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Published online before print December 19, 2005
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Surgical Research, Department of Trauma Surgery, University Hospital Essen, Germany
1 Correspondence: Surgical Research, Department of Trauma Surgery, University Hospital Essen, Virchowstr. 171, D-45147 Essen, Germany. E-mail: stefanie.flohe{at}medizin.uni-essen.de
ABSTRACT
Polymicrobial sepsis is associated with immunosuppression caused by the predominance of anti-inflammatory mediators and profound loss of lymphocytes through apoptosis. Dendritic cells (DC) are potent antigen-presenting cells and play a key role in T cell activation. We tested the hypothesis that DC are involved in sepsis-mediated immunosuppression in a mouse cecal ligation and puncture (CLP) model, which resembles human polymicrobial sepsis. At different time-points after CLP, DC from the spleen and peripheral lymph nodes were characterized in terms of expression of costimulatory molecules, cytokine synthesis, and subset composition. Splenic DC strongly up-regulated CD86 and CD40 but not CD80 as soon as 8 h after CLP. In contrast, lymph node DC equally increased the expression of CD86, CD40, and CD80. However, this process of maturation occurred later in the lymph nodes than in the spleen. Splenic DC from septic mice were unable to secrete interleukin (IL)-12, even upon stimulation with CpG or lipopolysaccharide + CD40 ligand, but released high levels of IL-10 in comparison to DC from control mice. Neutralization of endogenous IL-10 could not restore IL-12 secretion by DC of septic mice. In addition, the splenic CD4+CD8– and CD4–CD8+ subpopulations were lost during sepsis, and the remaining DC showed a reduced capacity for allogeneic T cell activation associated with decreased IL-2 synthesis. Thus, during sepsis, splenic DC acquire a state of aberrant responsiveness to bacterial stimuli, and two DC subtypes are selectively lost. These changes in DC behavior might contribute to impaired host response against bacteria during sepsis.
Key Words: bacterial infections • inflammation • cytokines • costimulation
INTRODUCTION
During sepsis, massive deterioration of the immune response occurs, which may result in multi-organ failure and death. Cecal ligation and puncture (CLP) in mice causes symptoms similar to those found in septic patients and serves as a model for polymicrobial sepsis. Pro- and anti-inflammatory cytokines are expressed in large amounts after CLP, and the balance of their expression seems to be critical for the outcome of sepsis. Interleukin (IL)-12 and its counterpart IL-10 play a major role during sepsis, as modulation of these cytokines influences sepsis-induced mortality after CLP in mice. Application of recombinant IL-10 prior to CLP increases survival [1
], and administration of IL-12 [2
] increases sepsis-induced mortality. However, the time of intervention appears to be decisive for disease development. Absence of IL-10 during the initial phase of sepsis is detrimental [3
], whereas neutralization of IL-10 at later time-points is beneficial [4
]. In addition to the predominance of anti-inflammatory mediators, T as well B lymphocytes undergo apoptosis in the spleen and other lymphoid organs during sepsis [5
]. The remaining T cells display reduced responsiveness to mitogens in terms of proliferation and secretion of IL-2 and interferon-
(IFN-) [6
]. This status of dysregulation of a broad range of the innate and adaptive immunity during sepsis has been entitled with terms such as "immunoparalysis."
It has to be emphasized that the changes of the immune system observed during polymicrobial sepsis cannot be simply reduced to effects mediated by bacteria-derived lipopolysaccharides (LPS). Mice, which are defective in the LPS signaling molecule Toll-like receptor 4, are susceptible to polymicrobial sepsis but not to LPS-induced shock [2 ]. T cell suppression is observed upon polymicrobial sepsis but not upon infusion with LPS [6 ]. These findings demonstrate that multiple factors contribute to polymicrobial sepsis-mediated immunomodulation. Although being a well-described phenomenon, the basic mechanisms leading to sepsis-caused immunoparalysis have not yet been characterized completely.
