1 × 103 CFU) of Lm, Lm-necro, Lm-pyro, Lm-apo, or a phosphate-buffered saline (PBS) control. We immunized at a very low dose (0.00001 50% lethal dose [LD50]) to allow for the possibility that preferential activation of a specific pathway of cell death might induce more robust protective immunity than in the absence of robust cell death. Following lethal challenge, mice immunized with the standard vaccine platform Lm cleared the infection and harbored low burdens of bacteria in their spleens and livers (Fig. 2A and B). In contrast, mice immunized with Lm-pyro or Lm-apo had 1 to 1.4 log-higher bacterial burdens in their spleens and livers while mice immunized with Lm-necro had a 2.8-log increase in bacterial burdens in their spleens and a 1.8-log increase in bacterial burdens in their livers (Fig. 2A and B). This result is in stark contrast to the traditional view of the immune response to sterile cell death whereby lytic, inflammatory types of death, such as necrosis or pyroptosis, are proimmunogenic and suggests that L. monocytogenes-induced cell death, including proinflammatory cell death, is detrimental to the development of L. monocytogenes-stimulated immunity.
L. monocytogenes strains that induce cell death fail to induce protective immunity. Thirty days postimmunization with 1 × 103 CFU of the indicated strain, mice were challenged with 2 LD50 of virulent L. monocytogenes expressing B8R20–27 and full-length OVA. Spleens (A) and livers (B) were harvested and analyzed for CFU burden 68 to 72 h postchallenge. Data are representative of at least three independent experiments with 4 to 5 mice per group. *, P < 0.05 (determined by Mann-Whitney U-test).
Activation of host cell death inhibits antigen-specific T-cell activation.CD8+ T cells are essential for controlling L. monocytogenes infection and conferring long-term protective immunity (37). We hypothesized that deficits in antigen-specific CD8+ T cells at the primary, memory, and/or recall stages were responsible for the observed deficits in protection. To test the hypothesis that cell death resulted in defective T-cell priming, we immunized mice with strains of L. monocytogenes that also secrete the model CD8+ T-cell antigens B8R and ovalbumin (OVA) and assessed activation of antigen-specific T cells during the primary response. Western blotting for the OVA-B8R fusion protein demonstrated that all strains expressed similar levels of antigen (see Fig. S2A in the supplemental material). At the peak of the primary immune response, mice immunized with Lm mounted a robust B8R-specifc CD8+ T-cell response as indicated by the percentage (Fig. 3A) and total number (Fig. 3B) of CD8+ T cells producing IFN-γ. In contrast, mice that were immunized with each of the cell death-inducing L. monocytogenes strains demonstrated impaired B8R-specific CD8+ T-cell responses (Fig. 3A and B). We next examined multifunctional T cells capable of producing IFN-γ, tumor necrosis factor alpha (TNF-α), and IL-2, which have recently been suggested to be the true effector cells of the immune response as well as being the cells that are capable of forming memory cells (38–40). As we saw with single positive IFN-γ-positive T cells, mice immunized with cell death-activating strains of L. monocytogenes activated fewer effector T cells than mice immunized with Lm. Finally, analysis of OVA-specific T-cell responses following immunization with Lm-pyro, Lm-apo, or Lm-necro resulted in similarly impaired multifunctional OVA-specific CD8+ T-cell responses, both in relative percentage and in total number (Fig. S2B and C). Taken together, these data suggest that increased L. monocytogenes-induced activation of necrosis, pyroptosis, or apoptosis during T-cell priming inhibits optimal primary CD8+ T-cell responses to multiple different antigens.
CD8+ T-cell responses are impaired following immunization with cell death-inducing strains of L. monocytogenes. Mice were immunized with 1 × 103 CFU of the indicated strain, and splenocytes were examined by ex vivo B8R peptide stimulation 7 days postimmunization. Percentage (A) and total number (B) of IFN-γ+ B8R-specific CD8+ T cells. Percentage (C) and total number (D) of multifunctional B8R-specific CD8+ T cells expressing IFN-γ, TNF-α, and IL-2. Splenocytes were first gated for a single-cell population, followed by expression of CD8α. CD8α+ cells were then selected for expression of IFN-γ, followed by TNF-α and IL-2. Data are representative of at least 2 independent experiments with 5 mice per group; ***, P < 0.001; ****, P < 0.0001.
Deficits in T-cell responses during the primary response could result in deficits in memory T cells, potentially explaining the loss of protective immunity. To test this hypothesis, we examined B8R-specific, CD44high CD8+ T cells present 30 days postimmunization using B8R-specific tetramers. Analysis of B8R-specific memory responses demonstrated similar contraction rates among all strains (95 to 98%), resulting in significantly impaired B8R-specific memory T-cell populations following immunization with Lm-pyro or Lm-necro, relative to immunization with Lm, when measured either by frequency or by total number (Fig. S2D and E). Taken together, these data suggest that deficits in primary T-cell responses are maintained in the memory T-cell pool but that cell death during T-cell priming does not have a specific effect on T-cell contraction.
