TLR4 and Toll-IL-1 Receptor Domain-Containing Adapter-Inducing IFN-, but Not MyD88, Regulate Escherichia coli-Induced Dendritic Cell Maturat
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免疫学杂志 2005年第14期
Abstract
Dendritic cells (DC) are short-lived, professional APCs that play a central role in the generation of adaptive immune responses. Induction of efficient immune responses is dependent on how long DCs survive in the host. Therefore, the regulation of DC apoptosis in vivo during infection remains an important question that requires further investigation. The impact of Escherichia coli bacteremia on DCs has never been analyzed. We show here that i.v. or i.p. administration of live or heat-killed E. coli in mice induces splenic DC migration, maturation, and apoptosis. We further characterize which TLR and Toll-IL-1R (TIR)-containing adaptor molecules regulate these processes in vivo. In this model, DC maturation is impaired in TLR2–/–, TLR4–/– and TIR domain-containing adapter-inducing IFN- (TRIF)–/– mice. In contrast, DC apoptosis is reduced only in TLR4–/– and TRIF–/– mice. As expected, DC apoptosis induced by the TLR4 ligand LPS is also abolished in these mice. Injection of the TLR9 ligand CpG-oligodeoxynucleotide (synthetic bacterial DNA) induces DC migration and maturation, but only modest DC apoptosis when compared with LPS and E. coli. Together, these results suggest that E. coli bacteremia directly impacts on DC maturation and survival in vivo through a TLR4-TRIF-dependent signaling pathway.
Introduction
Dendritic cells (DCs)3 are professional APCs critical for bridging innate and adaptive immune responses (1). Immature DCs express germline-encoded receptors that recognize pathogen-associated molecular patterns (PAMPs). These receptors allow them to rapidly detect microbial infections. During the past few years, the TLR family has emerged as a major group of signaling receptors for PAMPs (2). Most mammalian species have 10–15 TLRs. TLRs detect multiple PAMPs, including LPS, the major glycolipidic component of Gram-negative bacteria (detected by TLR4), bacterial lipoproteins and lipoteichoic acids (detected by TLR2) and the unmethylated CpG DNA of bacteria and viruses (detected by TLR9). TLRs recruit adaptor molecules that subsequently activate downstream signaling pathways and NF-B. Four adaptor molecules have been identified: MyD88 (3); Toll-IL-1R (TIR) domain-containing adaptor protein (4); TIR domain-containing adapter-inducing IFN- (TRIF)/TICAM-1 (TIR domain-containing adaptor-inducing IFN-) (5); and TRIF-related adapter molecule (TRAM)/TICAM-2 (6). Although the role of the various adaptors in the activation of specific signaling pathways still remains controversial, recent studies suggest that MyD88 and TRIF regulate independent pathways. In this model, TIR domain-containing adaptor protein is implicated in MyD88-dependant signaling, and TRAM (which is highly homologous to TIR) domain-containing adaptor protein) appears to function as a bridge between TLR4 and TRIF (7).
DC activation after TLR activation is characterized by several events: DCs 1) migrate to the T cell area, 2) lose their phagocytic capacity, 3) secrete proinflammatory cytokines, and 4) achieve a mature phenotype. During maturation, DCs increase their ability to present Ags and prime T cells by up-regulating their cell surface MHC II and CD40-, CD80-, and CD86-costimulatory molecules (1). This process is critical for mounting efficient innate immune responses to pathogens as well as for the development of T cell- and B cell-mediated adaptive responses. In addition, it has recently been reported that TLR activation can promote apoptosis of various cell lines (8, 9).
Sepsis is a major health problem (10, 11). During the onset of bacteremia in humans or rodents, massive apoptosis is observed inside lymphoid organs (11). Apoptosis can function as a highly efficient host mechanism to both limit the spread of intracellular pathogens and potentially down-regulate immune responses. In contrast, it is possible that pathogen-mediated induction of apoptosis is a strategy to evade immune responses. Despite the critical importance of DCs in the initiation and regulation of Th1 immune responses against bacteria (12, 13), very little is known on how bacteremia affects DCs in vivo.
Peritonitis continues to be a leading cause of morbidity and mortality (14, 15, 16). Although the overall mortality of sepsis is 35% (17), abdominal sepsis is associated with mortality rates of up to 80% (18). Although various bacteria have been identified as causative organisms in peritonitis, Escherichia coli belongs to the most common bacteria recovered from peritonitis (up to 60%; Ref. 19) and is the second most frequent bacterium to cause sepsis (up to 20%; Ref. 20).
In this work, we analyze whether E. coli bacteremia induce DC migration, maturation, and apoptosis in vivo. We further determine which TLRs and adaptor molecules are required for these effects on DC. Several components of E. coli have been reported to impact DCs. LPS induces DC activation (21) and apoptosis (22) in vivo. Bacterial DNA and its synthetic form CpG-oligodeoxynucleotides (CpG-ODN) also induce DC activation in vivo (23). Therefore, we analyzed the ability of E. coli bacteremia to induce DC maturation and apoptosis in vivo and compare them with those obtained after LPS and CpG-ODN administrations.