Dendritic cells (DC) are professional antigen-presenting cells (APC) and are sentinels of the immune system. Upon encounter of antigen, e.g., from invading pathogens, they migrate to the draining lymphoid organ, where they initiate the primary immune response [7 ]. Conventional DC express high levels of CD11c and major histocompatibility complex (MHC) class II molecules. During migration, these DC undergo major phenotypic and functional changes, termed maturation. Upon maturation, DC strongly up-regulate MHC and costimulatory molecules such as CD86, CD80, and CD40, which are all required for effective T cell activation. In addition, during maturation, DC secrete cytokines, which polarize T helper (Th) cells toward Th1 or Th2, depending on the type of the secreted cytokines. The DC-derived cytokine IL-12 favors the differentiation of Th1 cells and is important for the development of immunity against bacterial infections. In contrast, DC-derived IL-10 promotes the polarization of Th cells toward Th2, which mediates immunity against extracellular parasites [7 ].
As a result of high levels of MHC class II and costimulatory molecules, mature DC are potent T cell activators and are superior in comparison to other APC, such as macrophages or B cells [8 ]. In contrast, DC, which lack high levels of costimulatory molecules and/or do not secrete pro-inflammatory cytokines, are involved in tolerance induction. In this case, tolerance is mediated through activation of regulatory T (Treg) cells or through induction of T cell apoptosis [9 ]. Thus, depending on the pattern of costimulatory molecules and secreted cytokines, DC determine the fate of the immune response [10 ].
The role of DC during polymicrobial sepsis is largely unknown. As a result of their decisive role in T cell activation, a potentially deviated DC behavior could account for immunosuppression observed during sepsis. Indeed, apoptosis of DC in the spleen [11 , 12 ] and lymph nodes [13 ] during sepsis was described, suggesting the involvement of DC during disease development. In the present study, the maturation state, cytokine secretion pattern, and T cell-stimulatory capacity of DC during CLP-induced sepsis were analyzed. The spleen, representing a central lymphoid organ located next to the site of infection and peripheral lymph nodes, was chosen for DC analyses.
MATERIALS AND METHODS
Mice
Female BALB/c and C57BL/6J mice (all from Harlan Winkelmann, Borchen, Germany) were 8–10 weeks old and had access to standard rodent food and water ad libitum.
Culture medium (CM) and reagents
Very low endotoxin medium RPMI 1640 (Biochrom, Berlin, Germany), containing 10% heat-inactivated fetal calf serum, 10 mM HEPES, 2 mM L-glutamine, 0.06 mg/ml penicillin, 0.02 mg/ml gentamicin G, and 0.05 mM 2-mercaptoethanol, was used as CM. Murine recombinant granulocyte macrophage-colony stimulating factor (GM-CSF) and CD40 ligand (CD40L) were purchased from R&D Systems (Wiesbaden, Germany). Synthetic phosphorothioated CpG 1668 oligonucleotides [14
] were purchased from Qiagen (Köln, Germany). LPS (Escherichia coli 026:B6), ionomycin, and phorbol 12-myristate 13-acetate (PMA) were from Sigma Chemical Co. (Deisenhofen, Germany). All reagents were free of detectable LPS contaminations as tested using Limulus amoebocyte assay.
Induction of polymicrobial sepsis
The CLP model for polymicrobial sepsis developed by Chaudry et al. [15
] was used with some modifications. Briefly, after performing a mid-line laparotomy, the cecum was exposed and ligated, and the distal part was punctured once with a 17-gauge needle. A small amount of cecum contents was extruded through the perforation. After returning the cecum into the abdomen and application of 1 ml saline intraperitoneally (i.p.) for resuscitation, the abdomen was closed in a single layer technique. Control animals underwent a sham operation with laparotomy alone. Under these conditions, CLP caused severe sepsis with a mortality rate of 70% within 36 h.
Preparation of total spleen and lymph node cells
At different time-points after CLP or sham operation, spleens were removed, and single cell suspensions were prepared by collagenase digestion using 0.02 U/ml Blendzyme 2 (Roche, Grenzach-Wyhlen, Germany) at 37°C for 18 min. Spleens were meshed through a cell strainer, and cells were incubated in CM containing 5 mM EDTA for 5 min for dissociation of cell clusters. After washing, total spleen cells (106 cells/well) were cultured in 48-well plates, with or without 5 µg/ml CpG for 18 h. Nonadherent cells were harvested and used for intracellular stainings.
Inguinal and popliteal lymph nodes were isolated and flushed with CM using a 27-g needle. Remaining tissue was minced through a cell strainer. After pooling, single cells were incubated in CM containing 2 mM EDTA to dissociate cell clusters and used for fluorescence-associated cell sorter (FACS) stainings.