Finally, smaller pools of memory T cells could result in less robust recall responses following challenge. To test this hypothesis, we examined the recall T-cell population by analyzing CD8+ T cells after immunization and subsequent lethal challenge 30 days postimmunization. At 72 h postchallenge, when control of secondary infections is dependent on the primary immunizing strain (Fig. 2), there were significantly fewer total CD8+ T cells present in mice immunized with cell death-inducing L. monocytogenes (Fig. S2F), suggesting a deficit in CD8+ T-cell recall. Additionally, the frequency and, more significantly, the total number of IFN-γ+ (Fig. 4A and B) and multifunctional B8R-specific (Fig. 4C and D) CD8+ T cells were reduced in mice immunized with strains that triggered cell death. Fewer total multifunctional OVA-specific CD8+ T cells were also present following immunization with Lm-pyro, Lm-necro, or Lm-apo, again demonstrating that the deficits were not specific to the B8R antigen (Fig. S2G and H). Taken together, these results suggest that deficits in formation of primary CD8+ T cells persist through memory and recall populations and correlate with the lack of protective response (Fig. 2), suggesting that impairments in the generation of primary antigen-specific effector CD8+ T-cell responses are responsible for the deficits in protection.
Recall CD8+ T-cell responses are impaired following lethal challenge with L. monocytogenes. Thirty days postimmunization with 1 × 103 CFU of the indicated strain of L. monocytogenes, mice were challenged with a lethal dose of virulent L. monocytogenes expressing B8R20–27 and full-length OVA. Percentage (A) and total number (B) of IFN-γ+ B8R-specific CD8+ T cells at 72 h postchallenge, as examined by ex vivo peptide stimulation. Percentage (C) and total number (D) of multifunctional B8R-specific CD8+ T cells expressing IFN-γ, TNF-α, and IL-2. Splenocytes were first gated for a single-cell population, followed by expression of CD8+. CD8+ cells were then selected for expression of IFN-γ, followed by TNF-α and IL-2. Data are representative of at least 2 independent experiments with 5 mice per group. **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.
Induction of cell death does not impair antigen presentation.The three-signal hypothesis for T-cell generation states that antigen must be presented in the appropriate major histocompatibility complex (MHC) molecule and recognized by the T-cell receptor, costimulatory molecules must be present on both the antigen-presenting cell and the T cell, and finally, the appropriate inflammatory milieu must be present (31). Although activation of various cell death pathways during infection could impact each of these steps of T-cell activation, the most obvious hypothesis was that activation of cell death altered bacterial burdens and thus antigen presentation. Our use of the ΔactA ΔinlB background has previously been demonstrated to at least partially normalize differences in virulence and rates of clearance (34). However, when we assessed the impact of cell death on the levels of bacterial burdens in vivo, we observed differences in burdens of viable CFU within the first 48 h postinfection (see Fig. S3A and B in the supplemental material).
Previous reports suggest that the magnitude and kinetics of CD8+ T-cell expansion are determined within the first 24 h following L. monocytogenes infection and that expansion of antigen-specific T cells is independent of the duration and severity of infection (41). Additionally, previous work examining antigen presentation patterns following L. monocytogenes infection suggests that the amount of antigen present does not directly correlate with bacterial burden (42). Therefore, to assess whether or not our differences in bacterial burdens affected antigen presentation, and ultimately T-cell priming, we assessed antigen presentation using the B3Z T-cell hybridoma reporter cell line. The B3Z line acts independently of costimulation and activates expression of β-galactosidase upon recognition of the SIINFEKL epitope from ovalbumin (OVA257–264) (43). Consistent with analysis of antigen expression (Fig. S2A), analysis of SIINFEKL expression on CD11c+ DCs purified from the spleen 24 and 48 h postimmunization revealed similar levels of antigen presentation, independent of the induction of host cell death (Fig. 5A and B). These data, consistent with the previous reports that cross presentation is critical for L. monocytogenes-triggered immunity, suggest that L. monocytogenes-induced cell death does not lead to impairments in antigen presentation.
L. monocytogenes-stimulated cell death does not impair antigen presentation. (A, B) Antigen presentation of the SIINFEKL peptide was assessed using the B3Z hybridoma cell line at 24 (A) or 48 (B) hours postinfection. (C to F) To normalize burdens, mice were immunized with 1 dose of 1 × 103 CFU Lm, 1 dose of 1 × 105 CFU Lm-pyro or Lm-apo, or 2 doses of 1 × 107 CFU Lm-necro dosed 12 h apart. Bacterial burdens in spleens were examined at 24 (C) or 48 (D) hours following the first immunization. Seven days postimmunization, the percentage (E) and total number (F) of multifunctional B8R-specific CD8+ T cells expressing IFN-γ, TNF-α, and IL-2 were examined. Splenocytes were first gated for a single-cell population, followed by expression of CD8α. CD8α+ cells were then selected for expression of IFN-γ, followed by TNF-α and IL-2 (E, F). Data shown are the averages ± SD and are representative of at least 2 independent experiments with 3 mice per group (A to D) or 5 mice per group (E, F). ns, not significant; *, P < 0.05; **, P < 0.01.