Materials and Methods
Mice and reagents
Female BALB/c and C57BL/6 mice (6–8 wk old) were purchased from Harlan Sprague Dawley. MyD88–/– (3), TLR2–/– (24), and TLR4–/– (25) mice on the BALB/c background and TRIF–/– mice on the C57BL/6 background (Beutler, Scripps Research Institute; Ref. 5) were bred in the specific pathogen-free animal facility of the Institute of Molecular and Cellular Pharmacology, Nice-Sophia Antipolis (Valbonne, France). The maintenance and care of mice complied with the guidelines of the Institut National de la Santé et de la Recherche Médicale/Centre National de Recherche Scientifique/University of Nice-Sophia Antipolis Ethic Committee for the use of laboratory animals. CpG-ODN 1826 was obtained from Coley Pharmaceutical Group. Highly purified LPS (E. coli serotype 0111:B4) was purchased from Invivogen. E. coli (strain K514) was grown to stationary phase (8–10 h) in Luria-Bertani medium and further washed (2500 x g, 10 min, 4°C) three times in PBS before inoculation into mice. Heat-killed (HK) E. coli were obtained by a 60-min incubation at 70°C. We checked that the HK bacteria did not grow after inoculation of an overnight culture in Luria-Bertani medium.
In vivo treatment
Mice were given injections i.v. into the lateral tail vein with indicated doses of LPS, CpG-ODN, or E. coli in PBS. Control animals were injected with the same volume of PBS.
MLR
Spleens were cut into very small pieces and incubated (37°C, 30 min) into HBSS (Life Technologies) containing 4000 U/ml collagenase IV (Roche) and 0.1 mg/ml DNase I fraction IX (Sigma-Aldrich Chimie SARL). CD4+ T cells from the spleen were purified by negative depletion of CD8+, B220+, CD11b+, and MHC II+ cells using sheep anti-rat Ig-coated Dynabeads (Dynal). The indicated numbers of gamma-irradiated (3000 rads) spleen cells were incubated with 3 x 105 allogeneic CD4+ T cells in DMEM supplemented with L-glutamine (2 mM), heat-inactivated FCS (10%), 2-ME (5 x 10–5 M), penicillin (100 μg/ml), and streptomycin (100 U/ml) in U-bottom 96-well plates. Cell proliferation after 48 h of culture was determined by [3H]thymidine incorporation.
Cytofluorometric analysis
Spleen cells were analyzed by flow cytometry with a FACSCalibur cytofluorometer (BD Biosciences). The cells were first preincubated with saturating doses of 2.4G2 (a rat anti-mouse Fc receptor mAb; American Type Culture Collection) for 10 min to prevent Ab binding to Fc receptor and then labeled with PE-coupled N418 (anti-CD11c), APC-coupled 53-6.7 (anti-CD8), and fluorescein (FITC)-coupled Abs including L3T4 (anti-CD4), 3.23 (anti-CD40), 16–10A1 (anti-CD80), GL1 (anti-CD86), or 2G9 Ab (anti-IA/IE). All mAbs were purchased from BD Biosciences. Cells were gated according to size and scatter to eliminate dead cells and debris from analysis.
Confocal microscopy
Tissues were fixed for 3 h at 4°C in 1% paraformaldehyde (pH 7.0), washed in PBS, incubated overnight at 4°C in a solution of 30% sucrose under agitation, and washed again in PBS. Tissues were embedded in the Tissue-Tek OCT compound (Sakura) and frozen at –80°C, and 10-μm cryostat sections were prepared. Tissues sections were incubated in 100 mM Tris-HCl, pH 8.0, to eliminate OCT and then in a saturated solution containing 10% normal serum in 100 mM Tris-HCl, pH 8.0. Avidin-biotin sites were saturated using an avidin-biotin blocking kit (Vector). Tissues sections were incubated with any of the following mAbs: Alexa Fluor 488 phalloidin (Molecular Probes); APC-coupled N418 (anti-CD11c); GL1 (anti-CD86); 11-26c.2a (anti-IgD); or 30-H12 (anti-CD90.2, Thy-1.2; BD Biosciences) or MOMA-1 (BMA Biomedicals) in 100 mM Tris-HCl, pH 8.0. Uncoupled mAbs were stained with Alexa Fluor 568-coupled goat anti-rat IgG (Molecular Probes). Slides were mounted in DAKO medium (DakoCytomation). Labeled cells were visualized under a Leica laser scanning confocal microscope (TCS-SP) equipped with a DM-IRBE inverted microscope and an argon-krypton laser.
