During Polymicrobial Sepsis Dendritic Cells in Murine Lymph Nodes ...

Characterization of the Systemic Loss of Dendritic Cells in Murine Lymph Nodes During Polymicrobial Sepsis This information is current as of June 13, 2013.

Philip A. Efron, Antonio Martins, Douglas Minnich, Kevin Tinsley, Ricardo Ungaro, Frances R. Bahjat, Richard Hotchkiss, Michael Clare-Salzler and Lyle L. Moldawer J Immunol 2004; 173:3035-3043; ; http://www.jimmunol.org/content/173/5/3035

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The Journal of Immunology is published twice each month by The American Association of Immunologists, Inc., 9650 Rockville Pike, Bethesda, MD 20814-3994. Copyright © 2004 by The American Association of Immunologists All rights reserved. Print ISSN: 0022-1767 Online ISSN: 1550-6606.

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References

The Journal of Immunology

Characterization of the Systemic Loss of Dendritic Cells in Murine Lymph Nodes During Polymicrobial Sepsis1,2 Philip A. Efron,* Antonio Martins,* Douglas Minnich,* Kevin Tinsley,‡ Ricardo Ungaro,* Frances R. Bahjat,* Richard Hotchkiss,‡§ Michael Clare-Salzler,† and Lyle L. Moldawer3*

S

epsis represents a significant health problem worldwide. In the U.S. alone, sepsis is the twelfth most common cause of death, afflicting ⬎750,000 people and killing ⬎210,000 people annually (1–5). The annual incidence of sepsis in 1989 was 175 cases per 100,000 people, and this incidence has been steadily increasing, with current levels more than triple those seen in the early 1970s (3, 5). There are at present few therapeutic approaches for the treatment of septic patients. Results from clinical trials examining high dose steroids, platelet-activating factor antagonists, and drugs that inhibit the actions of TNF-␣, IL-1, PGs, and bradykinin have been disappointing, and only activated protein C (Xigris; Eli Lilly, Indianapolis, IN) has been approved for the treatment of severe sepsis. There is growing recognition that many of the immunological defects that accompany sepsis syndrome are in the acquired immune response, and contribute to the adverse clinical outcome. Defects in innate and acquired immunity have been reported in the septic patient (2, 6). In recent years, interest has focused on the increased lymphocyte apoptosis observed in experimental animals and patients with sepsis (7, 8). Increased apoptosis has been observed in the lymphoid tissue of mice subsequent to burns or

Departments of *Surgery and †Medicine, University of Florida College of Medicine, Gainesville, FL 32608; and Departments of ‡Anesthesiology, and §Surgery, Medicine, and Pathology, Washington University School of Medicine, St. Louis, MO 63110 Received for publication January 21, 2004. Accepted for publication June 29, 2004. 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 in part by Grants R37 GM-40561-15, and R01 GM-6304103, awarded by the National Institute of General Medical Sciences, U.S. Public Health Service. P.A.E. was supported by a National Institute of General Medical Sciencesfunded T32 Training Grant. 2 This study was presented at the Surgical Forum, American College of Surgeons 88th Annual Clinical Congress, San Francisco, CA, October 6 –10, 2002; and the 26th Annual Shock Society Meeting, Phoenix, AZ, June 7–10, 2003. 3 Address correspondence and reprint requests to Dr. Lyle L. Moldawer, Department of Surgery, University of Florida College of Medicine, Room 6116, Shands Hospital, Box 100286, Gainesville, FL 32610-0286. E-mail address: [email protected]

Copyright © 2004 by The American Association of Immunologists, Inc.

polymicrobial sepsis (cecal ligation and puncture (CLP))4 (9 –17). Human studies have verified this increased lymphocyte apoptosis. Septic patients are frequently lymphopenic (18); the lymphoid epithelia of the gastrointestinal tract and spleens of human patients who succumbed to sepsis and multiple organ dysfunction also have increased apoptosis. Spleens from septic patients also demonstrate increased levels of caspase 3 and decreased numbers of CD3⫹CD4⫹ Th and B cells (19, 20). We have further demonstrated the importance of lymphocyte apoptosis in the sepsis response by demonstrating improved outcome with specific caspase 3 inhibition (21, 22). Whether through overexpression of an antiapoptotic protein (Bcl-2), injection of a polycaspase inhibitor, or knockout of a major apoptotic enzyme (caspase 3), mice with decreased lymphocyte apoptosis during polymicrobial sepsis have improved survival (21, 22). Similar data have been observed in a porcine model of sepsis (23). Because murine lymphocyte apoptosis can occur in the absence of mature B and T cells as well as their secretory products, as demonstrated in Rag-1 mice (17), the cellular and humoral factors responsible for regulating lymphocyte apoptosis during sepsis have been the source of much investigation. The dendritic cell (DC) plays a pivotal role in lymphocyte apoptosis as well as in the immune suppression often seen in sepsis syndromes. First described by Steinman et al. (24 –26) as a novel cell population in the murine spleen, DCs exist in all lymphoid organs and most other tissues of the reticuloendothelial system (27). DCs are a potent APC, in which they serve as a critical link between the innate and acquired immune systems (6, 28 –32). Stereotypically, DCs differentiate from progenitor cells into immature, resting DCs that undergo maturation into an effector cell in response to a variety of exogenous stimuli, including microbial products (6, 33). After encountering a stimulus/Ag, DCs undergo this maturation or activation, resulting in a number of functional and phenotypic changes,