Isolation and culture of splenic DC
Splenic DC were purified using CD11c microbeads (Miltenyi, Mönchengladbach, Germany) according to the manufacturer’s recommendations. Purity was generally 85–90%, as confirmed by CD11c staining and FACS analyses. Purified, splenic DC were cultured in CM containing 0.3 ng/ml GM-CSF in the absence or presence of 100 ng/ml LPS + 2.5 µg/ml CD40L or 5 µg/ml CpG in 96-well flat-bottom plates (105 cells/well). In some experiments, stimulation of DC was performed in the presence of neutralizing anti-IL-10 antibodies (10 µg/ml, clone JES052A5, R&D Systems) or the respective rat immunoglobulin G1 (IgG1) isotype control (10 µg/ml, R&D Systems). Previous experiments have shown that 10 µg/ml anti-IL-10 antibodies were sufficient to neutralize the activity of 1000 pg/ml IL-10 (data not shown). After 18 h, supernatants were collected and stored at –20°C until analysis for IL-12 and IL-10 using the cytometric bead array mouse inflammation kit (BD Biosciences, Heidelberg, Germany).
FACS analyses
Cells (0.5–1x106) were washed with Cell Wash (BD Biosciences) and resuspended in 50 µl Cell Wash containing 1 µg FcBlock (BD Biosciences) for blocking unspecific binding. After 6 min, 50 µl antibody mixture (all antibodies from BD Biosciences) containing anti-CD11c-allophycocyanin (clone HL3) in combination with anti-CD40-fluorescein isothiocyanate (FITC; clone 3/23), anti-CD86-phycoerithrin (PE; clone GL1), anti-CD80-FITC (clone 16-10A1), anti-H2-Kd-PE (clone SF1-1.1), anti-IAd-FITC (clone 2G9), anti-CD8-FITC (clone 53-6.7), or anti-CD4-PE (clone RM4-5) was added. After 15 min on ice, cells were washed and resuspended in Cell Wash. For intracellular staining, cells were stimulated and incubated for the last 4–6 h in the presence of monensin (GolgiStop 0.66 µl/ml, BD Biosciences). After surface staining using anti-CD11c-allophycocyanin or anti-CD4-FITC antibodies, cells were fixed and permeabilized using Cytofix/Cytoperm (BD Biosciences) for 20 min at room temperature. Thereafter, intracellular cytokines were stained using anti-IL-12p40-PE (clone C15.6) or anti-IL-2-PE (clone JES6-5H4) antibodies. After washing with permeabilization buffer (BD Biosciences), cells were resuspended in Cell Wash. Appropriate isotype controls were used for all stainings. All data were acquired using a FACScalibur (BD Biosciences) and analyzed using CellQuest Pro software (BD Biosciences). For all stainings, the FL-3 channel was used to exclude autofluorescent cells. Living cells were selected according to forward- and side-scatter properties. The mean fluorescence intensity (MFI) of the respective molecules stained in combination with the anti-CD11c antibody was determined on gated, splenic CD11chi and lymph node CD11c+ DC. Quadrant lines in dot plots were set as indicated in the figure legends. Total DC number per spleen was calculated as % CD11chi cells x total number of cells per spleen/100.
Allogeneic T cell assay
Splenic CD3+ T cells from C57BL/6 mice were purified using the Pan T cell isolation kit (Miltenyi), according to the manufacturer’s instructions. Purity of T cells was usually >90% as determined by FACS analyses. T cells were labeled with carboxyfluorescein diacetate succinimidyl ester (CFSE; 500 nM for 12 min at 37°C, Molecular Probes, Göttingen, Germany), and 2 x 105 T cells were added to titrated amounts of purified, splenic DC per well (96-well, half-area, flat-bottom plate, Corning, Schiphol-Rijk, The Netherlands). All cultures were set up in triplicates. After 3 days, cells were harvested, and cell clusters were dissociated through addition of 7.5 mM EDTA for 15 min. After washing with Cell Wash, T cells were stained using anti-CD3-PE antibody (clone 145-2C11, BD Biosciences). A constant number of CaliBRITE allophycocyanin beads (BD Biosciences) were added to allow the acquisition of equal parts per culture. 7-Amino-actinomycin (4 µg/ml, Molecular Probes) was used to exclude dead cells. For data acquisition, a constant number of CaliBRITE beads were counted. Living T cells (CD3+7–AAD–) were gated, and the number of divided cells, showing less than the maximal CFSE fluorescence intensity, was determined. For analysis of IL-2 synthesis, cocultures were restimulated with 5 ng/ml PMA, 500 ng/ml ionomycin, and monensin for 4 h before staining for intracellular IL-2 and FACS analyses.