Despite determining that decreased burdens of viable bacteria did not correlate with decreased antigen presentation, we developed a dosing scheme that normalized burdens among all strains of L. monocytogenes within the first 48 h postinfection (Fig. 5C and D). By tetramer analysis or production of single-producing IFN-γ cells, we observed impaired numbers of B8R-specific CD8+ T cells following immunization with Lm-pyro or Lm-necro (Fig. S3C and D). Importantly, using this modified dosing scheme, evaluation of the multifunctional status demonstrated impaired primary CD8+ T-cell responses following immunization with any cell death-inducing strain of L. monocytogenes (Fig. 5E and F). The impairments observed in the multifunctional state mirrored the previously observed primary T-cell responses when in vivo burdens were not normalized (Fig. 3), suggesting that differences in bacterial burden alone are unlikely to explain reductions in immunity. Similarly, immunization with normalized bacterial burdens still resulted in significant impairments in protective immunity, though to a lesser extent (Fig. S3E and F), suggesting that while differences in burden may minimally contribute to differences in protective immunity (Fig. 2), cell death inhibits the development of immunity independent of bacterial burden. Taken together, these data suggest that activation of cell death by L. monocytogenes does not lead to deficits in antigen presentation/abundance and instead suggest that defects in costimulation or appropriate inflammation are more likely the cause of impaired T-cell priming associated with L. monocytogenes cell death.
Cell death inhibits expression of dendritic cell costimulatory molecule expression.As differential activation of cell death did not significantly influence antigen presentation, we hypothesized that differences in dendritic cell (DC) costimulatory molecule expression may be responsible for the loss of protective immunity observed following L. monocytogenes-induced cell death. We first examined the total number of CD11c+ cells present following immunization with normalized bacterial burdens. Consistent with a lack of difference in antigen presentation, we found no statistically significant difference in the total number of CD11c+ cells within the first 24 or 48 h of immunization, suggesting that, on a global scale, DCs are not being killed by our cell death strains of L. monocytogenes (Fig. 6A).
Dendritic cell costimulation is impaired following induction of cell death. Mice were immunized with the normalized burden-dosing scheme, and spleens were harvested at the indicated time points and analyzed for total numbers that were CD11c+ (A), CD11c+ CD86+ (B), CD8α+ CD11c+ (C), or CD8α+ CD86+ CD11c+ (D). Splenocytes were first gated for a single-cell population followed by gating on CD11c+. CD11c+ cells were then examined for expression of either CD86 or CD8α (B, C). CD11c+ CD8α+ cells were gated for expression of CD86 (D). Data are the combination of results from 2 independent experiments with 3 mice per group. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.
Although the number of DCs was relatively comparable between immunizations, their activation state varied depending on the type of cell death. Immunization with Lm-pyro or Lm-necro resulted in a significant upregulation of the costimulatory molecule CD86+ on DCs above control Lm levels at 24 h postimmunization both in the number of cells (Fig. 6B) and the intensity per cell (see Fig. S4A in the supplemental material), consistent with the inflammatory nature of these cell death pathways. By 48 h postimmunization, mice immunized with Lm-pyro still had significantly more mature DCs (Fig. 6B; Fig. S4A). Conversely, Lm-necro-immunized mice demonstrated a sharp decrease in CD86+ expression at 48 h compared to that of 24 h, reaching levels that were below those of any of the other strains (Fig. 6B), although still above those of unimmunized mice. Finally, immunization with Lm-apo followed the course of Lm immunization throughout the first 48 h postimmunization.
CD8α+ DCs are known to be important for cross-presentation and are essential for effective T-cell responses to L. monocytogenes (4). Immunization with any of our strains did not influence the total number of CD8α+ DCs at 24 h postimmunization; however, immunization with Lm-necro resulted in a steep decline in the number of CD8α+ DCs at 48 h postimmunization (Fig. 6C), potentially suggesting that the loss of this important subset of DCs is responsible for poor T-cell priming following immunization with Lm-necro. Furthermore, the patterns of CD86 expression observed on total DCs were mirrored on CD8α+ DC. Notably, immunization with Lm-pyro resulted in enhanced CD86+ expression over Lm at 24 h postimmunization both in the number of cells and in the intensity per cell, while immunization with Lm-necro resulted in significantly impaired costimulatory molecule expression at 48 h postimmunization (Fig. 6D; Fig. S4B). Taken together, these results suggest that differential induction of host cell death during L. monocytogenes infection modulates the expression of costimulatory molecules on both antigen-presenting CD11c+ cells and cross-presenting CD8α CD11c+ cells. Specifically, induction of pyroptosis results in a lasting robust activation of costimulatory molecule expression, while necrosis results in a robust activation of costimulatory molecule expression that quickly decreases. This premature loss of costimulatory molecule expression, coupled with the loss of critically important cross-presenting DCs, could partially explain the deficits in antigen-specific T-cell activation in the context of necrosis.
Cell death-specific inflammation independently inhibits CD8+ T-cell generation.Defects or hyperactivation of costimulatory molecule expression could be due to differences in the inflammatory environment caused by the activation of cell death pathways by L. monocytogenes. Furthermore, proper “Signal 3” cytokines, such as IL-12 or type I IFN, are required for optimal CD8+ T-cell expansion and the formation of memory cells (44). Given the importance of inflammation in generating optimal CD8+ T-cell responses and the known influence of cell death on the inflammatory milieu, we next hypothesized that cell death-specific inflammation may independently influence antigen-specific CD8+ T-cell generation. To test this hypothesis, we analyzed the presence of inflammatory cytokines, including IFN-γ, IL-12p70, TNF-α, monocyte chemoattractant protein 1 (MCP1), IL-6, and IL-10 following immunization according to the normalized-load dosing scheme. We observed significant increases in the inflammatory cytokines IFN-γ, TNF-α, MCP1, and IL-6 under one or both of the inflammatory cell death conditions, i.e., infection with Lm-pyro or Lm-necro, at 24 h postinfection above cytokine levels in Lm-infected mice (Fig. 7A to E), consistent with these being inflammatory cell death pathways. Notably, we observed substantial increases in IFN-γ (approximately 1,000-fold relative to Lm) 24 h postimmunization with Lm-pyro (Fig. 7A). With the exception of IL-6 in Lm-necro-treated mice (Fig. 7E), very few of these inflammatory changes persisted through 48 h, suggesting that immunizations with Lm-pyro and Lm-necro result in early, robust inflammatory cytokine production that quickly tapers off. Levels of the anti-inflammatory cytokine IL-10 were at or near the limit of detection for all strains both at 24 and at 48 h postinfection (Fig. 7F).
