Listeria-Infected Myeloid Dendritic Cells Produce IFN-, Priming T Cell Activation 1
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免疫学杂志 2005年第13期
Abstract
The intracellular bacterium Listeria monocytogenes infects dendritic cells (DC) and other APCs and induces potent cell-mediated protective immunity. However, heat-killed bacteria fail to do so. This study explored whether DC differentially respond to live and killed Listeria and how this affects T cell activation. To control for bacterial number, a replication-deficient strain, Lmdd, defective in D-alanine biosynthesis, was used. We found that DC internalize both live and heat-killed Lmdd and similarly up-regulate the expression of costimulatory molecules, a necessary step for T cell activation. However, only live Lmdd-infected DC stimulate T cells to express the early activation marker CD69 and enhance T cell activation upon TCR engagement. Infection with live, but not heat-killed, Lmdd induces myeloid DC to secrete copious amounts of IFN-, which requires bacterial cytosolic invasion. Exposure to high concentrations of IFN- sensitizes naive T cells for Ag-dependent activation.
Introduction
The intracellular, Gram-positive bacterium Listeria monocytogenes (Lm) 4 induces such a potent T cell-mediated immune response that it is one of the main models for studying T cell immunity in mice and is being developed as a vaccine vector to deliver Ags derived from infectious agents or tumors (1, 2, 3, 4). One of the reasons for the powerful immune response to Lm may be that Lm infects professional APCs: monocytes, tissue macrophages, and dendritic cells (DC). DC play a pivotal role in directing T cell responses. To activate naive T cells, DC must undergo a maturation process, in which DC up-regulate the expression of MHC, CD40, CD80, and CD86 surface molecules and cytokines. DC maturation can be induced by a variety of stimuli, including ligation of TLR, which recognize molecular patterns of infectious agents (5, 6, 7). TLR2 on DC binds to the Lm cell wall components, lipotechoic acid, and peptidoglycan (8), and TLR9 recognizes bacterial unmethylated CpG DNA (9).
DC are heterogeneous, and involvement of different DC subpopulations may dictate the outcome of subsequent immune responses. CD11c+ DC that also express CD11b, but not CD8, are termed myeloid DC (mDC). A second subset, plasmacytoid DC (pDC) (10), lack CD11b, but express B220, and are thought to be the major source of the type I IFNs in vivo (11). However, a recent report suggests that under certain conditions, viruses, such as lymphocytic choriomeningitis virus or influenza, can induce the production of high levels of type I IFN by non-pDC (12).
Immunization with killed Lm does not induce protective immunity (13). Similarly, adoptive transfer of DC infected with live Lm, but not with heat-killed (HK) Lm, protects against subsequent Lm infection (14). This is despite the fact that HK bacteria express a broad spectrum of immunostimulatory molecules capable of binding and activating TLRs. Why HK Lm fail to induce protective immunity remains unknown. Production of cytokines, such as IFN- and IL-12, or CD40 signaling may be important in inducing protective immunity after live Lm infection (15, 16, 17). Although immunization with HK Lm primes memory Lm-specific CD8 T cells, it does not induce them to differentiate into effector T cells and is much less efficient at activating their clonal expansion (18). The underlying mechanism for this differential effect remains to be defined. Although the effect of Lm infection on macrophages has been studied extensively (19, 20, 21, 22), few efforts have focused on how DC respond to Lm infection (23).
Because DC are believed to be the key professional APC capable of priming naive T cells (24), we compared the effects of treatment with live and HK Lm on mouse mDC. To minimize possible differential effects that might be due to increased numbers of live bacteria, an attenuated nonreplicating strain, Lmdd, deficient in D-alanine (D-Ala) biosynthesis (25), was used in this study. Lmdd does not replicate in the absence of exogenous D-Ala, but is nonetheless able to stimulate T cell immunity if D-Ala is provided during inoculation (25, 26). Both HK and live Lmdd similarly activate DC to up-regulate costimulatory molecules and secrete most cytokines. However, supernatants derived from DC cultured with live Lmdd, but not HK Lmdd, activated polyclonal T cells via a rapidly produced soluble factor independently of MHC-TCR ligation. This soluble factor was identified as IFN-. Microarray analysis, comparing genes expressed by DC treated with either live or HK Lmdd, and intracellular cytokine staining confirmed the induction of type I IFNs only by live Lmdd-treated DC. IFN- acts as a T cell commitment factor; it significantly decreases the dose-response threshold of naive T cells for subsequent activation by the TCR and enhances T cell priming. Using mutant bacteria unable to escape the phagolysosome, we found that cytosolic invasion is required to induce mDC to produce IFN-.
Materials and Methods
Mice and cells
Wild-type BALB/c and C57BL/6, and 2-microglobulin–/–, CIITA–/– mice in the H-2b background were obtained from The Jackson Laboratory. OT-1 mice (27) were provided by H. Eisen (Massachusetts Institute of Technology Center for Cancer Research, Cambridge, MA). MyD88–/– mice (28) were provided by M. Boes (Harvard Medical School, Boston, MA). Mouse bone marrow-derived DC (BMDC) were generated as previously described (29) by culture in GM-CSF and IL-4, followed by positive immunomagnetic selection using CD11c Ab-coated microbeads (Miltenyi Biotec). The selected BMDC were >98% CD11c+ DC by flow cytometric analysis. These DC are mDC and CD11c+ and B220–, and express high levels of CD11b and MHC class II and moderate levels of CD80 and CD86 (see Fig. 1A; data not shown). The pDC were generated by culturing mouse bone marrow cells in Flt3 ligand-conditioned medium for 9 days. The macrophage cell line RAW264.7 was obtained from American Type Culture Collection, and the mDC cell line DC2.4 (30) was provided by K. Rock (University of Massachusetts Medical School, Worcester, MA). Cells were cultured in RPMI 1640 medium containing 2 mM L-glutamine, 100 U/ml penicillin G, 50 μg/ml streptomycin sulfate, 50 μM 2-ME, and 10% FBS, unless otherwise indicated.
Bacteria
L. monocytogenes strain Lmdd was provided by F. Frankel (University of Pennsylvania, Philadelphia, PA) (25). DP-L3885 (inducible listeriolysin O (LLO)), DP-L1942 (Act-A–), and DP-L2612 (hly–) strains were provided by D. Higgins (Harvard Medical School, Boston, MA) (31, 32). Bacteria were grown in brain-heart infusion medium (BD Biosciences) supplemented with 100 μg/ml D-Ala and washed to remove D-Ala before use. HK bacteria were prepared by treatment at 60°C for 1 h. In some experiments, Lmdd were killed by treatment with 100 μg/ml gentamicin for 1 h, by sonication (1010 CFU/ml bacteria in PBS were sonicated three times for 20 s each time at 4°C; Heat Systems), or by physical lysis using 425- to 600-μm diameter glass beads (Sigma-Aldrich). For CFSE (Molecular Probes) staining, Lmdd were incubated with 1.5 μM CFSE in PBS at room temperature for 15 min, followed by three washes. Cells were infected with 5–10 CFU of bacteria/cell. Lm-treated cells were added to T cells 4 h after culture at 37°C.
Generation of culture supernatants
Supernatants harvested at the indicated time or 12 h after Lmdd treatment from DC plated at 106 cells/ml in serum-free RPMI 1640 were passed through a 0.2-μm pore size filter and stored at –80°C before use. In some experiments, Lmdd-infected cells were treated with the indicated concentration of cycloheximide 1 h after infection, then cultured overnight before harvesting supernatants. Supernatants were desalted using Econo-Pac desalting columns (Bio-Rad) to remove cycloheximide before addition to T cell cultures.
Microarray assay and RT-PCR
On day 7 of culture, mouse BMDC were enriched using anti-CD11c-conjugated magnetic beads and treated with medium, LPS (1 μg/ml), 10 CFU of HK or live Lmdd/cell for 6 h before extracting total RNA using the Qiagen RNeasy Mini kit. Approximately 10 μg of each sample was used to make labeled probes and was prepared for hybridization to mouse expression set A and B oligonucleotide arrays (Affymetrix), performed at the Microarray Core facility, Dana-Farber Cancer Institute. RNA was extracted and analyzed as described previously (33) using samples from two independent experiments. For RT-PCR, 1 μg of total RNA was used to generate cDNA using the TaqMan RT kit (Applied Biosystems). IFN- and -actin primers were previously described (34): ifnb: forward, 5'-ctggagcagctgaatggaaag; reverse, 5'-cttgaagtccgccctgtaggt; and -actin: forward, 5'-aggtgtgatggtgggaatgg; reverse, 5'-gcctcgtcacccacatagga. Two-step PCR was performed for 35 cycles at 95°C for 10 s and 62°C for 15 s.