4 Abbreviations used in this paper: CLP, cecal ligation and puncture; 7-AAD, 7-aminoactinomycin D; DC, dendritic cell.

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Dendritic cells (DCs) play a key role in critical illness and are depleted in spleens from septic patients and mice. To date, few studies have characterized the systemic effect of sepsis on DC populations in lymphoid tissues. We analyzed the phenotype of DCs and Th cells present in the local (mesenteric) and distant (inguinal and popliteal) lymph nodes of mice with induced polymicrobial sepsis (cecal ligation and puncture). Flow cytometry and immunohistochemical staining demonstrated that there was a significant local (mesenteric nodes) and partial systemic (inguinal, but not popliteal nodes) loss of DCs from lymph nodes in septic mice, and that this process was associated with increased apoptosis. This sepsis-induced loss of DCs occurred after CD3ⴙCD4ⴙ T cell activation and loss in the lymph nodes, and the loss of DCs was not preceded by any sustained increase in their maturation status. In addition, there was no preferential loss of either mature/activated (MHCIIhigh/CD86high) or immature (MHCIIlow/CD86low) DCs during sepsis. However, there was a preferential loss of CD8ⴙ DCs in the local and distant lymph nodes. The loss of DCs in lymphoid tissue, particularly CD8ⴙ lymphoid-derived DCs, may contribute to the alterations in acquired immune status that frequently accompany sepsis. The Journal of Immunology, 2004, 173: 3035–3043.

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Materials and Methods Mice Specific pathogen-free, female C57BL/6 mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and were used between 6 and 10 wk of age. Mice were maintained on standard rodent food and water ad libitum. These mice were allowed a minimum of 5 days to equilibrate to their environment before any experimental use. The studies were approved by the Institutional Animal Care and Use Committee at the University of Florida College of Medicine before their initiation.

Induction of polymicrobial sepsis For induction of polymicrobial sepsis, mice were subjected to a CLP, as previously described (46). In brief, a laparotomy was made and the cecum was isolated, ligated with 3-0 silk suture, and punctured through and through with an 18-gauge needle. The cecum was replaced into the abdomen, and the peritoneum and skin were closed using two surgical clips. Mice that underwent sham operations also had a laparotomy, but simply had their cecum briefly exposed, with subsequent peritoneal and skin closure. Only 20% of the mice undergoing CLP survive 72 h, and mortality begins to occur after 24 h (47). The current study examined the early DC response to this lethal polymicrobial sepsis. Six to 10 mice were sacrificed at 6, 12, and 24 h after surgery. In addition, an equal number of mice that did not undergo an operation were sacrificed. Subsequent to sacrifice, the popliteal and inguinal lymph nodes as well as the two mesenteric lymph nodes most proximal to the cecum were isolated and placed into iced PBS. Isolation of all six nodes took less than 10 min following euthanasia. The nodes of each operational and anatomical group were pooled together. The total number of mice used and lymph nodes collected for each group for each experiment were equivalent.

Lymph node single cell suspensions The lymph nodes were resuspended in 4 ml of HBSS without phenol red and with calcium and magnesium (Mediatech, Herndon, VA) containing 100 U/ml collagenase D solution (Boehringer Mannheim, Norwich, CT) and placed into a 60 ⫻ 15-mm tissue culture dish (Falcon; BD Biosciences, San Jose, CA) on ice. Subsequently, the lymph nodes were dissected with two 30-gauge needles. The suspension from the dish was removed and placed into 30 ml of HBSS without phenol red, calcium, or magnesium (Mediatech) on ice. The tissue fragments remaining in the dish were resuspended in 4 ml of HBSS with 400 ␮l/ml collagenase D and transferred to a 15-ml conical tube. These tubes were placed in a 37°C water bath for 30 min, and the solutions were pipetted vigorously, creating a single cell suspension. The cell suspension was filtered through a 70-␮m mesh filter (Falcon; BD Biosciences) into a 50-ml conical tube containing the previously acquired cells. The cells were suspended with 50 ml of HBSS and centrifuged at 400 ⫻ g for 10 min at 4°C. The supernatant was aspirated and the cells were resuspended in 50 ml of HBSS and centrifuged. The cells were then resuspended in 5 ml of HBSS and counted on a hemocytometer. Subsequently, 1 ⫻ 106 cells were dispensed into 5-ml polystyrene round-bottom tubes and stained for flow cytometry.

Flow cytometry After washing twice with HBSS containing 1% BSA, 1 mM EDTA (Fisher Scientific, Pittsburgh, PA), and 0.1% sodium azide (NaN3; Sigma-Aldrich, St. Louis, MO), the cells were resuspended in 4% BSA flow buffer, blocked with CD16/CD32 Fc Ab (BD Pharmingen, San Diego, CA), and then stained. Cells from each operation and node subtype were stained in one of five groups: 1) FITC-conjugated rat anti-mouse MHC class II mAb, PEconjugate hamster anti-mouse CD11c mAb, 7-aminoactinomycin D (7AAD), and biotin-conjugated rat anti-mouse CD86 mAb; 2) MHCII FITC, CD11c PE, 7-AAD, and biotin-conjugated rat anti-mouse CD8a mAb; 3) FITC-conjugated hamster anti-mouse CD69 mAb, PE-conjugated hamster anti-mouse CD3, 7-AAD, and biotin-conjugated rat anti-mouse CD4 mAb; and 4) FITC-conjugated rat anti-mouse CD69, CD3 PE, 7-AAD, and CD4 biotin (all Abs; BD Pharmingen). Also, additional cells were stained with MHCII FITC and CD11c PE, followed by two washes in cold 1⫻ binding buffer (10 mM HEPES/NaOH, 140 mM NaCl, 2.5 mM CaCl2 (all components; Sigma-Aldrich), pH 7.4). Cells were then resuspended in 100 ␮l of 1⫻ binding buffer and biotin-conjugated annexin V and 7-AAD (BD Pharmingen). Anti-biotinylated Ab labeled with allophycocyanin (BD Pharmingen) was used for indirect staining for biotin-conjugated samples (i.e., CD86, CD8, CD4, and annexin V). Samples were acquired and analyzed on one of two six-parameter FACSCalibur machines with CellQuest software (BD Biosciences) at the University of Florida Flow Cytometry Core Laboratory. FACSCalibur machines were calibrated before each use. A total of 0.2–1.0 ⫻ 106 events was collected per sample for DC analysis, with the same number of events being collected for all tubes of a given experiment. For non-annexin V staining, ⬃1 ⫻ 104 CD11c⫹/7-AAD⫺ events were collected per tube. For annexin V-stained samples, ⬃1 ⫻ 104 CD11c⫹ events were collected. A total of 1 ⫻ 105 events was collected for Th cell analysis, with ⬎1 ⫻ 104 CD3⫹/CD4⫹/7-AAD⫺ events being collected.