Statistical analyses
The paired Student’s t-test was used to compare differences between sham and CLP groups. The unpaired Student’s t-test was used to compare differences between replicates of cell cultures within one experiment. A P value <0.05 was considered to be significant.
RESULTS
Maturation of DC in the spleen during polymicrobial sepsis
To get insight into the effects of sepsis on splenic DC, these cells were analyzed at different time-points following CLP or sham operation. Spleens were removed by 8, 15, and 36 h after operation and stained for CD11c. DC were identified according to their high expression of CD11c (Fig. 1A
). In control animals, 0.94 ± 0.15% of total spleen cells were DC (Fig. 1B)
. The total number of DC per spleen was calculated. The number of DC per spleen from septic and sham mice did not differ until 15 h after CLP (Fig. 1C)
. However, 36 h after CLP, the number of DC in spleens from septic mice drastically declined down to 20% of DC usually found in spleens from control mice (Fig. 1C) .
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It has been reported that endogenous IL-10 in DC acts in an autocrine manner and regulates IL-12 synthesis of DC [16 ]. Thus, the putative role of endogenous IL-10 in impaired IL-12 synthesis from DC during sepsis was investigated. Splenic DC were purified 24 h after CLP or sham operation, and stimulation with CpG was performed in the presence or absence of neutralizing anti-IL-10 antibodies or the respective isotype control. Thereafter, supernatants were tested for IL-12. In response to CpG, DC from sham mice released high levels of IL-12, which were increased further in the presence of anti-IL-10 antibodies but not in the presence of the isotype control antibodies (Fig. 5 ). However, CpG-stimulated DC from septic mice did not release IL-12 in the absence or in the presence of anti-IL-10 or isotype control antibodies (Fig. 5) . Taken together, during sepsis, DC develop an inverse cytokine secretion pattern in response to bacterial stimuli associated with a predominance of IL-10.
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In the present study, we showed that polymicrobial sepsis induced strong maturation of splenic DC and a deviated cytokine secretion pattern of these DC toward IL-10. This maturation process was not only restricted to the spleen as a central lymphoid compartment but was also observed in lymph nodes distant to the site of infection. However, maturation of DC from lymph nodes of septic mice followed a slightly delayed kinetics in comparison to splenic DC. Additionally, we provide evidence that selectively, the splenic CD4+CD8– and CD4–CD8+ DC subpopulations were lost during disease development and that remaining DC from septic mice were inhibited in their capacity for T cell activation.
A variety of components released from Gram-negative as well as Gram-positive bacteria has been shown to induce maturation of DC [18 19 20 21 ]. Indeed, it has been reported that intraperitoneal bacteria reach liver and spleen rapidly via the blood circulation [22 ]. In addition, we detected bacteria in the spleen as early as 4 h after CLP (data not shown). From the peritoneal site of infection, direct bacterial spreading via the peritoneal cavity or via the blood circulation may be responsible for the rapid maturation of splenic DC. In contrast, DC from the popliteal or inguinal lymph nodes, which do not drain the initial site of infection, show a delayed kinetics of maturation.
Recently, Ding et al. [23 ] reported that peritoneal and splenic DC do not enhance the expression of CD86 or CD40 upon CLP-induced sepsis within 24 h. This observation is in marked contrast to our findings. However, they also report a lower IL-12 production in splenic DC from septic mice, which is in alignment with our findings. Different results in terms of DC surface marker expression might be explained by different gating strategies during FACS analysis. Another possible explanation for these partially contradictory results is the fact that the inbred strain they used (C3H/HeN) might be less susceptible for sepsis [24 ]. Efron et al. [13 ] reported that the total number of DC in popliteal and inguinal lymph nodes decreases within 24 h after CLP-induced sepsis without any signs for maturation in this timeframe. Our data confirm the absence of DC maturation at least during the early phase of sepsis up to 8 h. We only observed marginal effects in terms of DC maturation by 15 h after CLP, and obvious maturation of lymph node DC was detected by 36 h after CLP (Fig. 3) .