Chromatography and mass spectrometry analysis
Supernatants derived from Lmdd-infected BMDC or DC2.4 cells cultured in serum-free RPMI 1640 were harvested, passed through a 0.2-μm pore size filter, concentrated, and separated on a Superdex-200 column. The apparent m.w. of fractions active in inducing CD69 expression on naive C57BL/6 splenocytes were determined by comparing their migrations with those of m.w. standards. Approximately 20 L of serum-free RPMI 1640 supernatant derived from Lmdd-infected DC2.4 cells was separated by sequential High S (Bio-Rad), heparin, hydroxyapatite, and Blue gel (Amersham Biosciences) chromatography. Active fractions were pooled, subjected to tryptic digestion, and analyzed by MALDI-TOF mass spectrometry at the Harvard Medical School Mass Spectoscopy Core facility.
Flow cytometry
Flow cytometry was performed using FACSCalibur and CellQuest software (BD Biosciences) with fluorophore-conjugated Abs to IL-6 (MP5-20F3), IL-12 (C15.6), TNF- (MP6-XT3), I-Ab (AF6-120.1), CD11b (M1/70), CD11c (HL3), CD4 (GK1.5), CD8 (53-6.7), CD25 (clone PC61), CD40 (3/23), CD43 (1B11), CD69 (H1.2F3), CD62L (MEL-14), CD80 (16-10A1), and CD86 (GL1) from BD Pharmingen. Rat anti-mouse IFN mAb (F18) was purchased from Hycult Biotechnology. Rabbit anti-mouse polyclonal Abs against IFN- and IFN- were obtained from PBL Laboratory. For external staining, 2 x 105 cells/microtiter well were washed with FACS buffer (PBS containing 2% heat-inactivated FBS and 0.1% sodium azide) and incubated with an FcR-blocking Ab (BD Pharmingen) for 5 min, then incubated with saturating amounts of mAbs for 30 min at 4°C. For intracellular cytokine staining, cells were treated with 20 μM brefeldin A for the last 12 h of culture or as indicated, resuspended in 50 μl of FACS buffer, and permeabilized using the Fix and Perm kit according to the manufacturer’s protocol (Caltag Laboratories). Saturating amounts of fluorochrome-conjugated Abs were added to the permeabilization buffer, and cells were incubated at room temperature for 15 min. Cells were washed and resuspended in FACS buffer with 1% formaldehyde for analysis.
T cell proliferation and ELISPOT assays
Splenocytes from naive C57BL/6 mice or CD8 T cells from OT-1 transgenic mice were harvested and exposed to 1000 U/ml or the indicated concentration of mouse rIFN- (1.1 x 108 U/mg; PBL Laboratory) or IFN- (1.2 x 107 U/mg; PBL Laboratory) or supernatant derived from Lmdd-infected DC for 3 h and washed. Splenocytes were cultured with the indicated concentrations of anti-CD3 (BD Pharmingen). OT-1 CD8 T cells were cultured with irradiated splenocytes and the indicated concentrations of the cognate OVA peptide SIIFNEKL. Alternatively, splenocytes were directly cultured with the indicated concentration of anti-CD3 in the presence of IFN- or DC supernatant. T cell proliferation was measured by [3H]thymidine incorporation. IFN- production was measured by ELISPOT, as previously described (35), using an immunospot counter (Cellular Technology).
Results
Both HK and live Lmdd are similarly taken up by DC and induce DC maturation
To begin to define the differences in the immune-stimulating effects of live or HK Lmdd, we analyzed the responses of mouse BMDC and the DC cell line DC2.4 to live and HK Lmdd. Both BMDC and DC2.4 cells are myeloid lineage cells expressing CD11c and CD11b, but not B220 or CD8 (Fig. 1A). BMDC internalized both live and HK Lmdd. The number of internalized bacteria was similar 5 h after adding live or HK bacteria, as quantified by flow cytometry (mean fluorescence intensity (MFI), 798 HK vs 771 live Lmdd; Fig. 1B). Actin tails costained with CFSE-labeled bacteria were only evident in live Lmdd-infected DC (data not shown), suggesting that live Lmdd escape from the vacuole to the cytosol, whereas HK Lmdd remain in the vacuole (20). BMDC (Fig. 1C) and DC2.4 cells (data not shown) exposed to live or HK Lmdd for 24 h up-regulated the surface expression of molecules involved in T cell activation, such as CD40, CD80 (B7.1), CD86 (B7.2), and MHC class II (Fig. 1C). Although CD86 and the MHC class II molecule I-Ab were up-regulated to a similar extent, CD40 and B7.1 expression were slightly higher in live Lmdd-treated DC (CD40: MFI, 129 live vs 112 HK; CD80: MFI, 105 live vs 91 HK). Thus, both live and HK Lmdd were similarly internalized by DC, and both induced costimulatory molecules, with only subtle differences in magnitude.
HK and live Lmdd-treated DC differ in the ability to activate T cells
We next examined whether DC pulsed with live or HK Lmdd differentially activate T cells. Live bacteria-pulsed BMDC rapidly stimulated both CD4 and CD8 splenic T cells to express the early activation marker CD69, whereas HK Lmdd-treated DC had less of an effect (Fig. 2A). The reduced ability of HK Lmdd to activate DC capable of inducing CD69 expression on T cells was not due to heat inactivation of bacterial Ags or adjuvants, because Lmdd killed by other means (gentamicin, ultrasound, or glass beads) also did not efficiently activate DC (Fig. 2A). The Lmdd-infected DC2.4 DC line was also able to activate T cells to express CD69, whereas the infected RAW264.7 macrophage cell line had limited ability (Fig. 2A). We next examined whether T cells exposed to Lmdd-infected DC were fully activated. When CFSE-labeled CD8 T cells were cocultured with Lmdd-infected DC, they did not proliferate (Fig. 2, B and C), as determined by CFSE dilution. T cell activation markers, such as CD25 and CD43, were not up-regulated by 48 h (Fig. 2C), although CD62L expression was slightly down-regulated in T cells that had been cocultured with live Lmdd-pulsed DC (Fig. 2C). T cells did not produce IL-2, as determined by intracellular cytokine staining (data not shown). Therefore, T cell activation by Lmdd-infected DC was partial.
Because Lmdd infection introduces a great deal of foreign Ags and triggers DC maturation (Fig. 1), we examined whether activation of CD69 expression was dependent upon TCR engagement. T cells were cocultured with Lmdd-treated BMDC derived from MHC class I- or class II-null mice. CD69 expression on T cells was comparable after stimulation with wild-type or MHC-null DC (Fig. 2D). It was also comparable when T cells were cultured with Lmdd-infected allogeneic BMDC or syngeneic BMDC (Fig. 2D). These data suggest that live Lmdd-infected DC partially activate T cells through a TCR-independent process.
T cell activation by live Lmdd-infected DC is mediated by a secreted soluble factor(s)
Because the activation of T cells by live Lmdd-infected DC is independent of TCR-MHC engagement, we next determined whether it required cell-to-cell contact or was mediated by a soluble factor. Culture supernatant from live, but not HK, Lmdd-treated DC was fully able to induce CD69 expression on T cells (Fig. 3A). Supernatants derived from bacterial cultures without DC or from Lmdd-infected RAW246.7 macrophage cells had little capacity to activate CD69 expression (Fig. 3A), suggesting that the molecules that activate T cells were not produced by bacteria, but were secreted by DC after Lmdd infection. Secretion occurred rapidly within 5 h of infection, reached a peak by 12 h, and gradually declined after 20 h of culture (Fig. 3B). This molecule(s) was synthesized de novo, because treating DC with cycloheximide almost completely inhibited its production (Fig. 3C). Cycloheximide treatment did not affect cell viability, as assessed by forward and side scatter flow cytometry profiles, even at the highest concentration (data not shown).