including increased ability to migrate to lymphoid tissue, decreased endocytosis, up-regulation of membrane MHCII and costimulatory molecules, and increased specific cytokine expression. Many of these phenotypic changes result in the DC becoming a potent stimulator of T lymphocytes, and critical for T cell priming (6, 32). Nevertheless, the concept that DCs exist only in either an immature or mature phenotype is under challenge. Rather, depending upon the stimulus, its timing, and the environment in which the DC resides at the time of its stimulus, the ultimate DC phenotype and function will vary (6, 32) (our unpublished data). In addition, there are a number of DC subsets that have additional functions beyond Ag processing and presentation. One such cell is the murine CD8⫹ DC, previously thought to play a primary role in immune tolerance and anergy. Initially labeled lymphoid DCs because they were presumed to be derived from lymphoid precursors, CD8⫹ DCs are currently the subject of much debate in regard to their lineage and function. Although it has been demonstrated that these cells have the capacity to induce immune tolerance and are a likely candidate in mice as tolerogenic DCs, there are contrasting data demonstrating that CD8⫹ DCs also prime cells for a Th1 response, and that certain subsets of CD8⫺ DCs can induce tolerance (31, 32, 34 –38). Additionally, there is a subset of DCs known as follicular DCs, found in the B cell regions of lymphoid tissue, which maintain B cell function and growth. Although these cells are most likely derived from a different cell lineage, they are considered to be DCs based on their morphology and their ability to present Ag, albeit in a different manner, to B lymphocytes (36, 39 – 41). In vivo murine and human studies increasingly point to the importance of the loss of DCs and more specifically the apoptosis of DCs during sepsis. There is a significant loss of splenic DCs within 24 h of LPS administration, and LPS has been demonstrated to induce apoptosis of DCs in vivo (42– 44). Previous studies have illustrated a specific decrease in the splenic DC population in septic human patients when compared with patients who have suffered trauma (45). In addition, an apoptotic loss of splenic DCs in septic mice 24 – 48 h after the onset of polymicrobial sepsis (CLP) has also been demonstrated (39). The objectives of our study were to characterize the DC response in lymph nodes to polymicrobial sepsis, allowing us to simultaneously analyze the response in tissues that were and were not in the same body cavity with as well as having direct lymphatic drainage from the source of microbial infection. This included determining whether a relative and absolute change in the DC population of lymph node tissue occurred with time during polymicrobial sepsis, and whether apoptosis played a role in this change. In addition, we examined whether there was a specific increase or loss of mature DCs as well as CD8⫹ DCs in the lymph nodes. The loss of these specific DC subpopulations might further impair the ability of the host to contain and/or respond to the inflammation displayed in response to generalized peritonitis. Finally, we wished to analyze how these changes related temporally to the well-known loss of T lymphocytes during polymicrobial sepsis.

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Flow cytometry analysis

Data analysis

During analysis, debris and 7-AAD⫹ cells were excluded through gating, except in the annexin V-stained samples, which only had debris excluded. Gating based on observed populations (Fig. 1) was initially determined on the cells derived from lymph nodes obtained from healthy animals (no surgery) in contour plots and then applied to cells derived from sham- and CLP-treated animals. These populations were initially verified by staining with isotype control Abs. Each subtype of sample as defined by its operation, lymph node location, and time point was performed at least in triplicate.

Data were analyzed as a ratio of the percentage of positively stained cells from CLP- or sham-treated mice. This was calculated by dividing the percentage of positive staining cells in CLP or sham mice by the percentage of positive staining cells in the healthy, control mice from the same experiment (i.e., cells isolated from healthy mice the same day as cells isolated from CLP- or sham-treated mice).

Immunohistochemical staining

Data are reported as the mean ⫾ SEM. Data were analyzed using the statistical software program SigmaStat v.2.03 (SPSS, Chicago, IL). For multivariant comparison among groups, a two-way ANOVA was used with an all pairwise multiple comparison procedure being performed using the Fisher least significant difference method. Differences were considered significant at p ⬍ 0.05.