Immunohistochemistry
Immunohistochemical techniques used in this study were previously described (26). Briefly, spleens were fixed in Immunohistofix (Aphase), embedded in Immunohistowax (Aphase), and sectioned at 3–6 μm. Tissues sections were then treated with 1% blocking reagent (Boehringer) to saturate nonspecific binding sites. The endogenous peroxidase activity was neutralized by adding 3% H2O2 in PBS for 30 min. The slides were further treated to visualize apoptotic cells using TUNEL (Boehringer, Mannheim) and costaining with alkaline phosphatase substrates (SK-5300, blue precipitate; Vector Laboratories). A blocking kit (Vector Laboratories) was used to saturate the excess of biotin. Slides were incubated with the N418-biotinylated Ab (10 μg/ml in PBS-0.5% blocking reagent), washed, and then incubated with avidin-biotin-peroxidase complexes (Vectastain ABC kit) revealed with a solution of AEC tablets (Sigma-Aldrich). Digitalized images were captured using a charge-coupled device color camera (Ikegami Tsushinki).
Results
Live and HK E. coli induce DC migration and maturation in vivo
To model the potential effect of bacteremia on DCs in lymphoid organs, live or HK E. coli (5 x 107 bacteria) were injected i.v. into BALB/c mice. This treatment induced the migration of splenic DCs (CD11chigh cells) to T cell zones as shown by confocal microscopy on spleen sections from wild-type (wt) BALB/c mice (Fig. 1A and data not shown for HK E. coli). This result parallels previous findings using LPS (21). DCs migrate from red pulp and marginal zone to T cell-rich areas surrounding the central arteriole within a 60-h time frame. Marginal metallophilic macrophages (MOMA-1+ cells) are localized in the inner part of the marginal zone and form a discontinued ring delineating the border between red pulp and white pulp. DC migration to T cell zones occurs through areas of the marginal zone containing fewer MOMA-1+ cells (Fig. 1, A and B). Similar migration profiles are observed after i.v. administration of 105–108 bacteria, as well as with 50 μg of the TLR9 ligand CpG-ODN (data not shown). We also observed that CD11c+ cells, localized in the T cell area, up-regulate the expression of CD86 on inoculation of HK and live E. coli and LPS (Fig. 1C) as well as with HK and CpG-ODN (data not shown). In addition, high levels of constitutive CD86 were expressed on the CD11c– MOMA-1+ macrophage population in the marginal zone, as shown previously (27). Similar results with regard to DC migration and maturation were also seen in C57BL/6 mice (data not shown).
To confirm results observed using immunohistochemistry, flow cytometry was used to analyze cell surface expression of MHC II, CD40, CD80, and CD86 on splenic CD11chigh cells 9 h after administration of HK and live E. coli, LPS (50 μg), and CpG-ODN (50 μg) (Fig. 1D). The effects observed on DC maturation were similar whether E. coli were administered through the i.p. or i.v. route, and the kinetics was similar to that seen after i.v injection of LPS or CpG with peak expression of maturation markers observed at 6–12 h postinjection. Moreover, all CD11chigh DC subsets were affected (Fig. 1E for CD86; data not shown for CD40, CD80, and MHC II). Interestingly, because the administration of live and HK E. coli induces comparable up-regulation of costimulatory molecules on DCs, this suggests that structural components of E. coli are sufficient to induce complete DC maturation.
It has been recently found that commercial LPS preparations are frequently contaminated by lipoprotein and other bacterial components that interact with TLR2 (28). For these reasons, we injected highly purified E. coli LPS in all experiments. In addition, to prevent mice mortality after E. coli injections, we immunized them with an E. coli strain (K514) exhibiting a low virulence. However, short term experiments monitoring DC migration and maturation after injection of a virulent strain (018:K1) gave similar results (data not shown).
E. coli-induced DC maturation is dependent on TLR2, TLR4, and TRIF in vivo
To characterize the receptors and the intracellular signaling pathways involved in E. coli-induced DC migration and maturation, we inoculated wt, TLR2–/–, TLR4–/–, MyD88–/–, and TRIF–/– mice with 5 x 107 live E. coli, LPS (50 μg), and CpG-ODN (50 μg) (Fig. 2). Because DCs from wt BALB/c or C57BL/6 immunized recipients undergo comparable migration and maturation (data not shown), only data obtained with wt BALB/c mice are presented.
Discussion
Systemic LPS administration has been used in previous studies to visualize DC maturation and apoptosis in vivo (21, 22). However, to our knowledge, these data have never been validated in a more physiological model, such as a bacterial infection. In this study, we demonstrate that E. coli bacteremia induces DC migration, maturation, and apoptosis with magnitudes and kinetics similar to that of LPS in vivo. Although all DC subsets mature in response to E. coli or LPS, apoptosis preferentially affects CD8+ DCs that are described to preferentially induce Th1 responses (12, 13). It is conceivable that CD8+ DCs are more sensitive to apoptosis because they express higher levels of TLR4 (30). Along similar lines, preferential apoptosis of CD8+ DCs has recently been described in lymph nodes during polymicrobial sepsis (31). Thus, DC apoptosis can be a general consequence of bacterial infection. In the model of polymicrobial sepsis (31), DC maturation is not observed before DC apoptosis.