Live Lmdd-infected DC supernatant enhance Ag-dependent T cell activation
To explore the effects of the soluble factors secreted by Lmdd-infected DC on Ag-dependent T cell activation, splenocytes from naive mice were pulsed with anti-CD3 Ab in the presence or the absence of DC culture supernatant. Lmdd-treated DC supernatant significantly enhanced the numbers of T cells secreting IFN- after anti-CD3 treatment (Fig. 4A). At limiting concentrations of anti-CD3, the dose-response curve was shifted by about 1 log to the left in the presence of Lmdd-treated DC supernatant. HK Lmdd-treated DC supernatant slightly enhanced T cell activation, whereas supernatant from uninfected DC had no effect on promoting Ag-dependent T cell activation (Fig. 4A). In addition, T cell proliferation in response to stimulation by allogeneic splenocytes was significantly enhanced when T cells were continuously cultured in supernatant from Lmdd-infected DC (Fig. 4B). Pre-exposure of T cells to Lmdd-infected DC supernatant also improved T cell proliferation in response to allogeneic splenocytes (Fig. 4B), suggesting that the soluble factors secreted by live Lmdd-infected DC sensitized T cells for subsequent Ag-dependent activation. The continuous presence of supernatant, however, induced more proliferation than when T cells were washed before exposure to allogeneic splenocytes (Fig. 4B).
IFN- expression is induced by live, but not HK, Lmdd-treated DC
To identify the secreted molecules produced by Lmdd-infected DC that activate T cells, we looked for differential production of proinflammatory cytokines, such as IL-12, IL-6, and TNF-, by treated DC (23). HK or live Lmdd infection led to a comparable increase in IL-6 production (Fig. 5A). IL-12 and TNF- production by live Lmdd-treated DC was slightly higher than that by HK Lmdd-treated DC (Fig. 5, B and C, and data not shown). However, Lmdd-infected DC secreted much less TNF- than infected RAW264.7 cells (Fig. 5C). Although Escherichia coli-infected DC have been reported to produce IL-2 (36), no IL-2 was induced after Lmdd infection (data not shown). Because the production of these cytokines was similar after HK or live Lmdd infection, none of these cytokines is probably responsible for the difference in T cell activation.
To identify the soluble DC factor capable of inducing CD69 expression and sensitizing T cell activation, Lmdd-infected DC supernatant was separated by chromatography, testing fractions for their ability to up-regulate CD69 expression on T cells. The apparent Mr of the soluble factor was determined by Superdex 200 gel filtration to be 24–28 kDa (data not shown). Lmdd-infected DC supernatant was separated through sequential cation exchange High S, heparin, hydroxyapatite, and Blue gel columns. The final active fraction was subjected to tryptic digestion and MALDI-TOF mass spectroscopy analysis, and four peptides with sequences matching IFN- were identified (data not shown). This suggested that IFN- might be the soluble factor responsible for CD69 up-regulation. This result was unexpected, because IFN- is thought to be produced mostly by pDC.
To determine whether IFN- is the sought-after soluble factor produced by Lmdd-infected DC, we first looked by microarray analysis at whether mDC differentially up-regulate IFN- mRNA expression after Lm infection (Table I). CD11c+ BMDC were treated for 6 h with medium, LPS, or HK or live Lmdd before isolating mRNA. LPS-treated DC had limited capacity to activate T cells to express CD69 (data not shown). Twenty-seven genes were up-regulated by at least 10-fold after live Lmdd infection compared with untreated DC. Sixteen of these were also up-regulated at least 10-fold by treatment with HK Lmdd. The gene that showed the greatest modulation, however, was IFN-, whose expression was increased 67-fold. Moreover, IFN- expression was not enhanced by HK Lmdd and was only enhanced 4-fold by LPS. By semiquantitative RT-PCR, no IFN- mRNA was detected in untreated BMDC, but expression was up-regulated in Lmdd-infected BMDC (Fig. 6C). Samples from LPS- and HK Lmdd-treated DC had detectable IFN- expression, but substantially less than samples from Lmdd-infected DC. These data were also confirmed by quantitative real-time PCR (data not shown). Other genes that were up-regulated differentially by live Lmdd compared with both HK Lmdd and LPS were IFN- genes 2 (increased 19-fold) and 5 (increased 8-fold). IFN- expression was up-regulated in BMDC to a similar extent by live Lmdd (6-fold) or LPS (8-fold), but only 2-fold by HK Lmdd treatment. In addition, a number of IFN-inducible genes and cytokine and chemokine genes and receptors as well as some genes with unknown function were up-regulated substantially by 6 h, but they were also up-regulated in HK Lmdd-treated and/or LPS-treated DC (Table I). The inducible IFN-regulated factor 7 (IRF7) gene was significantly up-regulated by treatment with live Lmdd (13-fold) or LPS (15-fold) and less so by HK Lmdd (4.5-fold; Table I), indicating that the positive feedback loop of type I IFN gene expression was rapidly activated by all these treatments (37, 38). The low levels of IFNs induced by LPS or HK Lmdd appeared to be sufficient to activate a wide array of IFN-inducible genes. However, although IRF3 mRNA remained unchanged, IRF3 protein translocated from the cytoplasm to the nucleus only in DC treated with live Lmdd, as determined by subcellular fractionation and immunoblotting (data not shown). IRF3 nuclear translocation is the key to inducing IFN- expression (39).
Discussion
It has long been known that live Lm induce protective immunity, whereas killed bacteria fail to do so (13, 18). Unveiling the mechanism by which immune cells differentially respond to live vs dead Lm is important for understanding innate and adaptive immunity, and ultimately will help to design better vaccines against cancer and infectious diseases. Because DC are the key to priming immune responses, in this study we focused on differential effects of live and HK bacteria on DC activation and priming of T cells. We found that although live and HK Lm similarly cause DC to mature, as measured by up-regulating cell surface costimulatory molecules and MHC class II, only live bacteria-infected DC are able to activate CD69 expression on T cells efficiently and prime them for subsequent activation by the TCR. Using protein chemistry and microarray gene expression analysis, we identified the type I IFN, IFN-, as a soluble factor rapidly produced in large amounts (3000 U/106 cells) by BMDC upon infection by live Listeria, but not HK bacteria. Moreover, we were able to show that only high concentrations of type I IFNs are able to prime T cell activation. We therefore hypothesize that the exceptionally strong and protective immune response to Listeria is related to the amplification of T cell priming that occurs in the presence of high local concentrations of type I IFN secreted by infected DC.
The copious production of type I IFN in our studies was unanticipated, because our experiments were performed using mDC. Conventional wisdom holds that the major type I IFN-producing DC is the B220+CD11b– pDC (11). However, in our experiments the source of DC was either the B220–CD11b+ mDC cell line DC2.4 or BMDC that had been cultured in GM-CSF and IL-4 and selected for CD11c expression. These cultured BMDC are mDC, contain <1% B220+ cells, and are >98% CD11b+ (Fig. 1A). Furthermore, Lmdd infection did not alter the BMDC expression of cell surface markers that distinguish mDC and pDC (data not shown). Although the DC2.4 cell line is fully myeloid, we were concerned that contamination of our BMDC with a few pDC might account for the IFN production we measured in the BMDC cultures. However, when BMDC were cultured in the presence of Flt3 ligand to generate cultures enriched in pDC (30% B220+), the bioactivity of the Lmdd-infected DC-conditioned medium was not enhanced (Fig. 6B). These data therefore suggest that mDC produce high levels of type I IFN when infected with Listeria. Production of type I IFN by mDC was also documented in a recent study that showed that mDC produce large amounts of type I IFN after infection with viruses such as lymphocytic choriomeningitis virus (12).
Type I IFNs have multiple biological activities and play important roles in bridging innate immune responses and adaptive immunity (41). Both IFN- and IFN- trigger CD69 expression on T cells, but Lm infection triggered more IFN- than IFN- expression in mDCs. Ab-blocking experiments also suggested that most of the biological effect could be attributed to IFN-. In this study we also demonstrated that IFN- produced by Lm-infected DC acts as a commitment factor to decrease the Ag response threshold of T cells and enhance their priming. The initial exposure to IFN- partially activates naive T cells, preparing them for subsequent Ag-specific activation.
Stimulation of type I IFN production by mDC and their priming of T cell CD69 expression and sensitization for TCR stimulation require Lm escape from the phagolysosome. HK bacteria remain in the phagolysosome. Using bacterial mutants lacking or with inducible expression of LLO, which is required for cytosolic invasion, we also found that the production of IFN- and T cell activation by mDC correlate with the ability to escape the phagocytic vacuole. Portnoy and colleagues (20) previously reported production of IFN- by Lm-infected macrophages via a mechanism that requires bacterial cytosolic invasion. They postulated a bacterial cytosolic sensor that signals the presence of cytosolic bacteria and triggers the secretion of IFN- by macrophages. Our results suggest that a similar sensor and pathway are triggered by intracytoplasmic infection of mDC. The sensor remains to be identified. LPS activates a TLR4, MyD88-independent signaling pathway that induces IRF3 translocation and IFN- production via the Toll/IL-1R domain-containing adaptor (42, 43, 44). However, we found that LPS has a very limited capacity to stimulate BMDC to produce IFN- compared with live Lmdd, and LPS-activated DC do not efficiently stimulate T cells to express CD69. Supernatants from Lmdd-infected BMDC derived from mice deficient in MyD88, a key signaling molecule for TLR engagement, produce comparable amounts of IFN- (data not shown). It is, moreover, unlikely that TLR family proteins are responsible, because these receptors are displayed on the cell surface and within endosomes, but not in the cytosol. The nucleotide-binding oligomerization domain and Nacht, leucine-rich repeat, and pyrin domain-containing protein family proteins that are present in the cytosol and are able to sense Gram-positive and Gram-negative bacterial cell wall components are attractive candidates for the unknown sensor (45, 46, 47).