Results Sepsis induced a relative loss in the percentage of DCs present in the murine lymph nodes A significant loss in the number of CD11c⫹ DCs was observed in the local mesenteric nodes beginning 12 h post-CLP, and reached a 50% decline by 24 h (Fig. 2). In the systemic inguinal nodes, the number of CD11c⫹ DCs declined by a similar amount at 24 h post-CLP ( p ⬍ 0.05). A similar trend was observed in the systemic popliteal nodes, albeit statistical significance was not obtained. These data were verified by immunohistochemical staining in which there was a ubiquitous loss of DC positive staining throughout the T cell regions of the murine lymph node (Fig. 3). DC apoptosis and death were increased in the murine lymph nodes during polymicrobial sepsis There was also a significant increase ( p ⬍ 0.05) in the percentage of apoptotic and dead DCs in the mesenteric and inguinal nodes

FIGURE 1. DC and T cell flow cytometric analyses. After gating out debris and dead cells (7-AAD⫹ cells), DCs (CD11c⫹ events) were identified on a MHCII (x-axis)- vs CD11c (y-axis)-gated contour plot (A). Subsequently, gated DCs were analyzed for maturity on a MHCII (x-axis) vs CD86 (y-axis) contour plot (B); MHCIIhigh and CD86high were considered mature (upper right quadrant). In addition, gated DCs were analyzed for their expression of CD8 on a forward scatter vs CD8⫹ contour plot (C). For the analysis of DC apoptosis, dead cells (7-AAD⫹) were not gated out; rather, after gating out debris, DCs were determined by a MHCII vs CD11c contour plot and then gated to an annexin V⫹ (x-axis) vs 7-AAD⫹ (y-axis) contour plot (D). The lower right quadrant are annexin V⫹ DCs, while the upper right quadrant are annexin V⫹ and 7-AAD⫹ DCs. T cell analysis removed debris and dead cells (7-AAD⫹), as described above. Cells were considered CD3⫹CD4⫹ T cells if they were both CD3high and CD4high (upper right gate) on a CD3 (x-axis) vs CD4 (y-axis) contour plot (E). Subsequently, CD3⫹CD4⫹ T cells were analyzed for their activation status on CD69 (x-axis) vs CD4 (y-axis) and CD25 (x-axis) vs CD4 (y-axis) contour plots (F and G, respectively).


Immunohistochemical staining of samples obtained 24 h after CLP or the sham procedure was performed, as previously described (39). Lymph nodes were immediately isolated and placed into liquid nitrogen. Specimens were quickly frozen in OCT medium and cryosectioned in 10-␮m sections, and then immediately fixed in ice-cold acetone for 4 min and stored at ⫺80°C. All staining was performed at room temperature in a humidified chamber, and dilutions and washings were performed with PBS and 1% BSA. A purified hamster anti-mouse CD11c, integrin ␣␹-chain (BD Pharmingen), was used at a dilution of 1/20. After a 1-h incubation with the primary Ab, the slides were incubated with a 1/75 diluted Texas Red-conjugated AffiniPure goat anti-Armenian hamster IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) for 30 min. Subsequently, the slides were rinsed and stained with a 1/100 diluted polyclonal Ab for active caspase 3 (Cell Signaling Technology, Beverly, MA) for 1 h. A 1/1000 diluted secondary Alexa 488 Fluorophore goat anti-rabbit Ab (Molecular Probes, Eugene, OR) was incubated with the slides for 30 min. For follicular DC staining, a 1/30 diluted rat anti-mouse FDC Ab (clone FDC-M1; BD Pharmingen) was incubated with the slide for 1 h. Subsequently, the slide was incubated with 1/400 diluted Alexa Fluor 594 goat anti-rat Ab (Molecular Probes). Active caspase 3 colocalization was the same as that of colocalization of CD11c⫹ cells on lymph node tissue section slides (see above). Fluorescently labeled lymph node tissue was visualized using a Nikon Eclipse E600 microscope equipped with a dual filter cube selective for FITC and Texas Red. A minimum of 5–10 high-powered fields (⫻100) was evaluated in each sample.

Statistical analysis

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24 h after polymicrobial sepsis (Fig. 4). These data were also verified by immunohistochemical staining, specifically for expression of active caspase 3. Increased active caspase 3 was observed in DCs

FIGURE 3. Decrease in the DC population and indirect evidence of increased apoptosis of CD11c⫹ DCs in the mesenteric lymph nodes 24 h after CLP. A and B, Cells have been stained with an Ab directed against CD11c⫹ (red). C and D, Cells were stained for both CD11c⫹ and with an Ab directed against active caspase 3 (green). Cells expressing both CD11c⫹ and active caspase 3 are stained as yellow. CD11c⫹ DCs (stained red) are seen ubiquitously throughout T cell zones of the lymph node from sham-treated animals (A), while in B (CLP), there were areas in the T cell zones lacking DCs (arrows). Although some caspase 3 activity was seen in the lymph nodes of sham-treated mice (C), little activity was associated with the CD11c⫹ cells (double stained yellow). However, increased active caspase 3 was visible in the CD11c⫹ DCs in sepsis (arrows) (D). Similar results for interdigitating and follicular DCs were found in the inguinal and popliteal nodes (data not shown).

FIGURE 4. Apoptotic and dead DCs present in the murine lymph nodes. Results are presented as the ratio of the percentages of annexin V⫹ or annexin V⫹- and 7-AAD⫹-positive cells in CLP- and sham-treated animals compared with healthy controls. There was a significant increase in the relative number of apoptotic or dead cells 24 h in the mesenteric lymph nodes after CLP in the septic mice (p ⬍ 0.05). In addition, there was also a significant increase in the percentage of apoptotic/dead cells from 6 h post-CLP vs 24 h (p ⬍ 0.05). The septic inguinal nodes also had a significant increase in the percentage of apoptotic and dead DCs 24 h postoperation (p ⬍ 0.05). There was no significant difference in the level of apoptosis and death in the popliteal lymph nodes after either operation. There was a time-dependent change (6 vs 24 h, 12 vs 24 h, p ⬍ 0.05).

in the T cell regions (interdigitating DCs) 24 h after CLP (Fig. 4). In addition, active caspase 3 activity was also detected in the DCs in the B cell regions (follicular lymph DCs) of the murine lymph nodes 24 h following the induction of polymicrobial sepsis (Fig. 5).