As a possible consequence of DC apoptosis, we observed that E. coli bacteremia decreased the ability of total spleen cells to stimulate naive T cells during MLR. It is possible that the DC apoptosis that we observed could be a contributing factor to the immunosuppression associated with sepsis (32, 33). In agreement with this idea, reduction in DC numbers has been correlated in several systems with a reduced ability to mount T cell responses in vivo (13, 21). Moreover, DC loss has been associated with a decreased ability of recipient mice to produce systemic IL-12 in response to LPS injection (34). This suggests, perhaps not surprisingly, that massive DC apoptosis can specifically impair antibacterial Th1 responses.
The impact of TLRs, TRIF, and MyD88 in E. coli, LPS, and CpG-ODN-induced DC maturation and apoptosis is summarized in Table I. E. coli-induced DC maturation is reduced in TLR2–/– and TLR4–/– mice, demonstrating that both TLRs are important PAMP receptors for this pathogen. It also suggests that TLR2 and TLR4 participate in detection and control of E. coli infection. The requirement for TRIF in LPS-induced DC maturation has been described previously (29), but the role of TRIFs in E. coli bacteremia-induced DC maturation is a novel observation. This suggests a possible role for TRIF in the development of adaptive immune response against bacteria.
Although LPS-mediated DC activation requires signals emanating from TLR4 and the downstream binding of TIRAP, TRAM, TRIF, and MyD88 adaptor proteins (2), little has been described regarding the roles of these receptors and signaling molecules involved in LPS-induced DC apoptosis in vivo. However, it is known that the death receptor Fas is not involved in this process (21). We show here that LPS-induced splenic DC apoptosis in vivo depends on TLR4, TRIF, and MyD88 adaptor proteins. In addition, we demonstrate that the TLR4-mediated TRIF-dependent signaling pathway contributes to the control of DC apoptosis during E. coli bacteremia. Several recent in vitro studies using macrophages, bone marrow-derived DCs, and other cell lines show a direct role of TLR4 (8, 9), MyD88 (35), and TRIF (9) on apoptosis. Another in vitro study using TLR3 ligands suggests that TRIF regulates three distinct signaling pathways leading to the induction of the IFN- promoter, NF-B activation, and apoptosis via a FADD/caspase-8-dependent pathway (36).
CpG-ODN injection promoted DC migration and maturation but induced less DC apoptosis compared with LPS and E. coli; this was seen by several different assays (Figs. 3A, 5C, and 5D). CpG-ODN induced DC apoptosis is completely abrogated by MyD88 deficiency. Our in vivo results are in agreement with a recent work (37) showing that in vitro stimulation with LPS- but not CpG-induced apoptosis of bone marrow-derived DC. This study concluded that the ability of CpG to up-regulate Bcl-2, a potent anti-apoptotic molecule, counteracted the induction of programmed cell death of bone marrow-derived DC. Thus, the weak ability of CpG-ODN to induced DC apoptosis suggests a limited implication of bacterial DNA in E. coli-induced DC apoptosis and may explain its important role as a potent adjuvant in vivo.
Taken together, our results provide new insight into how Gram-negative bacteremia modulates DCs function and survival. Coevolution between bacteria and host immune systems has selected the emergence of PAMPs receptors to initiate innate immunity to these pathogens and control infection. However, during septicemia, bacteria subvert this detection system and induce massive DC apoptosis. This phenomenon most likely impairs or reduces the initiation and/or development of adaptive immune responses.
Acknowledgments
We thank Drs. P. M. Dubois and C. A. Benedict for criticism of this manuscript.
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 grants from the Belgian program on Interuniversity Poles of Attraction (to P.A.I.) initiated by the Belgian State, Prime Minister’s Office, Science Policy Programming, the Fonds National de la Recherche Scientifique (Crédit aux chercheurs, Belgium), the Ministère de l’Education Nationale, de la Recherche et de l’Enseignement Supérieur (France), and the Institut National de la Santé et de la Recherche Médicale. M.M. is Senior Research Associated from Fonds National de la Recherche Scientifique. E.M. is Research Associated from Fonds National de la Recherche Scientifique.
2 Address correspondence and reprint requests to Dr. Eric Muraille, Laboratoire de Parasitologie, Faculté de Médecine, Route de Lennik 808, 1070 Bruxelles, Université Libre de Bruxelles, Brussels, Belgium. E-mail address: emuraille@hotmail.com
3 Abbreviations used in this paper: DC, dendritic cell; PAMP, pathogen-associated molecular pattern; TIR, Toll-IL-1R; TRIF, TIR-domain-containing adapter-inducing interferon-; TRAM, TRIF-related adapter molecule; CpG-ODN, CpG-oligodeoxynucleotide; HK, heat killed; wt, wild type; TICAM, TIR domain-containing adaptor molecule.