In this study we found that pulsing bone marrow-derived mDC with either HK or live Lmdd induces DC to up-regulate costimulatory molecules on their surface and secrete proinflammatory cytokines, such as TNF-, IL-6, and IL-12. DC maturation and induction of proinflammatory cytokines were probably stimulated by engagement of TLR receptors by both HK and live Lmdd. Because the differential effects of live and HK bacteria were also evident when bacteria were killed by other means (such as antibiotic treatment) that do not cause denaturation, differences in TLR engagement of pathogenic patterning molecules in the bacteria were probably minimal. It is, therefore, not surprising that dead Lm also induce DC maturation. Although slightly more CD40 and CD80 were detected on the surface of BMDC infected with live Lmdd than HK Lmdd, the expression of other cell surface markers (class II MHC and CD86) and that of inflammatory cytokine production were comparable. Therefore, these factors are unlikely to explain the large difference in T cell activation we observed. However, the difference in type I IFN expression was dramatic, and Abs to type I IFNs could abrogate the effect of Lmdd-infected BMDC supernatants on T cells.
Our conclusions differ from those in a recent paper that compared GM-CSF-cultured BMDC maturation, DC inflammatory cytokine production, and stimulation of T cells to produce IFN- in response to infection with wild-type and LLO-deficient Lm (23). That study also found that bacterial cytosolic invasion was critical for T cell activation, but found more substantial differences in costimulatory molecule expression and proinflammatory cytokine expression, which they interpreted as the distinguishing feature. Our use of a replication-defective Lmdd strain may account for differences between their study and ours (25). In fact, they had twice as much infection after 4 h using wild-type as hly– bacteria. Another difference is that the other study examined DC maturation 18 h after Lm infection, whereas we observed DC maturation after 24 h. Typically, costimulatory molecule up-regulation occurs relatively slowly compared with IFN- induction, requiring 16–24 h (data not shown). Type I IFN production in response to bacterial invasion may accelerate DC maturation because it induces a positive loop of type I IFNs and IFN-inducible gene up-regulation (38) that enhances Ag presentation function and DC maturation (48). However, DC matured more slowly via TLR engagement may reach the same final state for efficient Ag presentation by 24 h. Accelerated DC maturation may lead to a more rapid and effective immune response. A final difference is that the myeloid DC used in the other study were generated from bone marrow by culture in GM-CSF, whereas we generated myeloid DC by culture in both IL-4 and GM-CSF. The combination of subtle differences in cytokines, timing, and bacterial replication may account for the different conclusions. However, which in vitro condition more faithfully recapitulates in vivo conditions is impossible to predict.
Both live and HK bacteria induce the rapid expression of multiple IFN-responsive genes in BMDC within 6 h, the time after Lmdd exposure at which we performed our microarray analysis. The downstream activation of these genes, many of which are also up-regulated by LPS engagement of TLR4, is probably triggered by low levels of IFNs generated after TLR signaling. Our results are similar to a gene expression analysis of Lm- or LPS-treated bone marrow-derived macrophages, which found that both bacterial invasion and TLR engagement were able to activate many IFN-responsive genes (49). However, high concentrations of type I IFN (250 U/ml IFN-, 5000 U/ml IFN- to activate CD69 on 50% of T cells) are needed to induce CD69 on T cells and prime T cells for antigenic exposure. These high concentrations of type I IFN decrease the Ag response threshold of naive T cells by 1 log. Therefore, weak signals that might otherwise be ignored or produce an ineffective, tolerogenic, or even suppressive regulatory immune response can be activated in the presence of high levels of type I IFN. The partial TCR-independent activation of T cells by high concentrations of type I IFN may also play a part in the bystander T cell apoptosis and lymphocyte depletion observed after Lm infection, because these partially activated T cells are prone to spontaneous apoptosis (data not shown) (50). Although removing nonspecific T cells may make room for expansion of Lm-specific T cells, it might also interfere with maintaining effective memory.
Our results and those reported by Portnoy and colleagues (20) suggest that in addition to TLR engagement, there is a second alarm system in APCs, such as DC and macrophages, that is activated when bacteria invade the cytosol. This alarm triggers the nuclear translocation of IRF3 and the production of copious quantities of type I IFNs that enhance T cell sensitivity to antigenic stimulation. This alarm system may be triggered by intracellular pathogens to enlist the aid of the cells (T cells) most capable of eliminating intracellular pathogens. The induction of type I IFN production by mDC after infection with lymphocytic choriomeningitis virus and a mutant strain of influenza (12) suggests that this second alarm might also be triggered by some viral infections. It will be of interest to determine whether other viruses and cytosolic pathogens also trigger exuberant type I IFN expression. Because T cells are not important for eliminating extracellular pathogens, it makes sense that this response would not be elicited by most bacteria and would not be triggered by cell surface receptors, like the TLR system. Our data also suggest that mDC produce much more IFN- than macrophages after live Lmdd infection, because supernatant from the infected RAW264.7 macrophage cell line barely activates T cells to express CD69 (Figs. 2A and 3A). Therefore, this second alarm system may function more effectively in DC than in macrophages.
This specialized type I IFN danger response may explain why Lm is such an effective vector for priming T cell responses to Ags. Because Lm causes serious disease in immunosuppressed individuals, pregnant women, and newborns, most investigators developing Lm as a vaccine vector agree that the bacterium must be attenuated before human use is contemplated (51). Our results suggest that attenuated bacteria unable to escape from the phagolysosome may be impaired in T cell priming and lead to ineffective vaccines. In fact, the vaccine strains that appear the most promising are able to invade the cytosol. Screening for type I IFN production may be a useful tool for predicting which attenuated bacteria are likely to be effective vaccine vectors.
Although this study suggests that type I IFN production in response to Lm cytosolic invasion should send a strong signal to enhance T cell immunity, mice deficient in the common type I IFNR are actually better able to handle Lm challenge (34, 52, 53). This is unexpected, because type I IFNs generally protect against other types of infection, especially by viruses. In fact, the harmful effect of type I IFNs may be peculiar to Lm infection (52, 54). Carrero et al. (52, 54) found that LLO, the bacterial pore-forming protein, acts as a bacterial toxin to induce T cell apoptosis, and that partially or fully activated T cells are particularly prone to LLO-induced apoptosis. Type I IFNs probably accelerate this process, because they induce partial activation of nonspecific T cells and facilitate full activation of specific T cells. This idiosyncratic effect of LLO on T cells and the induction of type I IFN need to be considered in developing Listeria-based vaccine vectors. Because of the pleiotropic effects of IFNs on different immune cells, however, the resistance of mice null for the type I IFN receptor to Lm may be due to more than one factor.
Acknowledgments
We thank Lianfa Shi, Manjunath Swamy, Howell Moffat, Pascale Cossart, and Jay Berzofsky for useful advice, and Fred Frankel and Darren Higgins for providing the strains of Lm used in this study.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work was supported by National Institutes of Health Grant AI056922 (to J.L.) and T32HL066987 (to A.S.).
2 Current address: Division of Infectious Diseases, Tufts University School of Veterinary Medicine, North Grafton, MA 01536.
3 Address correspondence and reprint requests to Dr. Judy Lieberman, CBR Institute for Biomedical Research, 200 Longwood Avenue, Boston, MA 02115. E-mail address: lieberman@cbr.med.harvard.edu
4 Abbreviations used in this paper: Lm, Listeria monocytogenes; DC, dendritic cell; BMDC, bone marrow-derived DC; D-Ala, D-alanine; HK, heat killed; IPTG, isopropyl--D-thiogalactoside; IRF, IFN-regulated factor; LLO, listeriolysin O; mDC, myeloid DC; MFI, mean fluorescence intensity; pDC, plasmacytoid DC.