Cell death-induced inflammation influences T-cell responses. Mice were immunized with the normalized burden-dosing scheme, and serum was examined at the indicated time point by cytokine bead array for levels of IFN-γ (A), IL-12p70 (B), TNF-α (C), MCP-1 (D), IL-6 (E), or IL-10 (F). Mice were immunized with 5 × 105 LPS-matured DCs from wild-type mice loaded with B8R20–27; 24 h after immunization, mice were boosted with 1 × 103 CFU of the indicated strain lacking B8R (G, H). Percentages (G) and total numbers (H) of IFN-γ-expressing CD8+ cells specific for B8R. Splenocytes were first gated for a single-cell population, followed by expression of CD8+. CD8+ cells were then selected for expression of IFN-γ. Data are the combination of results from 2 independent experiments with 3 mice per group (A to F), or data are representative of 2 independent experiments with 5 mice per group (G, H). Significance in comparison to Lm: *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001 compared to Lm, Significance in comparison to PBS: #, P < 0.05; ###, P < 0.001; ####, P < 0.0001.
To understand how these changes in inflammation impact the generation of antigen-specific T cells, we utilized a dendritic cell immunization and L. monocytogenes inflammatory boost strategy to separate antigen presentation and costimulation from the inflammatory stimulus provided by L. monocytogenes infection and its associated cell death. In this model, lipopolysaccharide (LPS)-matured dendritic cells were harvested from the spleens of mice previously implanted with Flt3-expressing B16 melanoma cells and were subsequently loaded with B8R peptides. B8R-loaded mature dendritic cells were adoptively transferred into naive mice, ensuring that all mice received equal amounts of antigen with proper costimulation. One day after B8R-dendritic cell immunization, mice were boosted with Lm, cell death-inducing strains of L. monocytogenes, or a PBS control. Importantly, these strains did not express the B8R antigen and provided only an inflammatory boost. This model has previously demonstrated that the inflammation provided by an L. monocytogenes infection significantly enhances the CD8+ T-cell response (45). Compared to mice boosted with PBS, mice boosted with Lm, Lm-necro, and Lm-apo had enhanced CD8+ T-cell responses as indicated by the percentages and total numbers of CD8+ T cells producing IFN-γ (Fig. 7G and H). Although boost with Lm-necro and Lm-apo provided improvement over a PBS boost in mice, this response did not reach Lm-stimulated levels, suggesting that the inflammatory milieu associated with these cell death pathways during infection is not as robust for T-cell activation as in their absence. Strikingly, boost with Lm-pyro failed to increase antigen-specific CD8+ T-cell responses beyond those in mice that received a PBS boost, and the level of the Lm-pyro boost responses even had a strong trend to be worse than the level of the PBS boost response (Fig. 7G and H). Importantly, differences in the ability of different L. monocytogenes strains to boost immunity were independent of any direct effect on DC viability, as rates of clearance of the adoptively transferred DCs were not affected by the secondary boost (Fig. S4C). These results suggest that the inflammatory milieu associated with each type of cell death is unique and independently influences the generation of L. monocytogenes-stimulated immunity. Importantly, the inflammatory milieu associated with L. monocytogenes-induced pyroptosis is suboptimal for generating cell-mediated immunity and likely contributes to the observed deficits in protective immunity following immunization with Lm-pyro.
DISCUSSION
The molecular mechanisms by which L. monocytogenes stimulates a robust CD8+ T-cell response capable of breaking self-tolerance remain largely unknown. As such, whether or not improvement of L. monocytogenes as an immunotherapeutic platform is possible is similarly unclear. We hypothesized that robust inflammation from lytic forms of cell death may improve L. monocytogenes-stimulated immunity, similar to observations from studies of sterile-environment-induced cell death. Thus, the goal of this study was first to generate tools and then to subsequently examine how cell death induced by L. monocytogenes influences the generation of protective immunity. We developed a unique set of L. monocytogenes strains that hyperactivate pyroptosis, apoptosis, or necrosis—forms of cell death that L. monocytogenes can naturally induce throughout infection (6–8), allowing us to examine this question within a single model system. In vitro characterization of these strains in myeloid-derived antigen-presenting cells demonstrated that they robustly activated specific cell death pathways. Importantly, activation of a single pathway by a given strain was not absolute. Nevertheless, following immunization with these strains, we investigated in detail in vivo how T-cell responses are affected and how cell death influences each of the 3 signals required for CD8+ T-cell generation. Our results suggest that in the context of L. monocytogenes-induced immunity, modulation of induction of cell death interferes with different signals required for optimal CD8+ T-cell activation, ultimately inhibiting the host's ability to generate protective immunity. More specifically, strains engineered to hyperinduce necrosis relative to other cell death pathways in primarily infected antigen-presenting cells were associated with a hyperinflammatory state and a premature loss of both CD8α+ DCs and expression of critical costimulatory molecules. In contrast, skewing the cell death responses toward pyroptosis hyperstimulated expression of costimulatory molecules and resulted in an inflammatory milieu that was suboptimal for the generation of CD8+ T-cell-mediated immunity.