Received for publication November 10, 2004. Accepted for publication April 27, 2005.
References
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Dendritic cells (DC) are short-lived, professional APCs that play a central role in the generation of adaptive immune responses. Induction of efficient immune responses is dependent on how long DCs survive in the host. Therefore, the regulation of DC apoptosis in vivo during infection remains an important question that requires further investigation. The impact of Escherichia coli bacteremia on DCs has never been analyzed. We show here that i.v. or i.p. administration of live or heat-killed E. coli in mice induces splenic DC migration, maturation, and apoptosis. We further characterize which TLR and Toll-IL-1R (TIR)-containing adaptor molecules regulate these processes in vivo. In this model, DC maturation is impaired in TLR2–/–, TLR4–/– and TIR domain-containing adapter-inducing IFN- (TRIF)–/– mice. In contrast, DC apoptosis is reduced only in TLR4–/– and TRIF–/– mice. As expected, DC apoptosis induced by the TLR4 ligand LPS is also abolished in these mice. Injection of the TLR9 ligand CpG-oligodeoxynucleotide (synthetic bacterial DNA) induces DC migration and maturation, but only modest DC apoptosis when compared with LPS and E. coli. Together, these results suggest that E. coli bacteremia directly impacts on DC maturation and survival in vivo through a TLR4-TRIF-dependent signaling pathway.
Introduction
Dendritic cells (DCs)3 are professional APCs critical for bridging innate and adaptive immune responses (1). Immature DCs express germline-encoded receptors that recognize pathogen-associated molecular patterns (PAMPs). These receptors allow them to rapidly detect microbial infections. During the past few years, the TLR family has emerged as a major group of signaling receptors for PAMPs (2). Most mammalian species have 10–15 TLRs. TLRs detect multiple PAMPs, including LPS, the major glycolipidic component of Gram-negative bacteria (detected by TLR4), bacterial lipoproteins and lipoteichoic acids (detected by TLR2) and the unmethylated CpG DNA of bacteria and viruses (detected by TLR9). TLRs recruit adaptor molecules that subsequently activate downstream signaling pathways and NF-B. Four adaptor molecules have been identified: MyD88 (3); Toll-IL-1R (TIR) domain-containing adaptor protein (4); TIR domain-containing adapter-inducing IFN- (TRIF)/TICAM-1 (TIR domain-containing adaptor-inducing IFN-) (5); and TRIF-related adapter molecule (TRAM)/TICAM-2 (6). Although the role of the various adaptors in the activation of specific signaling pathways still remains controversial, recent studies suggest that MyD88 and TRIF regulate independent pathways. In this model, TIR domain-containing adaptor protein is implicated in MyD88-dependant signaling, and TRAM (which is highly homologous to TIR) domain-containing adaptor protein) appears to function as a bridge between TLR4 and TRIF (7).
DC activation after TLR activation is characterized by several events: DCs 1) migrate to the T cell area, 2) lose their phagocytic capacity, 3) secrete proinflammatory cytokines, and 4) achieve a mature phenotype. During maturation, DCs increase their ability to present Ags and prime T cells by up-regulating their cell surface MHC II and CD40-, CD80-, and CD86-costimulatory molecules (1). This process is critical for mounting efficient innate immune responses to pathogens as well as for the development of T cell- and B cell-mediated adaptive responses. In addition, it has recently been reported that TLR activation can promote apoptosis of various cell lines (8, 9).
Sepsis is a major health problem (10, 11). During the onset of bacteremia in humans or rodents, massive apoptosis is observed inside lymphoid organs (11). Apoptosis can function as a highly efficient host mechanism to both limit the spread of intracellular pathogens and potentially down-regulate immune responses. In contrast, it is possible that pathogen-mediated induction of apoptosis is a strategy to evade immune responses. Despite the critical importance of DCs in the initiation and regulation of Th1 immune responses against bacteria (12, 13), very little is known on how bacteremia affects DCs in vivo.
Peritonitis continues to be a leading cause of morbidity and mortality (14, 15, 16). Although the overall mortality of sepsis is 35% (17), abdominal sepsis is associated with mortality rates of up to 80% (18). Although various bacteria have been identified as causative organisms in peritonitis, Escherichia coli belongs to the most common bacteria recovered from peritonitis (up to 60%; Ref. 19) and is the second most frequent bacterium to cause sepsis (up to 20%; Ref. 20).
In this work, we analyze whether E. coli bacteremia induce DC migration, maturation, and apoptosis in vivo. We further determine which TLRs and adaptor molecules are required for these effects on DC. Several components of E. coli have been reported to impact DCs. LPS induces DC activation (21) and apoptosis (22) in vivo. Bacterial DNA and its synthetic form CpG-oligodeoxynucleotides (CpG-ODN) also induce DC activation in vivo (23). Therefore, we analyzed the ability of E. coli bacteremia to induce DC maturation and apoptosis in vivo and compare them with those obtained after LPS and CpG-ODN administrations.