Received for publication January 7, 2005. Accepted for publication April 29, 2005.
References
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The intracellular bacterium Listeria monocytogenes infects dendritic cells (DC) and other APCs and induces potent cell-mediated protective immunity. However, heat-killed bacteria fail to do so. This study explored whether DC differentially respond to live and killed Listeria and how this affects T cell activation. To control for bacterial number, a replication-deficient strain, Lmdd, defective in D-alanine biosynthesis, was used. We found that DC internalize both live and heat-killed Lmdd and similarly up-regulate the expression of costimulatory molecules, a necessary step for T cell activation. However, only live Lmdd-infected DC stimulate T cells to express the early activation marker CD69 and enhance T cell activation upon TCR engagement. Infection with live, but not heat-killed, Lmdd induces myeloid DC to secrete copious amounts of IFN-, which requires bacterial cytosolic invasion. Exposure to high concentrations of IFN- sensitizes naive T cells for Ag-dependent activation.
Introduction
The intracellular, Gram-positive bacterium Listeria monocytogenes (Lm) 4 induces such a potent T cell-mediated immune response that it is one of the main models for studying T cell immunity in mice and is being developed as a vaccine vector to deliver Ags derived from infectious agents or tumors (1, 2, 3, 4). One of the reasons for the powerful immune response to Lm may be that Lm infects professional APCs: monocytes, tissue macrophages, and dendritic cells (DC). DC play a pivotal role in directing T cell responses. To activate naive T cells, DC must undergo a maturation process, in which DC up-regulate the expression of MHC, CD40, CD80, and CD86 surface molecules and cytokines. DC maturation can be induced by a variety of stimuli, including ligation of TLR, which recognize molecular patterns of infectious agents (5, 6, 7). TLR2 on DC binds to the Lm cell wall components, lipotechoic acid, and peptidoglycan (8), and TLR9 recognizes bacterial unmethylated CpG DNA (9).
DC are heterogeneous, and involvement of different DC subpopulations may dictate the outcome of subsequent immune responses. CD11c+ DC that also express CD11b, but not CD8, are termed myeloid DC (mDC). A second subset, plasmacytoid DC (pDC) (10), lack CD11b, but express B220, and are thought to be the major source of the type I IFNs in vivo (11). However, a recent report suggests that under certain conditions, viruses, such as lymphocytic choriomeningitis virus or influenza, can induce the production of high levels of type I IFN by non-pDC (12).
Immunization with killed Lm does not induce protective immunity (13). Similarly, adoptive transfer of DC infected with live Lm, but not with heat-killed (HK) Lm, protects against subsequent Lm infection (14). This is despite the fact that HK bacteria express a broad spectrum of immunostimulatory molecules capable of binding and activating TLRs. Why HK Lm fail to induce protective immunity remains unknown. Production of cytokines, such as IFN- and IL-12, or CD40 signaling may be important in inducing protective immunity after live Lm infection (15, 16, 17). Although immunization with HK Lm primes memory Lm-specific CD8 T cells, it does not induce them to differentiate into effector T cells and is much less efficient at activating their clonal expansion (18). The underlying mechanism for this differential effect remains to be defined. Although the effect of Lm infection on macrophages has been studied extensively (19, 20, 21, 22), few efforts have focused on how DC respond to Lm infection (23).
Because DC are believed to be the key professional APC capable of priming naive T cells (24), we compared the effects of treatment with live and HK Lm on mouse mDC. To minimize possible differential effects that might be due to increased numbers of live bacteria, an attenuated nonreplicating strain, Lmdd, deficient in D-alanine (D-Ala) biosynthesis (25), was used in this study. Lmdd does not replicate in the absence of exogenous D-Ala, but is nonetheless able to stimulate T cell immunity if D-Ala is provided during inoculation (25, 26). Both HK and live Lmdd similarly activate DC to up-regulate costimulatory molecules and secrete most cytokines. However, supernatants derived from DC cultured with live Lmdd, but not HK Lmdd, activated polyclonal T cells via a rapidly produced soluble factor independently of MHC-TCR ligation. This soluble factor was identified as IFN-. Microarray analysis, comparing genes expressed by DC treated with either live or HK Lmdd, and intracellular cytokine staining confirmed the induction of type I IFNs only by live Lmdd-treated DC. IFN- acts as a T cell commitment factor; it significantly decreases the dose-response threshold of naive T cells for subsequent activation by the TCR and enhances T cell priming. Using mutant bacteria unable to escape the phagolysosome, we found that cytosolic invasion is required to induce mDC to produce IFN-.
Materials and Methods
Mice and cells
Wild-type BALB/c and C57BL/6, and 2-microglobulin–/–, CIITA–/– mice in the H-2b background were obtained from The Jackson Laboratory. OT-1 mice (27) were provided by H. Eisen (Massachusetts Institute of Technology Center for Cancer Research, Cambridge, MA). MyD88–/– mice (28) were provided by M. Boes (Harvard Medical School, Boston, MA). Mouse bone marrow-derived DC (BMDC) were generated as previously described (29) by culture in GM-CSF and IL-4, followed by positive immunomagnetic selection using CD11c Ab-coated microbeads (Miltenyi Biotec). The selected BMDC were >98% CD11c+ DC by flow cytometric analysis. These DC are mDC and CD11c+ and B220–, and express high levels of CD11b and MHC class II and moderate levels of CD80 and CD86 (see Fig. 1A; data not shown). The pDC were generated by culturing mouse bone marrow cells in Flt3 ligand-conditioned medium for 9 days. The macrophage cell line RAW264.7 was obtained from American Type Culture Collection, and the mDC cell line DC2.4 (30) was provided by K. Rock (University of Massachusetts Medical School, Worcester, MA). Cells were cultured in RPMI 1640 medium containing 2 mM L-glutamine, 100 U/ml penicillin G, 50 μg/ml streptomycin sulfate, 50 μM 2-ME, and 10% FBS, unless otherwise indicated.
Bacteria
L. monocytogenes strain Lmdd was provided by F. Frankel (University of Pennsylvania, Philadelphia, PA) (25). DP-L3885 (inducible listeriolysin O (LLO)), DP-L1942 (Act-A–), and DP-L2612 (hly–) strains were provided by D. Higgins (Harvard Medical School, Boston, MA) (31, 32). Bacteria were grown in brain-heart infusion medium (BD Biosciences) supplemented with 100 μg/ml D-Ala and washed to remove D-Ala before use. HK bacteria were prepared by treatment at 60°C for 1 h. In some experiments, Lmdd were killed by treatment with 100 μg/ml gentamicin for 1 h, by sonication (1010 CFU/ml bacteria in PBS were sonicated three times for 20 s each time at 4°C; Heat Systems), or by physical lysis using 425- to 600-μm diameter glass beads (Sigma-Aldrich). For CFSE (Molecular Probes) staining, Lmdd were incubated with 1.5 μM CFSE in PBS at room temperature for 15 min, followed by three washes. Cells were infected with 5–10 CFU of bacteria/cell. Lm-treated cells were added to T cells 4 h after culture at 37°C.
Generation of culture supernatants
Supernatants harvested at the indicated time or 12 h after Lmdd treatment from DC plated at 106 cells/ml in serum-free RPMI 1640 were passed through a 0.2-μm pore size filter and stored at –80°C before use. In some experiments, Lmdd-infected cells were treated with the indicated concentration of cycloheximide 1 h after infection, then cultured overnight before harvesting supernatants. Supernatants were desalted using Econo-Pac desalting columns (Bio-Rad) to remove cycloheximide before addition to T cell cultures.
Microarray assay and RT-PCR
On day 7 of culture, mouse BMDC were enriched using anti-CD11c-conjugated magnetic beads and treated with medium, LPS (1 μg/ml), 10 CFU of HK or live Lmdd/cell for 6 h before extracting total RNA using the Qiagen RNeasy Mini kit. Approximately 10 μg of each sample was used to make labeled probes and was prepared for hybridization to mouse expression set A and B oligonucleotide arrays (Affymetrix), performed at the Microarray Core facility, Dana-Farber Cancer Institute. RNA was extracted and analyzed as described previously (33) using samples from two independent experiments. For RT-PCR, 1 μg of total RNA was used to generate cDNA using the TaqMan RT kit (Applied Biosystems). IFN- and -actin primers were previously described (34): ifnb: forward, 5'-ctggagcagctgaatggaaag; reverse, 5'-cttgaagtccgccctgtaggt; and -actin: forward, 5'-aggtgtgatggtgggaatgg; reverse, 5'-gcctcgtcacccacatagga. Two-step PCR was performed for 35 cycles at 95°C for 10 s and 62°C for 15 s.