Based on the literature of sterile cell death, we hypothesized that strains activating proinflammatory necrosis would be the most immunogenic strains in our study. However, immunization with Lm-necro provided the least protective immunity. LLO has previously been demonstrated to induce lymphocyte apoptosis in vitro and in purified form in vivo (46). Although we did not observe increases in T-cell apoptosis associated with Lm-necro infection (data not shown), we cannot rule out the possibility that LLO-induced bystander T-cell apoptosis contributes to the loss of immunity with Lm-necro. Based on our data, however, our favored hypothesis is that impairments in DC costimulation 48 h postimmunization (Fig. 6B and D), coupled with the loss of the critically important CD8α+ DC subset, is largely responsible for the protective immunity defect following immunization with Lm-necro. Consistent with the sterile cell death literature, we find that when we separate the act of necrosis from antigen presentation and DC costimulation, L. monocytogenes-stimulated necrosis can supplement a significant CD8+ T-cell response. Necrosis, though traditionally immunogenic, can be tolerogenic following chemotherapy treatments. However, tumor immunogenicity can be rescued following necrosis-induced cell death by administration of an anti-CD40-activating antibody (47). CD40, a critically important costimulatory molecule, induces upregulation of other costimulatory molecules required during antigen presentation, including CD80 and CD86 (48), and can substitute for CD4+ T-cell help to prime and fully activate CD8+ T cells (49). Seven days postimmunization with Lm-necro, we found fewer total CD8+ T cells, suggesting that the potential loss of CD40 or loss of CD4+ T-cell help is detrimental in our system. Importantly, CD40 activation is better than other inflammatory signals, including LPS and TNF-α, at inducing costimulatory molecule expression (48). Similarly, the presence of L. monocytogenes PAMPs in our model was unable to sustain appropriate CD86 expression and ultimately unable to provide robust protective immunity. Though surprising based on the sterile cell death literature, our work is also in line with observations of necrotic cell death induced by Mycobacterium tuberculosis. Virulent M. tuberculosis can actively inhibit apoptosis and induce necrosis to promote its pathogenesis (50) and as such delay the cross-priming essential for initiating a T-cell response (51). Though not examined in the context of cell death, administration of an anti-CD40-activating antibody in the presence of M. tuberculosis was able to increase cytotoxic T-cell functions, further suggesting that overcoming impairments in CD40 is essential in mounting an effective cell-mediated immune response, particularly in the context of necrosis.
In a sterile environment, apoptosis traditionally has been viewed as tolerogenic and can result in impairments in DC activation (11). In our model, we do not see impairments in DC activation or in antigen presentation. However, when we look at how inflammation induced by apoptosis impacts cell-mediated immunity, we see that the apoptotic inflammatory environment fails to stimulate CD8+ T-cell development to that of Lm levels, although it does stimulate these cells modestly. Apoptosis can result in the production of immunosuppressive mediators, such as TGF-β, IL-10, and PGE2 (52, 53), and suppression of the proinflammatory cytokine IL-12 (54). Apoptosis in an infection-based setting produces a unique environment whereby there is simultaneous recognition of anti-inflammatory signals from apoptosis and proinflammatory signals from PAMPs, which in the context of Citrobacter rodentium infection polarizes the immune system to result in a Th17 response (55). The role of IL-17-producing T cells has been controversial in regard to L. monocytogenes infection. IL-17 production from γδ T cells has been shown to be required for induction of primary CD8+ T-cell responses (56); however, adoptively transferred IL-17+ cells that express the inhibitory marker CTLA4 were unable to clear L. monocytogenes infection and failed to develop cytolytic activity and effector cytokine responses (57). These data suggest that IL-17-producing cells could be important in inducing robust primary CD8+ T-cell responses; however, the surface markers on the T cells may limit the effect of IL-17-producing T cells. These different T-cell populations could explain why we see CD8+ T cells that lack classic “Th1” effector functions and consequently fail to provide protective immunity.
Pyroptosis is a more recently identified form of cell death, and little is known about how it influences the adaptive immune response in either a sterile or infection-based environment. The inflammasome receptor AIM2 has recently been implicated in mounting fully protective humoral and cell-mediated responses to DNA vaccines (23). The Nlrp3 inflammasome signaling pathway has also been shown to be essential in mounting an antitumor response through the production of IL-1β (26). Finally, inflammasome adaptor protein apoptosis-associated speck-like protein containing CARD (ASC) and caspase-1 are needed for protective humoral immunity against influenza challenge and modified vaccine Ankara virus (24, 25). Our lab has previously examined the role of pyroptosis on cell-mediated immunity, and we observed impairments in CD8+ T-cell responses following robust inflammasome activation by L. monocytogenes (34, 58). The discrepancy between the role of the inflammasome in L. monocytogenes-stimulated immunity and other pathogens may be due to the specific route of immunization, the desired polarization of the immune response, or simply, differences in response to different pathogens. Here, we characterize the mechanism by which inflammasome activation inhibits the generation of antigen-specific T-cell responses to L. monocytogenes immunization. Specifically, we found that immunization with Lm-pyro does not result in defective antigen presentation, nor does it cause lack of CD86 costimulatory molecules within the first 48 h of immunization on the critical cross-presenting CD8α DC population. In fact, immunization with Lm-pyro results in increased DC activation throughout the first 48 h postimmunization compared to Lm, potentially due to increased IFN-γ levels. It is unclear how early, increased costimulatory levels influence the development of immunity in this model system. Recent reports suggest that the levels of costimulation can impact the proportion of effector versus memory cells that are formed (59), and it is possible that early, increased costimulation is altering the ultimate response generated from antigen-specific CD8+ T cells following pyroptosis. In addition to the potential role of increased DC costimulation, pyroptosis may also negatively influence the inflammatory milieu optimal for supporting L. monocytogenes-stimulated immunity.