Materials and Methods
Mice and reagents
Female BALB/c and C57BL/6 mice (6–8 wk old) were purchased from Harlan Sprague Dawley. MyD88–/– (3), TLR2–/– (24), and TLR4–/– (25) mice on the BALB/c background and TRIF–/– mice on the C57BL/6 background (Beutler, Scripps Research Institute; Ref. 5) were bred in the specific pathogen-free animal facility of the Institute of Molecular and Cellular Pharmacology, Nice-Sophia Antipolis (Valbonne, France). The maintenance and care of mice complied with the guidelines of the Institut National de la Santé et de la Recherche Médicale/Centre National de Recherche Scientifique/University of Nice-Sophia Antipolis Ethic Committee for the use of laboratory animals. CpG-ODN 1826 was obtained from Coley Pharmaceutical Group. Highly purified LPS (E. coli serotype 0111:B4) was purchased from Invivogen. E. coli (strain K514) was grown to stationary phase (8–10 h) in Luria-Bertani medium and further washed (2500 x g, 10 min, 4°C) three times in PBS before inoculation into mice. Heat-killed (HK) E. coli were obtained by a 60-min incubation at 70°C. We checked that the HK bacteria did not grow after inoculation of an overnight culture in Luria-Bertani medium.
In vivo treatment
Mice were given injections i.v. into the lateral tail vein with indicated doses of LPS, CpG-ODN, or E. coli in PBS. Control animals were injected with the same volume of PBS.
MLR
Spleens were cut into very small pieces and incubated (37°C, 30 min) into HBSS (Life Technologies) containing 4000 U/ml collagenase IV (Roche) and 0.1 mg/ml DNase I fraction IX (Sigma-Aldrich Chimie SARL). CD4+ T cells from the spleen were purified by negative depletion of CD8+, B220+, CD11b+, and MHC II+ cells using sheep anti-rat Ig-coated Dynabeads (Dynal). The indicated numbers of gamma-irradiated (3000 rads) spleen cells were incubated with 3 x 105 allogeneic CD4+ T cells in DMEM supplemented with L-glutamine (2 mM), heat-inactivated FCS (10%), 2-ME (5 x 10–5 M), penicillin (100 μg/ml), and streptomycin (100 U/ml) in U-bottom 96-well plates. Cell proliferation after 48 h of culture was determined by [3H]thymidine incorporation.
Cytofluorometric analysis
Spleen cells were analyzed by flow cytometry with a FACSCalibur cytofluorometer (BD Biosciences). The cells were first preincubated with saturating doses of 2.4G2 (a rat anti-mouse Fc receptor mAb; American Type Culture Collection) for 10 min to prevent Ab binding to Fc receptor and then labeled with PE-coupled N418 (anti-CD11c), APC-coupled 53-6.7 (anti-CD8), and fluorescein (FITC)-coupled Abs including L3T4 (anti-CD4), 3.23 (anti-CD40), 16–10A1 (anti-CD80), GL1 (anti-CD86), or 2G9 Ab (anti-IA/IE). All mAbs were purchased from BD Biosciences. Cells were gated according to size and scatter to eliminate dead cells and debris from analysis.
Confocal microscopy
Tissues were fixed for 3 h at 4°C in 1% paraformaldehyde (pH 7.0), washed in PBS, incubated overnight at 4°C in a solution of 30% sucrose under agitation, and washed again in PBS. Tissues were embedded in the Tissue-Tek OCT compound (Sakura) and frozen at –80°C, and 10-μm cryostat sections were prepared. Tissues sections were incubated in 100 mM Tris-HCl, pH 8.0, to eliminate OCT and then in a saturated solution containing 10% normal serum in 100 mM Tris-HCl, pH 8.0. Avidin-biotin sites were saturated using an avidin-biotin blocking kit (Vector). Tissues sections were incubated with any of the following mAbs: Alexa Fluor 488 phalloidin (Molecular Probes); APC-coupled N418 (anti-CD11c); GL1 (anti-CD86); 11-26c.2a (anti-IgD); or 30-H12 (anti-CD90.2, Thy-1.2; BD Biosciences) or MOMA-1 (BMA Biomedicals) in 100 mM Tris-HCl, pH 8.0. Uncoupled mAbs were stained with Alexa Fluor 568-coupled goat anti-rat IgG (Molecular Probes). Slides were mounted in DAKO medium (DakoCytomation). Labeled cells were visualized under a Leica laser scanning confocal microscope (TCS-SP) equipped with a DM-IRBE inverted microscope and an argon-krypton laser.