FIGURE 2. Relative levels of DCs present in murine lymph nodes postCLP. Results are presented as a percentage of cells from CLP septic- and sham-treated animals compared with healthy controls evaluated on the same day. The hours postinduction of sepsis are presented on the x-axis, while the ratio of CD11c⫹ events is presented on the y-axis (percentage of CD11c⫹ events from CLP- or sham-treated animals/percentage of CD11c⫹ events from healthy mice). DC populations in mesenteric lymph nodes from CLP mice were significantly different from those of sham mice (p ⬍ 0.05 CLP vs sham). A significant decrease was seen at 12 h post-CLP, and continued at 24 h (p ⬍ 0.05). In addition, a significant difference was noted in the CLP group between the 6- and 12-h samples, as well as between the 12- and 24-h samples (p ⬍ 0.05). As a group, septic mice also differed from sham mice in the percentage of DCs in the inguinal lymph nodes (p ⬍ 0.05). The relative decrease in DC population in CLP mice was significant 24 h postoperation, compared with sham mice (p ⬍ 0.05). Popliteal lymph nodes of septic mice displayed a similar decrease in their DC population as compared with sham mice, although statistical significance was not reached.

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FIGURE 5. Indirect evidence of apoptosis of follicular DCs in the mesenteric lymph nodes 24 h after CLP. A, Cells have been stained with a mAb directed against FDC-M1 (red). B, Cells were stained for both FDC-M1 and with a mAb directed against active caspase 3 (green). The follicular DCs (stained red) are seen in the B cell regions of the lymph node (arrows). Active caspase 3 (green) can be seen in the follicular DC regions, as well in double-stained (yellow) follicular DCs (arrows).

Polymicrobial sepsis neither increased the percentage of mature DCs present in the murine lymph nodes early in septic response nor caused a specific loss of mature DCs later during the septic response

CLP induces a relative loss in the CD8⫹ DCs in the murine lymph nodes

The systemic loss of DCs followed the loss in CD3⫹CD4⫹ T cells during polymicrobial sepsis Significant decreases were seen in the CD3⫹CD4⫹ T cell population 6 –12 h post-CLP, while losses in the DC population were

A 50% decrease in the percentage of CD8⫹ DCs was observed in mesenteric and popliteal lymph nodes 12 and 24 h after CLP ( p ⬍ 0.05) (Fig. 6). Although not significant, a similar trend was displayed in the inguinal nodes from septic animals. Polymicrobial sepsis caused a loss of CD3⫹CD4⫹ T cells associated with T cell activation that precedes the loss of DCs in murine lymph nodes In general, CD3⫹CD4⫹ T cells from the lymph nodes of septic mice had an increased expression of activation markers, as measured by surface expression of CD25 (IL-2R) and CD69 (very early activation Ag). Also, there was a decreased percentage of CD3⫹CD4⫹ T cells present in the lymph nodes of septic mice (Fig. 7). Specifically, mesenteric lymph nodes had a small, but significant decrease in the relative percentage of CD3⫹CD4⫹ T cells within 6 h of the onset of sepsis ( p ⬍ 0.05). Although there was no significant change in their expression of CD25, there was an increase in CD69 expression. The most dramatic changes were Table I. Comparison of mature DCs (MHCIIhigh/CD86high) present in the lymph node of septic and sham-treated micea Hours Postoperation Operation and Lymph Node

CLP mesenteric lymph nodes Sham mesenteric lymph nodes CLP inguinal lymph nodes Sham inguinal lymph nodes CLP popliteal lymph nodes Sham popliteal lymph nodes

6

12

24

1.10 ⫾ 0.08

0.87 ⫾ 0.03

1.33 ⫾ 0.19

1.07 ⫾ 0.28

1.02 ⫾ 0.05

1.38 ⫾ 0.09

0.92 ⫾ 0.07

0.82 ⫾ 0.14

1.25 ⫾ 0.27

0.81 ⫾ 0.06

0.99 ⫾ 0.07

1.09 ⫾ 0.11

1.10 ⫾ 0.11

1.25 ⫾ 0.16

1.16 ⫾ 0.08

0.99 ⫾ 0.13

1.12 ⫾ 0.18

1.05 ⫾ 0.05

a Numbers represent the ratio of MCHIIhigh/CD86high DCs present, determined by the ratio of MCHIIhigh/CD86high DCs present in the CLP- or sham-treated mice vs those present in the healthy control mice, and the SEM. There was no significant difference in the level of maturity of the DCs present in the lymph nodes during polymicrobial sepsis as compared with sham-operated mice.

FIGURE 6. Relative levels of CD8⫹ DCs present in murine lymph nodes post-CLP. Results are presented as the ratio of the percentages of CD8⫹ DCs in CLP- and sham-treated animals compared with healthy controls. The mesenteric CD8⫹ DC populations were different in CLP mice as compared with sham mice (p ⬍ 0.05). Although there was also a time-dependent difference in both groups (6 vs 24 h, p ⬍ 0.001), beginning at 12 h and continuing to 24 h, there was also a significant decrease in the CD8⫹ DC population in the mesenteric nodes from CLP mice (p ⬍ 0.05). The decrease in CD8⫹ DCs from inguinal nodes followed a similar trend to the mesenteric and inguinal lymph nodes from septic mice, although statistical significance was not obtained. Once again, there was a time-dependent difference in both the septic and nonseptic populations (12 vs 24 h, p ⬍ 0.05). Popliteal nodes from CLP- and sham-treated mice were different (p ⬍ 0.05), and there was a time-dependent decrease (6 vs 12 h, and 12 vs 24 h, p ⬍ 0.001 and p ⬍ 0.05, respectively) in CD8⫹ DC populations. Beginning at 12 h and continuing to 24 h, there was significant drop in the relative CD8⫹ DC population in popliteal lymph nodes from septic mice.