The isolation of cell death-associated inflammation independently of antigen presentation and costimulation illuminates a pattern of immune activation different from our standard immunization model. In this context, Lm-apo and Lm-necro are able to provide an effective boost over PBS alone almost to Lm levels, whereas Lm-pyro fails to provide any increased generation in CD8+ T cells with effector function over that in PBS. One hypothesis is that early, robust inflammation as occurs following pyroptosis is a signal to the immune system to prevent full expansion of effector CD8+ T cells. Pyroptosis has a unique inflammatory milieu largely characterized by the release of IL-1β and IL-18 and the production of eicosanoids (60). Eicosanoids are lipid signaling molecules that exert a variety of both pro- and anti-inflammatory effects. Prostaglandin E2 has been one of the most studied eicosanoids and can suppress T-cell cytotoxic functions and T-cell activation (61). Additionally, PGE2 can skew the balance of Th1 versus Th2 cells toward Th2 (62) and can induce T-regulatory cells (Tregs) in an antigen-specific manner (63). We hypothesize that eicosanoids, and more specifically prostaglandin E2, downstream of pyroptosis may be responsible for the inhibition of protective immunity seen in our system. Eicosanoid production may result in a failure to activate CD8+ T-cell effector functions or skew antigen-specific cells to produce IL-4, IL-5, and IL-10, which would inhibit the protective response. Additionally, pyroptosis-dependent eicosanoid production may be increasing the role of Tregs, further dampening an effective CD8+ T-cell response.
When we compare our results to those in the sterile cell death literature, interesting differences emerge: classically, apoptosis has been considered tolerogenic while necrosis has been considered immunogenic (11, 13–16). This dichotomy has recently been muddled with the identification of immunogenic apoptosis and the direct study of cell death in vivo as opposed to in vitro. In our model system, all forms of cell death induced by L. monocytogenes are detrimental to forming a protective immune response. The majority of the sterile cell death literature has involved the injection of a bolus of dead cells in the context of antigen, a one-time stimulation event that is likely quickly cleared (11). In contrast, our model system likely results in a continued release of dead cells for as long as L. monocytogenes retains its virulence. Additionally, the location of injected dead cells has impacted the outcome of sterile immunity studies, as apoptotic cells injected intravenously promote tolerance while apoptotic cells injected subcutaneously promote immunity, potentially due to the type of cell that initially encounters the dying cell (reviewed in reference 64). In our model system, cells are dying in vivo in their natural location. Thus, it is important to consider that our observations may not be due just to the presence of the immune system seeing cell death in the context of an infection but also to the continued presence of dead cells or the location of the dying cells. Furthermore, although it is clear that skewing cell death in the context of L. monocytogenes infection influences the outcome of immunity, unlike what is reported in the sterile cell death literature, cell death induced by the strains of L. monocytogenes in this study were not on/off or even binary, as it is clear that even in vitro multiple forms of cell death take place (e.g., caspase 3/7 activation in Lm-pyro- or Lm-necro-infected cells or LDH release in Lm-apo-infected cells). Additionally, how cell death of primarily infected antigen-presenting cells influences death pathways of bystander cells is an important future question. Finally, the impact of the presence of PAMPs and DAMPs together is unique in our model system compared to studies in the sterile cell death literature, and as such, studies in the future could focus on sterile cell death in the context of pathogen-derived signals (PAMPs) to determine how this influences the resulting immune response.
L. monocytogenes largely avoids activation of cell death of its host cell to promote its pathogenesis, and an unintended consequence of this appears to be the activation of a robust protective immune response. Indeed, even the apoptotic death of uninfected bystander cells due to increased type I IFN production has been demonstrated to be detrimental in the context of L. monocytogenes-stimulated CD8+ T-cell responses (65). We suggest that the recognition of cell death in this system alerts the host to a pathological, tissue-damaging infection that results in the quicker clearance of pathological strains but, as a trade-off, diminishes the formation of a long-lasting protective response. When L. monocytogenes overactivates innate immune signaling, as through inflammasome activation or type I IFN production, adaptive immunity is ultimately impaired (34, 65). These results suggest a careful balance between clearing a host-damaging infection and mounting a long-lasting protective response, where the host is willing to sacrifice long-term gains for immediate survival. Our results suggest that for effective CD8+ T-cell-mediated vaccine development, inhibition or avoidance of host cell death will be critical in developing effective therapies and should be carefully considered when selecting infectious platforms for the generation of novel vaccines, including L. monocytogenes-based immunotherapies.
MATERIALS AND METHODS
Ethics statement.This work was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All protocols were reviewed and approved by the University of Wisconsin—Madison Institutional Animal Care and Use Committee.