Immunohistochemistry
Immunohistochemical techniques used in this study were previously described (26). Briefly, spleens were fixed in Immunohistofix (Aphase), embedded in Immunohistowax (Aphase), and sectioned at 3–6 μm. Tissues sections were then treated with 1% blocking reagent (Boehringer) to saturate nonspecific binding sites. The endogenous peroxidase activity was neutralized by adding 3% H2O2 in PBS for 30 min. The slides were further treated to visualize apoptotic cells using TUNEL (Boehringer, Mannheim) and costaining with alkaline phosphatase substrates (SK-5300, blue precipitate; Vector Laboratories). A blocking kit (Vector Laboratories) was used to saturate the excess of biotin. Slides were incubated with the N418-biotinylated Ab (10 μg/ml in PBS-0.5% blocking reagent), washed, and then incubated with avidin-biotin-peroxidase complexes (Vectastain ABC kit) revealed with a solution of AEC tablets (Sigma-Aldrich). Digitalized images were captured using a charge-coupled device color camera (Ikegami Tsushinki).
Results
Live and HK E. coli induce DC migration and maturation in vivo
To model the potential effect of bacteremia on DCs in lymphoid organs, live or HK E. coli (5 x 107 bacteria) were injected i.v. into BALB/c mice. This treatment induced the migration of splenic DCs (CD11chigh cells) to T cell zones as shown by confocal microscopy on spleen sections from wild-type (wt) BALB/c mice (Fig. 1A and data not shown for HK E. coli). This result parallels previous findings using LPS (21). DCs migrate from red pulp and marginal zone to T cell-rich areas surrounding the central arteriole within a 60-h time frame. Marginal metallophilic macrophages (MOMA-1+ cells) are localized in the inner part of the marginal zone and form a discontinued ring delineating the border between red pulp and white pulp. DC migration to T cell zones occurs through areas of the marginal zone containing fewer MOMA-1+ cells (Fig. 1, A and B). Similar migration profiles are observed after i.v. administration of 105–108 bacteria, as well as with 50 μg of the TLR9 ligand CpG-ODN (data not shown). We also observed that CD11c+ cells, localized in the T cell area, up-regulate the expression of CD86 on inoculation of HK and live E. coli and LPS (Fig. 1C) as well as with HK and CpG-ODN (data not shown). In addition, high levels of constitutive CD86 were expressed on the CD11c– MOMA-1+ macrophage population in the marginal zone, as shown previously (27). Similar results with regard to DC migration and maturation were also seen in C57BL/6 mice (data not shown).
To confirm results observed using immunohistochemistry, flow cytometry was used to analyze cell surface expression of MHC II, CD40, CD80, and CD86 on splenic CD11chigh cells 9 h after administration of HK and live E. coli, LPS (50 μg), and CpG-ODN (50 μg) (Fig. 1D). The effects observed on DC maturation were similar whether E. coli were administered through the i.p. or i.v. route, and the kinetics was similar to that seen after i.v injection of LPS or CpG with peak expression of maturation markers observed at 6–12 h postinjection. Moreover, all CD11chigh DC subsets were affected (Fig. 1E for CD86; data not shown for CD40, CD80, and MHC II). Interestingly, because the administration of live and HK E. coli induces comparable up-regulation of costimulatory molecules on DCs, this suggests that structural components of E. coli are sufficient to induce complete DC maturation.
It has been recently found that commercial LPS preparations are frequently contaminated by lipoprotein and other bacterial components that interact with TLR2 (28). For these reasons, we injected highly purified E. coli LPS in all experiments. In addition, to prevent mice mortality after E. coli injections, we immunized them with an E. coli strain (K514) exhibiting a low virulence. However, short term experiments monitoring DC migration and maturation after injection of a virulent strain (018:K1) gave similar results (data not shown).
E. coli-induced DC maturation is dependent on TLR2, TLR4, and TRIF in vivo
To characterize the receptors and the intracellular signaling pathways involved in E. coli-induced DC migration and maturation, we inoculated wt, TLR2–/–, TLR4–/–, MyD88–/–, and TRIF–/– mice with 5 x 107 live E. coli, LPS (50 μg), and CpG-ODN (50 μg) (Fig. 2). Because DCs from wt BALB/c or C57BL/6 immunized recipients undergo comparable migration and maturation (data not shown), only data obtained with wt BALB/c mice are presented.
Discussion
Systemic LPS administration has been used in previous studies to visualize DC maturation and apoptosis in vivo (21, 22). However, to our knowledge, these data have never been validated in a more physiological model, such as a bacterial infection. In this study, we demonstrate that E. coli bacteremia induces DC migration, maturation, and apoptosis with magnitudes and kinetics similar to that of LPS in vivo. Although all DC subsets mature in response to E. coli or LPS, apoptosis preferentially affects CD8+ DCs that are described to preferentially induce Th1 responses (12, 13). It is conceivable that CD8+ DCs are more sensitive to apoptosis because they express higher levels of TLR4 (30). Along similar lines, preferential apoptosis of CD8+ DCs has recently been described in lymph nodes during polymicrobial sepsis (31). Thus, DC apoptosis can be a general consequence of bacterial infection. In the model of polymicrobial sepsis (31), DC maturation is not observed before DC apoptosis.