We could demonstrate that sepsis did not only induce profound maturation of DC but also modulated splenic DC in terms of cytokine secretion. Within 4–8 h after CLP, DC became unable to secrete bioactive IL-12 in response to LPS + CD40L or to CpG, both well-known inducers of IL-12 (Fig. 4)
. DC secrete IL-12 only transiently for 
16 h after stimulation [25
] before they become unresponsive to further restimulation. This characteristic has been termed "exhaustion" of DC and might represent a mechanism for protection from infection-induced immunopathology caused by unregulated IL-12 production. A state of impaired IL-12 secretion by splenic DC upon restimulation with microbial components has been reported for DC in other models of infectious diseases [26
, 27
]. However, such an immunosuppressive state reported for splenic DC in the above-mentioned models of infection is preceded by a transient phase of IL-12 release and might be explained by DC exhaustion. Our observations are in contrast to the findings in the above-mentioned models of infection, as we found unresponsiveness to restimulation of splenic DC during sepsis but not a preceding phase of IL-12 secretion. Instead, DC from septic mice secreted prominent levels of IL-10 ex vivo, which were further enhanced upon stimulation with otherwise IL-12-inducing agents (Fig. 4B)
. The reason for this aberrant cytokine response of splenic DC during sepsis remains unknown. DC-derived IL-10 seemed not to be responsible for suppressed IL-12 production, as neutralization of IL-10 did not restore IL-12 production by splenic DC from septic mice (Fig. 5)
. Thus, during sepsis, splenic DC fail to secrete IL-12, which is required for the induction of immunity against bacterial infections.
It is interesting that sepsis seems not to be associated with a general inhibition of IL-12 production by DC, as it has been reported that DC in the peritoneal cavity secrete substantial amounts of IL-12 during sepsis [23 ]. The underlying mechanisms for this compartment-dependent IL-12 release by DC remain unclear. We suggest that there exist differences in local microenvironmental factors between spleen and peritoneal cavity during sepsis, which might cause different responses of DC to bacteria. Potential candidates for such factors could be prostaglandin E2 (PGE2), IL-10, and transforming growth factor-ß, which are known to suppress IL-12 production by DC [28 ]. IL-10 and PGE2 have been found in serum and/or spleen early during sepsis [29 , 30 ]. PGE2 inhibits the release of IL-12 by DC stimulated with bacterial products but does not affect the up-regulation of costimulatory molecules [31 ]. In contrast, IL-10 inhibits IL-12 synthesis and maturation of DC [32 , 33 ]. As splenic DC from septic mice showed maturation and suppression of IL-12 synthesis (Figs. 2 and 4) , it is unlikely that circulating IL-10 alone mediates the development of the deviated DC function during sepsis, which we describe here. Rather, we hypothesize that DC in the peritoneal cavity representing the primary site of infection are stimulated directly by gut-derived microbes and are able to respond with adequate IL-12 production. In contrast, splenic DC previously exposed to sepsis-induced mediators such as circulating PGE2 are therefore instructed for an impaired IL-12 secretion when stimulated by spreading bacteria or circulating bacterial products.
Evidence for compartmentalization of the cytokine release during CLP-induced sepsis has also been described for the release of IFN-. Mononuclear cells from liver and from the peritoneal cavity but not from the spleen produce IFN- [34
]. This compartmentalization of cytokine production might explain why intravenous (i.v.) application of an adenoviral vector coding for IL-10 fails to improve survival after CLP, whereas subcutaneous application protects from CLP-induced mortality [35
]. Splenic DC seem to be predisposed for IL-10 secretion during sepsis (Fig. 4B)
. Thus, additional IL-10 applied via the i.v. route would further support the predominance of anti-inflammatory mediators in the spleen. In this context, it would be of interest whether DC from peripheral lymph nodes of septic mice are impaired in IL-12 secretion as we observed for splenic DC. However, the low number of lymph node DC, especially in septic mice, was not sufficient for setting up the required cell cultures. Therefore, the behavior of lymph node DC remains unknown.