No significant differences were observed in the percentage of mature DCs (MHCIIhigh and CD86high) in the lymph nodes of CLPvs sham-operated mice (Table I). This was true in the local and distant lymph nodes at 6, 12, and 24 h postoperation.

seen in the inguinal nodes, simultaneously demonstrating an increase in the activation status of CD3⫹CD4⫹ T cells with the relative loss of the CD3⫹CD4⫹ T cells with time. Popliteal lymph nodes also exhibited a loss in their CD3⫹CD4⫹ T cell population with an increased activation status of the remaining living CD3⫹CD4⫹ T cells.

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FIGURE 8. Loss of DCs in relation to the loss of CD3⫹CD4⫹ cells. Ratios (right y-axis) of CD3⫹CD4⫹ cells in CLP- and sham-treated mice as compared with healthy controls (—) are displayed as the line graph, while ratios (left y-axis) of CD11c⫹ DCs in CLP- and sham-treated mice are displayed in the bar graph. Systemically, the loss of CD3⫹CD4⫹ cells from murine lymph nodes occurs within 12 h following CLP, while the loss of DCs usually occurred between 12 and 24 h following CLP.

and no-operation mice, although these reductions did not reach statistical significance. Thus, the relative loss of DCs and CD3⫹CD4⫹ T cells observed in septic murine lymph nodes represents an absolute

not observed until 12–24 h after induction of polymicrobial sepsis (Fig. 8). There was a decrease in the absolute population of cells present in the lymph nodes of septic mice Total cell counts from pooled lymph nodes from each experiment were compared, demonstrating that the total number of cells present in the lymph nodes from septic mice was never greater than that of the cell numbers from sham or healthy mice (Fig. 9). In general, at 12 and 24 h post-CLP, there were a fewer number of total lymph node cells in the septic mice as compared with sham

FIGURE 9. Total cell counts of pooled lymph nodes. There was no change in the cell counts in lymph nodes from CLP as compared with sham operation over 24 h, although CLP total cell number tended to decline. More importantly, cell counts were never significantly greater in lymph nodes from CLP than from sham or no-operation cell counts. This confirms that the relative loss of DCs and CD3⫹CD4⫹ cells, as determined by flow cytometry, most likely represents an absolute decline in the number of DCs, CD8⫹ DCs, and CD3⫹CD4⫹ cells.


FIGURE 7. Relative population of CD3⫹CD4⫹ T cells in the murine lymph nodes and their activation status as measured by expression of CD25 and CD69. The presence of CD3⫹CD4⫹ T cells in the lymph nodes is presented as a line graph displaying the hours postoperation (x-axis) and the ratio of cells (right y-axis) from CLP- and sham-treated animals vs cells from healthy controls. The activation status of these CD3⫹CD4⫹ T cells is displayed in the bar graph as the hours postoperation (x-axis) and the ratio of CD25⫹- as well as CD69⫹-positive cells (left y-axis) in CLP- and shamtreated animals compared with healthy controls. Although the difference in the percentage of CD3⫹CD4⫹ cells from mesenteric lymph node populations in CLP and sham-treated mice was small, it did obtain statistical significance (CLP vs sham, p ⬍ 0.001). There was no difference in the CD25⫹ expression, but CLP-treated mice had increased CD69⫹ expression, compared with sham-treated mice (CLP vs sham, p ⬍ 0.05). In the inguinal nodes, there was a time-dependent decrease in the CD3⫹CD4⫹ cell populations and an increase in the activation status postprocedure (6 vs 12 h, 6 vs 24 h, p ⬍ 0.001). Septic mice not only had significantly lower CD3⫹CD4⫹ cell populations, but those CD3⫹CD4⫹ cells had significantly higher levels of CD25 and CD69 expression (CLP vs sham, p ⬍ 0.001). This decrease in the CD3⫹CD4⫹ cell population as well as the increased activation status in the setting of polymicrobial sepsis become significant at 12 h postprocedure and continued at 24 h postprocedure (CLP, 12 h vs sham, 12 h; CLP, 24 h vs sham, 24 h, p ⬍ 0.001; except for CLP, 12 h CD25 vs sham, 12 h CD25, p ⬍ 0.05). Popliteal lymph nodes from CLP mice were different from popliteal lymph nodes from sham mice in their relative decreased population of CD3⫹CD4⫹ cells as well as their increased activation status of those Th cells (CLP vs sham CD3⫹CD4⫹, p ⬍ 0.05; CLP vs sham CD25⫹ or CD69⫹, p ⬍ 0.001).

SEPSIS AND THE DC

The Journal of Immunology decrease in the cell population rather than an increase in another cell population.