Chromatography and mass spectrometry analysis
Supernatants derived from Lmdd-infected BMDC or DC2.4 cells cultured in serum-free RPMI 1640 were harvested, passed through a 0.2-μm pore size filter, concentrated, and separated on a Superdex-200 column. The apparent m.w. of fractions active in inducing CD69 expression on naive C57BL/6 splenocytes were determined by comparing their migrations with those of m.w. standards. Approximately 20 L of serum-free RPMI 1640 supernatant derived from Lmdd-infected DC2.4 cells was separated by sequential High S (Bio-Rad), heparin, hydroxyapatite, and Blue gel (Amersham Biosciences) chromatography. Active fractions were pooled, subjected to tryptic digestion, and analyzed by MALDI-TOF mass spectrometry at the Harvard Medical School Mass Spectoscopy Core facility.
Flow cytometry
Flow cytometry was performed using FACSCalibur and CellQuest software (BD Biosciences) with fluorophore-conjugated Abs to IL-6 (MP5-20F3), IL-12 (C15.6), TNF- (MP6-XT3), I-Ab (AF6-120.1), CD11b (M1/70), CD11c (HL3), CD4 (GK1.5), CD8 (53-6.7), CD25 (clone PC61), CD40 (3/23), CD43 (1B11), CD69 (H1.2F3), CD62L (MEL-14), CD80 (16-10A1), and CD86 (GL1) from BD Pharmingen. Rat anti-mouse IFN mAb (F18) was purchased from Hycult Biotechnology. Rabbit anti-mouse polyclonal Abs against IFN- and IFN- were obtained from PBL Laboratory. For external staining, 2 x 105 cells/microtiter well were washed with FACS buffer (PBS containing 2% heat-inactivated FBS and 0.1% sodium azide) and incubated with an FcR-blocking Ab (BD Pharmingen) for 5 min, then incubated with saturating amounts of mAbs for 30 min at 4°C. For intracellular cytokine staining, cells were treated with 20 μM brefeldin A for the last 12 h of culture or as indicated, resuspended in 50 μl of FACS buffer, and permeabilized using the Fix and Perm kit according to the manufacturer’s protocol (Caltag Laboratories). Saturating amounts of fluorochrome-conjugated Abs were added to the permeabilization buffer, and cells were incubated at room temperature for 15 min. Cells were washed and resuspended in FACS buffer with 1% formaldehyde for analysis.
T cell proliferation and ELISPOT assays
Splenocytes from naive C57BL/6 mice or CD8 T cells from OT-1 transgenic mice were harvested and exposed to 1000 U/ml or the indicated concentration of mouse rIFN- (1.1 x 108 U/mg; PBL Laboratory) or IFN- (1.2 x 107 U/mg; PBL Laboratory) or supernatant derived from Lmdd-infected DC for 3 h and washed. Splenocytes were cultured with the indicated concentrations of anti-CD3 (BD Pharmingen). OT-1 CD8 T cells were cultured with irradiated splenocytes and the indicated concentrations of the cognate OVA peptide SIIFNEKL. Alternatively, splenocytes were directly cultured with the indicated concentration of anti-CD3 in the presence of IFN- or DC supernatant. T cell proliferation was measured by [3H]thymidine incorporation. IFN- production was measured by ELISPOT, as previously described (35), using an immunospot counter (Cellular Technology).
Results
Both HK and live Lmdd are similarly taken up by DC and induce DC maturation
To begin to define the differences in the immune-stimulating effects of live or HK Lmdd, we analyzed the responses of mouse BMDC and the DC cell line DC2.4 to live and HK Lmdd. Both BMDC and DC2.4 cells are myeloid lineage cells expressing CD11c and CD11b, but not B220 or CD8 (Fig. 1A). BMDC internalized both live and HK Lmdd. The number of internalized bacteria was similar 5 h after adding live or HK bacteria, as quantified by flow cytometry (mean fluorescence intensity (MFI), 798 HK vs 771 live Lmdd; Fig. 1B). Actin tails costained with CFSE-labeled bacteria were only evident in live Lmdd-infected DC (data not shown), suggesting that live Lmdd escape from the vacuole to the cytosol, whereas HK Lmdd remain in the vacuole (20). BMDC (Fig. 1C) and DC2.4 cells (data not shown) exposed to live or HK Lmdd for 24 h up-regulated the surface expression of molecules involved in T cell activation, such as CD40, CD80 (B7.1), CD86 (B7.2), and MHC class II (Fig. 1C). Although CD86 and the MHC class II molecule I-Ab were up-regulated to a similar extent, CD40 and B7.1 expression were slightly higher in live Lmdd-treated DC (CD40: MFI, 129 live vs 112 HK; CD80: MFI, 105 live vs 91 HK). Thus, both live and HK Lmdd were similarly internalized by DC, and both induced costimulatory molecules, with only subtle differences in magnitude.
HK and live Lmdd-treated DC differ in the ability to activate T cells
We next examined whether DC pulsed with live or HK Lmdd differentially activate T cells. Live bacteria-pulsed BMDC rapidly stimulated both CD4 and CD8 splenic T cells to express the early activation marker CD69, whereas HK Lmdd-treated DC had less of an effect (Fig. 2A). The reduced ability of HK Lmdd to activate DC capable of inducing CD69 expression on T cells was not due to heat inactivation of bacterial Ags or adjuvants, because Lmdd killed by other means (gentamicin, ultrasound, or glass beads) also did not efficiently activate DC (Fig. 2A). The Lmdd-infected DC2.4 DC line was also able to activate T cells to express CD69, whereas the infected RAW264.7 macrophage cell line had limited ability (Fig. 2A). We next examined whether T cells exposed to Lmdd-infected DC were fully activated. When CFSE-labeled CD8 T cells were cocultured with Lmdd-infected DC, they did not proliferate (Fig. 2, B and C), as determined by CFSE dilution. T cell activation markers, such as CD25 and CD43, were not up-regulated by 48 h (Fig. 2C), although CD62L expression was slightly down-regulated in T cells that had been cocultured with live Lmdd-pulsed DC (Fig. 2C). T cells did not produce IL-2, as determined by intracellular cytokine staining (data not shown). Therefore, T cell activation by Lmdd-infected DC was partial.
Because Lmdd infection introduces a great deal of foreign Ags and triggers DC maturation (Fig. 1), we examined whether activation of CD69 expression was dependent upon TCR engagement. T cells were cocultured with Lmdd-treated BMDC derived from MHC class I- or class II-null mice. CD69 expression on T cells was comparable after stimulation with wild-type or MHC-null DC (Fig. 2D). It was also comparable when T cells were cultured with Lmdd-infected allogeneic BMDC or syngeneic BMDC (Fig. 2D). These data suggest that live Lmdd-infected DC partially activate T cells through a TCR-independent process.
T cell activation by live Lmdd-infected DC is mediated by a secreted soluble factor(s)
Because the activation of T cells by live Lmdd-infected DC is independent of TCR-MHC engagement, we next determined whether it required cell-to-cell contact or was mediated by a soluble factor. Culture supernatant from live, but not HK, Lmdd-treated DC was fully able to induce CD69 expression on T cells (Fig. 3A). Supernatants derived from bacterial cultures without DC or from Lmdd-infected RAW246.7 macrophage cells had little capacity to activate CD69 expression (Fig. 3A), suggesting that the molecules that activate T cells were not produced by bacteria, but were secreted by DC after Lmdd infection. Secretion occurred rapidly within 5 h of infection, reached a peak by 12 h, and gradually declined after 20 h of culture (Fig. 3B). This molecule(s) was synthesized de novo, because treating DC with cycloheximide almost completely inhibited its production (Fig. 3C). Cycloheximide treatment did not affect cell viability, as assessed by forward and side scatter flow cytometry profiles, even at the highest concentration (data not shown).