Strains and strain construction.All L. monocytogenes strains used in this study were in the 10403s background and were cultured in brain heart infusion (BHI) medium. Lm, the parent vaccine strain, was constructed with in-frame deletions of actA and inlB, as previously described (33). For construction of the Lm-apo strain (JDS247), codon-optimized mouse bims was constructed using a g-block (IDT, Coralville, IA) with the restriction enzymes BamHI and NotI and cloned downstream of the actA promoter in frame with the secretion signal of the amino-terminal 300 bp of the actA gene in the site-specific integration vector pPL1. The construct was integrated into the comK locus of the attenuated ΔactA ΔinlB background as previously described (33, 66). Strain Lm-pyro (JDS124) was engineered to express Legionella pneumophila flaA under the control of the actA promoter and in frame with the amino-terminal 300 bp of the ActA gene in the site-specific integration vector pPL1 in the L. monocytogenes 10403s ΔactA ΔinlB background as previously described (34). Strain Lm-necro (JDS331) was constructed by introducing the point mutations T130G and T132C to induce the amino acid change S44A and the point mutation C1381A to induce the amino acid change L461T into the hly gene to reconstruct the S44A L461T listeriolysin O (LLO) mutant. These mutations result in increased LLO expression (S44A) and LLO activity at a neutral pH (L461T) as previously described (35). This mutated version of hly was introduced into the L. monocytogenes 10403s ΔactA ΔinlB background via double homologous recombination using pKSV7 as previously described (67). All immunizing strains expressed full-length OVA and the B8R20–27 epitope unless otherwise indicated. The B8R20–27 epitope and full-length OVA were engineered as a single fusion protein behind the actA promoter and in frame with the secretion signal of the amino-terminal 300 bp of the ActA gene (ActAN100) in the site-specific pPL2e vector as previously described (34).
Mouse strains and macrophages.Six- to 8-week-old C57BL/6 female mice were obtained from NCI/Charles River NCI facility or The Jackson Laboratory. Caspase-1/11−/− mice were a kind gift from Russell Vance (University of California, Berkeley, CA). CD11c-yfp mice were a kind gift from Matyas Sandor (University of Wisconsin, Madison, WI). Bone marrow-derived macrophages and dendritic cells (DCs) were made from 6- to 8-week-old female C57BL/6 or caspase-1/11−/− mice as previously described (68, 69). Briefly, macrophages were cultured from bone marrow in the presence of M-CSF derived from transfected 3T3 cell supernatant for 6 days, with an additional supplement of M-CSF medium 3 days postharvest. Dendritic cells were cultured in 100-mm dishes in the presence of 20 ng/ml recombinant GM-CSF (BD Biosciences, San Jose, CA) for 7 to 10 days with additional supplementation of 20-ng/ml GM-CSF-containing medium every third day.
Cell death characterization.For lactate dehydrogenase (LDH) release assays, bone marrow-derived macrophages (BMDMs) or dendritic cells (BMDCs, GM-CSF stimulated cells) were plated in 24-well plates at 5 × 105 cells per well and primed with 100 ng/ml Pam3CSK4 (Invivogen, San Diego, CA) overnight before infection. L. monocytogenes was grown slanted overnight at 30°C. BMDMs or BMDCs were infected at a multiplicity of infection (MOI) of 2, plates were spun at 500 × g for 5 min, and the infection was allowed to proceed for 15 min. Medium was aspirated and replaced with fresh medium containing 50 μg/ml gentamicin to kill extracellular bacteria. Medium with gentamicin was present on the cells for an additional 15 min before replacing with fresh medium without gentamicin. LDH in cell supernatants was assayed at the indicated time points after initial infection as previously described (75).
For caspase 3/7 activation, BMDMs or BMDCs (GM-CSF stimulated cells) were plated in white-bottom 96-well plates at 1 × 105 cells per well in 100 μl medium with 100 ng/ml Pam3CSK4 and allowed to adhere overnight. Infection at an MOI of 5 was allowed to proceed for 30 min, and medium was aspirated and replaced with fresh medium with 50 μg/ml gentamicin and 100 ng/ml Pam3CSK4. Caspase 3/7 activation was measured at the indicated time points with a Caspase-Glo 3-7 assay kit (Promega, Madison, WI) according to the manufacturer's instructions.
For annexin V characterization, BMDMs were plated in 35-mm non-tissue culture (non-TC)-treated petri dishes at 1 × 106 cells per dish. Cells were infected as described for the LDH assay. At the end of the assay, cells were lifted with PBS and stained with annexin V-fluorescein isothiocyanate (FITC) and propidium iodide per the manufacturer's instructions (Ebioscience, San Diego, CA).
OVA expression.Overnight cultures were grown in PrfA-inducing medium (LB-morpholinepropanesulfonic acid [LB-MOPS] with 25 mM glucose-1-phosphate and 0.2% activated charcoal) (70) to induce ActA expression, back-diluted 1:20 into fresh PrfA-inducing medium, and grown at 37°C for 5.5 h. Cultures were pelleted, and the supernatant was precipitated with trichloroacetic acid (TCA). Samples were separated on a 10% SDS-PAGE gel. A rabbit polyclonal antibody against the N-terminal portion of ActA (71) was used to detect the secreted fusion protein (ActAN100-OVA-B8R) at a 1:4,000 dilution, and a mouse monoclonal antibody was used to detect the constitutively expressed protein (72) and loading control, p60 (Adipogen, San Diego, CA), at a dilution of 1:2,000.