As a possible consequence of DC apoptosis, we observed that E. coli bacteremia decreased the ability of total spleen cells to stimulate naive T cells during MLR. It is possible that the DC apoptosis that we observed could be a contributing factor to the immunosuppression associated with sepsis (32, 33). In agreement with this idea, reduction in DC numbers has been correlated in several systems with a reduced ability to mount T cell responses in vivo (13, 21). Moreover, DC loss has been associated with a decreased ability of recipient mice to produce systemic IL-12 in response to LPS injection (34). This suggests, perhaps not surprisingly, that massive DC apoptosis can specifically impair antibacterial Th1 responses.
The impact of TLRs, TRIF, and MyD88 in E. coli, LPS, and CpG-ODN-induced DC maturation and apoptosis is summarized in Table I. E. coli-induced DC maturation is reduced in TLR2–/– and TLR4–/– mice, demonstrating that both TLRs are important PAMP receptors for this pathogen. It also suggests that TLR2 and TLR4 participate in detection and control of E. coli infection. The requirement for TRIF in LPS-induced DC maturation has been described previously (29), but the role of TRIFs in E. coli bacteremia-induced DC maturation is a novel observation. This suggests a possible role for TRIF in the development of adaptive immune response against bacteria.
Although LPS-mediated DC activation requires signals emanating from TLR4 and the downstream binding of TIRAP, TRAM, TRIF, and MyD88 adaptor proteins (2), little has been described regarding the roles of these receptors and signaling molecules involved in LPS-induced DC apoptosis in vivo. However, it is known that the death receptor Fas is not involved in this process (21). We show here that LPS-induced splenic DC apoptosis in vivo depends on TLR4, TRIF, and MyD88 adaptor proteins. In addition, we demonstrate that the TLR4-mediated TRIF-dependent signaling pathway contributes to the control of DC apoptosis during E. coli bacteremia. Several recent in vitro studies using macrophages, bone marrow-derived DCs, and other cell lines show a direct role of TLR4 (8, 9), MyD88 (35), and TRIF (9) on apoptosis. Another in vitro study using TLR3 ligands suggests that TRIF regulates three distinct signaling pathways leading to the induction of the IFN- promoter, NF-B activation, and apoptosis via a FADD/caspase-8-dependent pathway (36).
CpG-ODN injection promoted DC migration and maturation but induced less DC apoptosis compared with LPS and E. coli; this was seen by several different assays (Figs. 3A, 5C, and 5D). CpG-ODN induced DC apoptosis is completely abrogated by MyD88 deficiency. Our in vivo results are in agreement with a recent work (37) showing that in vitro stimulation with LPS- but not CpG-induced apoptosis of bone marrow-derived DC. This study concluded that the ability of CpG to up-regulate Bcl-2, a potent anti-apoptotic molecule, counteracted the induction of programmed cell death of bone marrow-derived DC. Thus, the weak ability of CpG-ODN to induced DC apoptosis suggests a limited implication of bacterial DNA in E. coli-induced DC apoptosis and may explain its important role as a potent adjuvant in vivo.
Taken together, our results provide new insight into how Gram-negative bacteremia modulates DCs function and survival. Coevolution between bacteria and host immune systems has selected the emergence of PAMPs receptors to initiate innate immunity to these pathogens and control infection. However, during septicemia, bacteria subvert this detection system and induce massive DC apoptosis. This phenomenon most likely impairs or reduces the initiation and/or development of adaptive immune responses.
Acknowledgments
We thank Drs. P. M. Dubois and C. A. Benedict for criticism of this manuscript.
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 grants from the Belgian program on Interuniversity Poles of Attraction (to P.A.I.) initiated by the Belgian State, Prime Minister’s Office, Science Policy Programming, the Fonds National de la Recherche Scientifique (Crédit aux chercheurs, Belgium), the Ministère de l’Education Nationale, de la Recherche et de l’Enseignement Supérieur (France), and the Institut National de la Santé et de la Recherche Médicale. M.M. is Senior Research Associated from Fonds National de la Recherche Scientifique. E.M. is Research Associated from Fonds National de la Recherche Scientifique.
2 Address correspondence and reprint requests to Dr. Eric Muraille, Laboratoire de Parasitologie, Faculté de Médecine, Route de Lennik 808, 1070 Bruxelles, Université Libre de Bruxelles, Brussels, Belgium. E-mail address: emuraille@hotmail.com
3 Abbreviations used in this paper: DC, dendritic cell; PAMP, pathogen-associated molecular pattern; TIR, Toll-IL-1R; TRIF, TIR-domain-containing adapter-inducing interferon-; TRAM, TRIF-related adapter molecule; CpG-ODN, CpG-oligodeoxynucleotide; HK, heat killed; wt, wild type; TICAM, TIR domain-containing adaptor molecule.
Received for publication November 10, 2004. Accepted for publication April 27, 2005.
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