Discussion

an exaggerated proinflammatory state. However, CD8␣⫹ can also induce a Th1 response (32), so their specific loss could also play a role in the immune paralysis witness later in the septic response. The overall loss of DCs during the progression of sepsis may contribute to suppression of the acquired immune response. The possible relationship between the loss of DCs and the sustained immune suppression may occur at several levels. Being an integral part of both the innate and acquired immune systems, DCs migrate throughout the body and act as sentinels by constantly sampling their environment (31). Because DCs are the most potent APCs, their loss could significantly affect the capacity of the host to mount an acquired immune system. This not only includes lymphocytes, but other cells known to interact with DCs, including NK cells, which are increasingly being shown to play a role in the sepsis response (50 –54). DCs may drive apoptosis in selected lymphocyte populations (36, 39 – 41). Inadequate stimulation or a lack of costimulation from DCs engenders anergic or apoptotic T cells (31, 55, 56). Thus, the increased activation of Th cells without proper costimulation during sepsis could lead to their anergy or apoptosis (57). One mechanism of this inadequate costimulation may simply be the loss of either CD3⫹CD4⫹ T cells or DCs during systemic infection, and thus, decreased overall costimulation. Another possible mechanism is a sepsis-induced change in DC function. Sepsis most likely alters the paracrine cytokine profile expressed by DCs at the point of naive T cell interaction, a process that plays a role in determining Th cell polarization (31). For example, DCs are able to secrete IL-12, a cytokine that induces the expansion of Th1 clones, for 8 –12 h after maturation (58). DCs exposed to endotoxin in vivo multiple times have a reduced capacity to express IL-12 (59). As mentioned above, naive T cells that do not receive appropriate costimulation may become anergic. These anergic T cells can, in turn, suppress DC function (60). Thus, it is possible that a feedback loop may occur later in the sepsis response in which immunosuppressive T cells pathologically suppress appropriate DC function, creating a vicious cycle. Finally, the increased presence of apoptotic cells themselves during polymicrobial sepsis can worsen outcome. We have previously demonstrated that the injection of apoptotic splenocytes into mice worsens survival during CLP as compared with the injection of necrotic cells, which improves survival (61). The endocytosis of apoptotic material by the remaining DCs could also affect their ability to function and their viability (62). Other mechanisms besides apoptosis may also be responsible for the decline in DC populations in lymphoid tissue during sepsis. The acquired immune system uses a significant number of attractant and migration molecules, with CD3⫹CD4⫹ T cells and DCs interacting through chemokines and their cell surface receptors. Initially, DCs migrate to inflammatory tissue due to chemoattractants produced by the cells of that environment (63– 65). As the DC matures, the DC’s chemokine membrane receptor profile changes, and thus, its sensitivity to chemokines of lymphoid origin increases (31). Thus, changes in the ability of the DC to appropriately migrate may contribute to the decline of DCs in the lymphoid tissue during sepsis. In addition, it is possible that a lack of functional precursors could contribute to this decline of the DC population in the lymph nodes. It has been reported in a murine burn and sepsis model that there is significant decline in DC precursors after injury (66). Human studies have demonstrated a similar phenomenon, with peripheral blood monocytes having a decreased capacity to differentiate into immature DCs in trauma patients (67). The changes in DC numbers and phenotypes may be exacerbated by the severity of the CLP model as well as the strain of


Our present findings demonstrate for the first time that there is a general loss in the DC population from both local and distant murine lymph nodes during the progression of polymicrobial sepsis. This loss in both relative and absolute numbers of DCs occurs temporally after the loss of mouse CD3⫹CD4⫹ T lymphocytes. At least some of the loss of DCs from these lymph organs could be attributed to increased apoptosis. Surprisingly, there was no specific increase in the maturation status of the DCs present in the lymph nodes during the evolution of the septic process. Our data are therefore consistent with the demonstrated loss of DCs from the spleen reported by our laboratories (39). In addition, we have demonstrated that there was some systemic loss of DCs in nondirectly draining nonabdominal compartment lymphoid tissue distant from the site of infection. As DCs are considered vital for communication between the innate and acquired immune systems, their loss in all lymphoid tissues would significantly impact the immune status of the host. In addition, this systemic loss of DCs distant to the site of infection reinforces the importance and complexity of the multifactorial response of the host to both systemic exogenous and endogenous factors during sepsis. The loss of DCs from the lymph nodes occurs after the loss of CD3⫹CD4⫹ T cells in the same lymph node tissue. The rapidity of the loss of CD3⫹CD4⫹ T cells during induction of murine polymicrobial sepsis (within 6 h) is similar to that seen in other systemic inflammatory syndromes. We have previously reported that there was a rapid increase in the rate of lymphocyte caspase 3-dependent apoptosis in the thymus and spleen within 3 h following a scald burn in mice (48). In both the burn injury model reported previously and the present studies with polymicrobial sepsis, the increased apoptotic loss of CD3⫹CD4⫹ T cells would have preceded the loss of DCs. Although it is not possible from these experiments to conclude that one process is dependent upon the other, the findings are not consistent with the proposal that mature or apoptotic DCs drive the activation-induced cell death of CD3⫹CD4⫹ T cells. There was neither an increased presence of mature DCs nor a specific loss of immature or mature DCs at any point during the development of polymicrobial sepsis. Our data are not consistent with the increased proportion of activated or mature DCs seen following LPS treatment. Although LPS administration does not fully recapitulate the complexity of the sepsis response, much of what we know about the DC role in the sepsis response comes from studies in which DCs have been exposed to bacterial endotoxin, and to date, it has provided a good model for examining the DC response to microbial products and, presumably, microbial invasion. For example, DCs express MHC class II molecules very early after LPS stimulation (49), and there is up-regulation of T cell costimulatory molecules, such as CD86. A possible explanation for this disparity between the response to in vitro LPS exposure and CLP may be the complexity of the septic disease response. Numerous other bacterial products, glucocorticoids, catecholamines, and host cytokines are produced during the early septic response, many of which have differing effects on DC and T cell maturation and function. Despite no overall effect of sepsis on the DC maturation status, CLP did induce a preferential loss of CD8⫹ DCs from the murine lymph nodes. Although much debated, murine CD8⫹ cells are known to play a role in immune tolerance (35). Splenic CD8␣⫹ DEC205⫹ DCs induce very little allogeneic response from CD8⫹ T cells (35). The depletion of CD8⫹ DCs could be responsible for exacerbation of the immune response, which would contribute to