In vivo infections.All mice were immunized intravenously with logarithmic phase L. monocytogenes ΔactA ΔinlB diluted in 200 μl of phosphate-buffered saline (PBS) at the doses indicated. For acute infections, mice were sacrificed at 24 or 48 h postinfection. For challenge studies, mice immunized 30 days prior were challenged with 2 LD50 (2 × 105 CFU) of virulent L. monocytogenes expressing full-length OVA and the B8R20–27 and analyzed 68 to 72 h postinfection. Organs were homogenized in 0.1% Nonidet P-40 in PBS and plated on LB plates to quantify bacterial loads.
T-cell and dendritic cell analysis.Spleens taken at the indicated time points postimmunization were made into a single cell suspension, and red blood cells were lysed in ammonium-chloride-potassium (ACK) buffer. Total splenocytes were counted with a Z1 Coulter Counter, and a total of 1.7 × 106 splenocytes were stained for analysis. For primary and recall responses, splenocytes were stimulated ex vivo for 5 h with 2 μM OVA257–264 (SIINFEKL) or B8R20–27 (TSYKFESV) in the presence of brefeldin A (Ebioscience). Stimulated cells were surface stained with anti-CD4 (clone RM4-5) and anti-CD8α (clone 53-6.7) antibodies, fixed and permeabilized using IC fixation and permeabilization buffer (Ebioscience), and then stained intracellularly for IFN-γ (clone XMG1.2), TNF-α (clone MP6-XT22), and IL-2 (clone JES6-5H4). For tetramer analysis, cells were stained with B8R-tetramer (NIH Tetramer Facility, Atlanta, GA), followed by surface staining with anti-CD8α and anti-CD44 (clone IM7) antibodies. Dendritic cells were stained with markers anti-CD11c (clone N418), anti-CD86 (clone GL1), and anti-CD8α. All fluorophore-conjugated antibodies were obtained from Ebioscience. Samples were acquired using an LSRII flow cytometer (BD Biosciences) with fluorescence-activated cell sorter (FACS) DIVA software (BD Biosciences). Data were analyzed using FlowJo software (Treestar, Ashland, OR).
Antigen presentation.Spleens were removed from immunized mice 24 and 48 h postinfection, collagenase treated, and made into a single-cell suspension. CD11c+ cells were isolated using CD11c+ magnetic beads (Miltenyi, San Diego, CA) on an AutoMACS Pro machine (Miltenyi) per the manufacturer's instructions. CD11c+ cells (8 × 105) were incubated with 1 × 105 SIINFEKL-specific B3Z T cells as previously described (73) in the presence of 50 μg/ml gentamicin to prevent extracellular L. monocytogenes growth. Tetracycline (75 μg/ml) was also added to prevent further OVA secretion from L. monocytogenes during ex vivo incubation. Cells were incubated for 20 to 24 h, washed, and lysed with 0.5% NP-40 containing 0.15 mM chlorophenol red-β-d-galactopyranoside (CPRG) in PBS. Absorbance was read at 595 nm 24 h postlysis. The assay was sensitive enough to detect 10 pM OVA peptide presented based on standard curves run in parallel.
Dendritic cell immunization and L. monocytogenes boost.LPS-matured Flt3L-stimulated DCs were isolated as previously described (74). Briefly, mice were injected subcutaneously with Flt3L-expressing B16 cells for 3 weeks. Sixteen hours before splenocyte harvest, mice were injected with 2 μg lipopolysaccharide (Invivogen). Splenocytes from Flt3L tumor-bearing mice were pulsed with 2 μM B8R20–27 peptide and purified using a CD11c+ cell isolation kit (Miltenyi). Peptide-loaded cells (5 × 105) were injected intravenously into 6- to 8-week-old female C57BL/6 mice. Isolated cells were checked for purity and were >85% pure. Twenty-four hours following DC immunization, mice received an inflammatory boost with 1 × 103 CFU of L. monocytogenes lacking B8R expression or received a PBS control.
Cytokine analysis.Blood was obtained by cardiac puncture, and serum was analyzed by cytokine bead array with the mouse inflammation kit (BD Biosciences, San Jose, CA) according to the manufacturer's instructions. Samples were acquired using an LSRII flow cytometer with FACS DIVA software.
Statistical analysis.Statistical analysis was performed using GraphPad Prism Software (La Jolla, CA), and results were analyzed with a one-way analysis of variance (ANOVA) with Bonferroni's correction unless otherwise indicated.
ACKNOWLEDGMENTS
We thank John Harty for generously providing the B16Flt3L-expressing cell line and Nilabh Shastri for generously providing the B3Z hybridoma cell line. We also thank Suresh Marulasiddappa for critical reading of the manuscript. We acknowledge the NIH Tetramer Core Facility (contract HHSN272201300006C) for provision of MHC-I B8R tetramers.
This work is supported by grants from the NIH (R01 CA188034 to J.-D.S. and F30 CA210912 to E.T.) and an AAI Careers in Immunology Fellowship to E.T. and J.-D.S. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
FOOTNOTES
- Received 2 November 2016.
- Accepted 4 November 2016.
- Accepted manuscript posted online 7 November 2016.
Supplemental material for this article may be found at https://doi.org/10.1128/IAI.00733-16 .
- Copyright © 2016 American Society for Microbiology.