Live Lmdd-infected DC supernatant enhance Ag-dependent T cell activation
To explore the effects of the soluble factors secreted by Lmdd-infected DC on Ag-dependent T cell activation, splenocytes from naive mice were pulsed with anti-CD3 Ab in the presence or the absence of DC culture supernatant. Lmdd-treated DC supernatant significantly enhanced the numbers of T cells secreting IFN- after anti-CD3 treatment (Fig. 4A). At limiting concentrations of anti-CD3, the dose-response curve was shifted by about 1 log to the left in the presence of Lmdd-treated DC supernatant. HK Lmdd-treated DC supernatant slightly enhanced T cell activation, whereas supernatant from uninfected DC had no effect on promoting Ag-dependent T cell activation (Fig. 4A). In addition, T cell proliferation in response to stimulation by allogeneic splenocytes was significantly enhanced when T cells were continuously cultured in supernatant from Lmdd-infected DC (Fig. 4B). Pre-exposure of T cells to Lmdd-infected DC supernatant also improved T cell proliferation in response to allogeneic splenocytes (Fig. 4B), suggesting that the soluble factors secreted by live Lmdd-infected DC sensitized T cells for subsequent Ag-dependent activation. The continuous presence of supernatant, however, induced more proliferation than when T cells were washed before exposure to allogeneic splenocytes (Fig. 4B).
IFN- expression is induced by live, but not HK, Lmdd-treated DC
To identify the secreted molecules produced by Lmdd-infected DC that activate T cells, we looked for differential production of proinflammatory cytokines, such as IL-12, IL-6, and TNF-, by treated DC (23). HK or live Lmdd infection led to a comparable increase in IL-6 production (Fig. 5A). IL-12 and TNF- production by live Lmdd-treated DC was slightly higher than that by HK Lmdd-treated DC (Fig. 5, B and C, and data not shown). However, Lmdd-infected DC secreted much less TNF- than infected RAW264.7 cells (Fig. 5C). Although Escherichia coli-infected DC have been reported to produce IL-2 (36), no IL-2 was induced after Lmdd infection (data not shown). Because the production of these cytokines was similar after HK or live Lmdd infection, none of these cytokines is probably responsible for the difference in T cell activation.
To identify the soluble DC factor capable of inducing CD69 expression and sensitizing T cell activation, Lmdd-infected DC supernatant was separated by chromatography, testing fractions for their ability to up-regulate CD69 expression on T cells. The apparent Mr of the soluble factor was determined by Superdex 200 gel filtration to be 24–28 kDa (data not shown). Lmdd-infected DC supernatant was separated through sequential cation exchange High S, heparin, hydroxyapatite, and Blue gel columns. The final active fraction was subjected to tryptic digestion and MALDI-TOF mass spectroscopy analysis, and four peptides with sequences matching IFN- were identified (data not shown). This suggested that IFN- might be the soluble factor responsible for CD69 up-regulation. This result was unexpected, because IFN- is thought to be produced mostly by pDC.
To determine whether IFN- is the sought-after soluble factor produced by Lmdd-infected DC, we first looked by microarray analysis at whether mDC differentially up-regulate IFN- mRNA expression after Lm infection (Table I). CD11c+ BMDC were treated for 6 h with medium, LPS, or HK or live Lmdd before isolating mRNA. LPS-treated DC had limited capacity to activate T cells to express CD69 (data not shown). Twenty-seven genes were up-regulated by at least 10-fold after live Lmdd infection compared with untreated DC. Sixteen of these were also up-regulated at least 10-fold by treatment with HK Lmdd. The gene that showed the greatest modulation, however, was IFN-, whose expression was increased 67-fold. Moreover, IFN- expression was not enhanced by HK Lmdd and was only enhanced 4-fold by LPS. By semiquantitative RT-PCR, no IFN- mRNA was detected in untreated BMDC, but expression was up-regulated in Lmdd-infected BMDC (Fig. 6C). Samples from LPS- and HK Lmdd-treated DC had detectable IFN- expression, but substantially less than samples from Lmdd-infected DC. These data were also confirmed by quantitative real-time PCR (data not shown). Other genes that were up-regulated differentially by live Lmdd compared with both HK Lmdd and LPS were IFN- genes 2 (increased 19-fold) and 5 (increased 8-fold). IFN- expression was up-regulated in BMDC to a similar extent by live Lmdd (6-fold) or LPS (8-fold), but only 2-fold by HK Lmdd treatment. In addition, a number of IFN-inducible genes and cytokine and chemokine genes and receptors as well as some genes with unknown function were up-regulated substantially by 6 h, but they were also up-regulated in HK Lmdd-treated and/or LPS-treated DC (Table I). The inducible IFN-regulated factor 7 (IRF7) gene was significantly up-regulated by treatment with live Lmdd (13-fold) or LPS (15-fold) and less so by HK Lmdd (4.5-fold; Table I), indicating that the positive feedback loop of type I IFN gene expression was rapidly activated by all these treatments (37, 38). The low levels of IFNs induced by LPS or HK Lmdd appeared to be sufficient to activate a wide array of IFN-inducible genes. However, although IRF3 mRNA remained unchanged, IRF3 protein translocated from the cytoplasm to the nucleus only in DC treated with live Lmdd, as determined by subcellular fractionation and immunoblotting (data not shown). IRF3 nuclear translocation is the key to inducing IFN- expression (39).
Discussion
It has long been known that live Lm induce protective immunity, whereas killed bacteria fail to do so (13, 18). Unveiling the mechanism by which immune cells differentially respond to live vs dead Lm is important for understanding innate and adaptive immunity, and ultimately will help to design better vaccines against cancer and infectious diseases. Because DC are the key to priming immune responses, in this study we focused on differential effects of live and HK bacteria on DC activation and priming of T cells. We found that although live and HK Lm similarly cause DC to mature, as measured by up-regulating cell surface costimulatory molecules and MHC class II, only live bacteria-infected DC are able to activate CD69 expression on T cells efficiently and prime them for subsequent activation by the TCR. Using protein chemistry and microarray gene expression analysis, we identified the type I IFN, IFN-, as a soluble factor rapidly produced in large amounts (3000 U/106 cells) by BMDC upon infection by live Listeria, but not HK bacteria. Moreover, we were able to show that only high concentrations of type I IFNs are able to prime T cell activation. We therefore hypothesize that the exceptionally strong and protective immune response to Listeria is related to the amplification of T cell priming that occurs in the presence of high local concentrations of type I IFN secreted by infected DC.
The copious production of type I IFN in our studies was unanticipated, because our experiments were performed using mDC. Conventional wisdom holds that the major type I IFN-producing DC is the B220+CD11b– pDC (11). However, in our experiments the source of DC was either the B220–CD11b+ mDC cell line DC2.4 or BMDC that had been cultured in GM-CSF and IL-4 and selected for CD11c expression. These cultured BMDC are mDC, contain <1% B220+ cells, and are >98% CD11b+ (Fig. 1A). Furthermore, Lmdd infection did not alter the BMDC expression of cell surface markers that distinguish mDC and pDC (data not shown). Although the DC2.4 cell line is fully myeloid, we were concerned that contamination of our BMDC with a few pDC might account for the IFN production we measured in the BMDC cultures. However, when BMDC were cultured in the presence of Flt3 ligand to generate cultures enriched in pDC (30% B220+), the bioactivity of the Lmdd-infected DC-conditioned medium was not enhanced (Fig. 6B). These data therefore suggest that mDC produce high levels of type I IFN when infected with Listeria. Production of type I IFN by mDC was also documented in a recent study that showed that mDC produce large amounts of type I IFN after infection with viruses such as lymphocytic choriomeningitis virus (12).
Type I IFNs have multiple biological activities and play important roles in bridging innate immune responses and adaptive immunity (41). Both IFN- and IFN- trigger CD69 expression on T cells, but Lm infection triggered more IFN- than IFN- expression in mDCs. Ab-blocking experiments also suggested that most of the biological effect could be attributed to IFN-. In this study we also demonstrated that IFN- produced by Lm-infected DC acts as a commitment factor to decrease the Ag response threshold of T cells and enhance their priming. The initial exposure to IFN- partially activates naive T cells, preparing them for subsequent Ag-specific activation.
Stimulation of type I IFN production by mDC and their priming of T cell CD69 expression and sensitization for TCR stimulation require Lm escape from the phagolysosome. HK bacteria remain in the phagolysosome. Using bacterial mutants lacking or with inducible expression of LLO, which is required for cytosolic invasion, we also found that the production of IFN- and T cell activation by mDC correlate with the ability to escape the phagocytic vacuole. Portnoy and colleagues (20) previously reported production of IFN- by Lm-infected macrophages via a mechanism that requires bacterial cytosolic invasion. They postulated a bacterial cytosolic sensor that signals the presence of cytosolic bacteria and triggers the secretion of IFN- by macrophages. Our results suggest that a similar sensor and pathway are triggered by intracytoplasmic infection of mDC. The sensor remains to be identified. LPS activates a TLR4, MyD88-independent signaling pathway that induces IRF3 translocation and IFN- production via the Toll/IL-1R domain-containing adaptor (42, 43, 44). However, we found that LPS has a very limited capacity to stimulate BMDC to produce IFN- compared with live Lmdd, and LPS-activated DC do not efficiently stimulate T cells to express CD69. Supernatants from Lmdd-infected BMDC derived from mice deficient in MyD88, a key signaling molecule for TLR engagement, produce comparable amounts of IFN- (data not shown). It is, moreover, unlikely that TLR family proteins are responsible, because these receptors are displayed on the cell surface and within endosomes, but not in the cytosol. The nucleotide-binding oligomerization domain and Nacht, leucine-rich repeat, and pyrin domain-containing protein family proteins that are present in the cytosol and are able to sense Gram-positive and Gram-negative bacterial cell wall components are attractive candidates for the unknown sensor (45, 46, 47).