The predominance of IL-10 released from splenic DC during sepsis as shown here might polarize Th cells toward Th2 and in parallel, might inhibit the development of a Th1 response, which is required for the effective clearance of bacterial infections [16 ]. The finding that splenocytes from septic mice secrete elevated levels of Th2 cytokines upon mitogenic stimulation [6 ] supports this assumption. The suppressive activity of IL-10 on Th1 cell polarization and proliferation might explain the unexpected finding that DC from septic mice were inferior to DC from controls in T cell activation, despite the expression of high levels of costimulatory molecules (Fig. 6) .
Recently, DC with the "semi-mature" phenotype of MHC class II+/CD86+ but negative for secretion of proinflammatory cytokines were found to induce tolerance through Treg cells [36 , 37 ], which inhibit the proliferation and cytokine release of antigen-specific Th1 cells. It is tempting to speculate that semi-mature DC develop in the spleen during sepsis and activate Treg cells, thereby preventing the induction of a protective Th1 immune response. This hypothesis is supported by the observation that elevated numbers of Treg cells were found in septic patients [38 ].
During late sepsis, the number of splenic DC expressing CD11c was strongly reduced. CD11c is a stable population marker for murine DC, and its expression remains unchanged upon maturation. So far, there is no evidence that DC down-regulate CD11c expression in vitro or in vivo. Therefore, we assume that the reduced number of splenic CD11c+ cells shown here represents a loss of DC similar to the loss of CD11c+ DC, which has been observed, e.g., during sepsis, endotoxemia, and viral infections [12 , 18 , 39 ]. We could further show that the reduced number of splenic DC during late sepsis is mediated by a selective loss of the CD4+CD8– and CD4–CD8+ DC subpopulations. In contrast, the total number of CD4–CD8– DC remained unaffected, and CD4–CD8– DC became a main subpopulation by 36 h after CLP (Fig. 7) . The three DC subpopulations arise from distinct DC lineages, and in vivo, there is no conversion of one subtype into another, e.g., through down-regulation of CD4 or CD8 expression [40 ]. Likewise, CD4+ or CD8+ DC remain clearly positive for these markers, even upon maturation in vitro [17 ]. Thus, we assume that the CD4+CD8– and CD4–CD8+ DC are lost during sepsis rather than that these subtypes transform into CD4–CD8– DC.
One might speculate that the reduced T cell-stimulatory capacity and the impaired IL-12 production by DC observed 36 h after CLP are a phenomenon characteristic for CD4–CD8– DC. However, several reports argue against divergent properties of the splenic DC subpopulations with regard to maturation and T cell activation [41 , 42 ]. Moreover, the finding that splenic DC failed to secrete IL-12, even during the early phase of sepsis (8 h after CLP, Fig. 4B ) when all DC subtypes were present (Fig. 7B) , further contradicts the assumption of a CD4–CD8– DC-restricted phenomenon.
The disappearance of two DC subsets during sepsis may occur through migration and/or death. As apoptosis of DC has already been reported to be associated with sepsis (in humans and in mice) [11 12 13 ], we suppose that the selective loss of CD4+CD8– and CD4–CD8+ DC, which we show here, is the consequence of apoptosis. It is well known that DC undergo apoptosis after reaching the final maturation stage, e.g., upon administration of LPS [43 ]. Such a programmed DC death might represent a tool for regulation of the immune response. However, we never observed an enhanced maturation stage of CD4+CD8– and CD4–CD8+ DC in comparison with CD4–CD8– DC, which could result in the predominant loss of these DC subtypes (data not shown). Thus, the mechanisms of selective DC subset depletion remain speculative.
In summary, we describe here profound maturation of DC during polymicrobial sepsis. However, DC acquire a phenotype that might favor the development of Th2 and/or Treg cells and might inhibit effective immunity against the bacterial infection through Th1 cell polarization. Changes in DC phenotype or functions could therefore contribute to sepsis-mediated immunosuppression. Treatment regimens, which restore the capacity of DC for Th1 cell polarization or the regular DC number [44 ], seem to represent promising approaches to counteract immunoparalysis during sepsis.
ACKNOWLEDGEMENTS
This work is supported by DFG Grants FL-391 (to S. B. F.) and FL-353/2.1 (to S. F.). We are grateful to Dr. E. Kreuzfelder and B. Nyadu for technical support in terms of FACS analyses.
Received July 26, 2005; revised October 18, 2005; accepted November 10, 2005.
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