In this study we found that pulsing bone marrow-derived mDC with either HK or live Lmdd induces DC to up-regulate costimulatory molecules on their surface and secrete proinflammatory cytokines, such as TNF-, IL-6, and IL-12. DC maturation and induction of proinflammatory cytokines were probably stimulated by engagement of TLR receptors by both HK and live Lmdd. Because the differential effects of live and HK bacteria were also evident when bacteria were killed by other means (such as antibiotic treatment) that do not cause denaturation, differences in TLR engagement of pathogenic patterning molecules in the bacteria were probably minimal. It is, therefore, not surprising that dead Lm also induce DC maturation. Although slightly more CD40 and CD80 were detected on the surface of BMDC infected with live Lmdd than HK Lmdd, the expression of other cell surface markers (class II MHC and CD86) and that of inflammatory cytokine production were comparable. Therefore, these factors are unlikely to explain the large difference in T cell activation we observed. However, the difference in type I IFN expression was dramatic, and Abs to type I IFNs could abrogate the effect of Lmdd-infected BMDC supernatants on T cells.
Our conclusions differ from those in a recent paper that compared GM-CSF-cultured BMDC maturation, DC inflammatory cytokine production, and stimulation of T cells to produce IFN- in response to infection with wild-type and LLO-deficient Lm (23). That study also found that bacterial cytosolic invasion was critical for T cell activation, but found more substantial differences in costimulatory molecule expression and proinflammatory cytokine expression, which they interpreted as the distinguishing feature. Our use of a replication-defective Lmdd strain may account for differences between their study and ours (25). In fact, they had twice as much infection after 4 h using wild-type as hly– bacteria. Another difference is that the other study examined DC maturation 18 h after Lm infection, whereas we observed DC maturation after 24 h. Typically, costimulatory molecule up-regulation occurs relatively slowly compared with IFN- induction, requiring 16–24 h (data not shown). Type I IFN production in response to bacterial invasion may accelerate DC maturation because it induces a positive loop of type I IFNs and IFN-inducible gene up-regulation (38) that enhances Ag presentation function and DC maturation (48). However, DC matured more slowly via TLR engagement may reach the same final state for efficient Ag presentation by 24 h. Accelerated DC maturation may lead to a more rapid and effective immune response. A final difference is that the myeloid DC used in the other study were generated from bone marrow by culture in GM-CSF, whereas we generated myeloid DC by culture in both IL-4 and GM-CSF. The combination of subtle differences in cytokines, timing, and bacterial replication may account for the different conclusions. However, which in vitro condition more faithfully recapitulates in vivo conditions is impossible to predict.
Both live and HK bacteria induce the rapid expression of multiple IFN-responsive genes in BMDC within 6 h, the time after Lmdd exposure at which we performed our microarray analysis. The downstream activation of these genes, many of which are also up-regulated by LPS engagement of TLR4, is probably triggered by low levels of IFNs generated after TLR signaling. Our results are similar to a gene expression analysis of Lm- or LPS-treated bone marrow-derived macrophages, which found that both bacterial invasion and TLR engagement were able to activate many IFN-responsive genes (49). However, high concentrations of type I IFN (250 U/ml IFN-, 5000 U/ml IFN- to activate CD69 on 50% of T cells) are needed to induce CD69 on T cells and prime T cells for antigenic exposure. These high concentrations of type I IFN decrease the Ag response threshold of naive T cells by 1 log. Therefore, weak signals that might otherwise be ignored or produce an ineffective, tolerogenic, or even suppressive regulatory immune response can be activated in the presence of high levels of type I IFN. The partial TCR-independent activation of T cells by high concentrations of type I IFN may also play a part in the bystander T cell apoptosis and lymphocyte depletion observed after Lm infection, because these partially activated T cells are prone to spontaneous apoptosis (data not shown) (50). Although removing nonspecific T cells may make room for expansion of Lm-specific T cells, it might also interfere with maintaining effective memory.
Our results and those reported by Portnoy and colleagues (20) suggest that in addition to TLR engagement, there is a second alarm system in APCs, such as DC and macrophages, that is activated when bacteria invade the cytosol. This alarm triggers the nuclear translocation of IRF3 and the production of copious quantities of type I IFNs that enhance T cell sensitivity to antigenic stimulation. This alarm system may be triggered by intracellular pathogens to enlist the aid of the cells (T cells) most capable of eliminating intracellular pathogens. The induction of type I IFN production by mDC after infection with lymphocytic choriomeningitis virus and a mutant strain of influenza (12) suggests that this second alarm might also be triggered by some viral infections. It will be of interest to determine whether other viruses and cytosolic pathogens also trigger exuberant type I IFN expression. Because T cells are not important for eliminating extracellular pathogens, it makes sense that this response would not be elicited by most bacteria and would not be triggered by cell surface receptors, like the TLR system. Our data also suggest that mDC produce much more IFN- than macrophages after live Lmdd infection, because supernatant from the infected RAW264.7 macrophage cell line barely activates T cells to express CD69 (Figs. 2A and 3A). Therefore, this second alarm system may function more effectively in DC than in macrophages.
This specialized type I IFN danger response may explain why Lm is such an effective vector for priming T cell responses to Ags. Because Lm causes serious disease in immunosuppressed individuals, pregnant women, and newborns, most investigators developing Lm as a vaccine vector agree that the bacterium must be attenuated before human use is contemplated (51). Our results suggest that attenuated bacteria unable to escape from the phagolysosome may be impaired in T cell priming and lead to ineffective vaccines. In fact, the vaccine strains that appear the most promising are able to invade the cytosol. Screening for type I IFN production may be a useful tool for predicting which attenuated bacteria are likely to be effective vaccine vectors.
Although this study suggests that type I IFN production in response to Lm cytosolic invasion should send a strong signal to enhance T cell immunity, mice deficient in the common type I IFNR are actually better able to handle Lm challenge (34, 52, 53). This is unexpected, because type I IFNs generally protect against other types of infection, especially by viruses. In fact, the harmful effect of type I IFNs may be peculiar to Lm infection (52, 54). Carrero et al. (52, 54) found that LLO, the bacterial pore-forming protein, acts as a bacterial toxin to induce T cell apoptosis, and that partially or fully activated T cells are particularly prone to LLO-induced apoptosis. Type I IFNs probably accelerate this process, because they induce partial activation of nonspecific T cells and facilitate full activation of specific T cells. This idiosyncratic effect of LLO on T cells and the induction of type I IFN need to be considered in developing Listeria-based vaccine vectors. Because of the pleiotropic effects of IFNs on different immune cells, however, the resistance of mice null for the type I IFN receptor to Lm may be due to more than one factor.
Acknowledgments
We thank Lianfa Shi, Manjunath Swamy, Howell Moffat, Pascale Cossart, and Jay Berzofsky for useful advice, and Fred Frankel and Darren Higgins for providing the strains of Lm used in this study.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work was supported by National Institutes of Health Grant AI056922 (to J.L.) and T32HL066987 (to A.S.).
2 Current address: Division of Infectious Diseases, Tufts University School of Veterinary Medicine, North Grafton, MA 01536.
3 Address correspondence and reprint requests to Dr. Judy Lieberman, CBR Institute for Biomedical Research, 200 Longwood Avenue, Boston, MA 02115. E-mail address: lieberman@cbr.med.harvard.edu
4 Abbreviations used in this paper: Lm, Listeria monocytogenes; DC, dendritic cell; BMDC, bone marrow-derived DC; D-Ala, D-alanine; HK, heat killed; IPTG, isopropyl--D-thiogalactoside; IRF, IFN-regulated factor; LLO, listeriolysin O; mDC, myeloid DC; MFI, mean fluorescence intensity; pDC, plasmacytoid DC.
Received for publication January 7, 2005. Accepted for publication April 29, 2005.
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