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3042 mouse used. Different inbred strains of mice have different responses and outcomes to sepsis (68 –70). Because we did not examine the DC response in CLP models with varying degrees of severity, it is possible that the dramatic decrease in the DC number and increase in DC apoptosis could be exaggerated due to the particular strain of mouse and/or the severity of the model. However, increased DC loss and apoptosis were seen in the spleens of C57BL/6 mice following a less severe model of sepsis (39). In summary, polymicrobial sepsis induces both a significant systemic and local loss of DC cells in the lymph nodes, and this process follows the apoptotic loss of CD3⫹CD4⫹ T cells known to occur during sepsis syndromes. This decline in the DC population does not appear to affect DCs of a particular activation state, although CD8⫹ DCs are significantly depleted, possibly contributing to the unregulated proinflammatory state seen in polymicrobial sepsis. The DC depletion in lymph nodes that is due, at least in part, to apoptosis, could contribute to global immune suppression seen in sepsis syndromes.

1. Hotchkiss, R. S., and I. E. Karl. 2003. The pathophysiology and treatment of sepsis. N. Engl. J. Med. 348:138. 2. Oberholzer, A., C. Oberholzer, and L. L. Moldawer. 2001. Sepsis syndromes: understanding the role of innate and acquired immunity. Shock 16:83. 3. Angus, D. C., and R. S. Wax. 2001. Epidemiology of sepsis: an update. Crit. Care Med. 29:S109. 4. Hoyert, D. L., K. D. Kochanek, and S. L. Murphy. 1999. Deaths: final data for 1997. Natl. Vital Stat. Rep. 47:1. 5. Stone, R. 1994. Search for sepsis drugs goes on despite past failures. Science 264:365. 6. Efron, P., and L. L. Moldawer. 2003. Sepsis and the dendritic cell. Shock 20:386. 7. Oberholzer, C., A. Oberholzer, M. Clare-Salzler, and L. L. Moldawer. 2001. Apoptosis in sepsis: a new target for therapeutic exploration. FASEB J. 15:879. 8. Oberholzer, A., C. Oberholzer, R. M. Minter, and L. L. Moldawer. 2001. Considering immunomodulatory therapies in the septic patient: should apoptosis be a potential therapeutic target? Immunol. Lett. 75:221. 9. Fukuzuka, K., C. K. Edwards III, M. Clare-Salzer, E. M. Copeland III, L. L. Moldawer, and D. W. Mozingo. 2000. Glucocorticoid and Fas ligand induced mucosal lymphocyte apoptosis after burn injury. J. Trauma 49:710. 10. Fukuzuka, K., C. K. Edwards III, M. Clare-Salzler, E. M. Copeland III, L. L. Moldawer, and D. W. Mozingo. 2000. Glucocorticoid-induced, caspasedependent organ apoptosis early after burn injury. Am. J. Physiol. Regul. Integr. Comp. Physiol. 278:R1005. 11. Woodside, K. J., M. Spies, X. W. Wu, J. Song, S. S. Quadeer, J. A. Daller, and S. E. Wolf. 2003. Decreased lymphocyte apoptosis by anti-tumor necrosis factor antibody in Peyer’s patches after severe burn. Shock 20:70. 12. Ayala, A., Y. X. Xu, C. S. Chung, and I. H. Chaudry. 1999. Does Fas ligand or endotoxin contribute to thymic apoptosis during polymicrobial sepsis? Shock 11:211. 13. Ayala, A., C. D. Herdon, D. L. Lehman, C. A. Ayala, and I. H. Chaudry. 1996. Differential induction of apoptosis in lymphoid tissues during sepsis: variation in onset, frequency, and the nature of the mediators. Blood 87:4261. 14. Ayala, A., Y. Xin Xu, C. A. Ayala, D. E. Sonefeld, S. M. Karr, T. A. Evans, and I. H. Chaudry. 1998. Increased mucosal B-lymphocyte apoptosis during polymicrobial sepsis is a Fas ligand but not an endotoxin-mediated process. Blood 91:1362. 15. Chung, C. S., Y. X. Xu, W. Wang, I. H. Chaudry, and A. Ayala. 1998. Is Fas ligand or endotoxin responsible for mucosal lymphocyte apoptosis in sepsis? Arch. Surg. 133:1213. 16. Chung, C. S., Y. X. Xu, I. H. Chaudry, and A. Ayala. 1998. Sepsis induces increased apoptosis in lamina propria mononuclear cells which is associated with altered cytokine gene expression. J. Surg. Res. 77:63. 17. Hotchkiss, R. S., P. E. Swanson, J. P. Cobb, A. Jacobson, T. G. Buchman, and I. E. Karl. 1997. Apoptosis in lymphoid and parenchymal cells during sepsis: findings in normal and T- and B-cell-deficient mice. Crit. Care Med. 25:1298. 18. Cheadle, W. G., R. M. Pemberton, D. Robinson, D. H. Livingston, J. L. Rodriguez, and H. C. Polk, Jr. 1993. Lymphocyte subset responses to trauma and sepsis. J. Trauma 35:844. 19. Hotchkiss, R. S., P. E. Swanson, B. D. Freeman, K. W. Tinsley, J. P. Cobb, G. M. Matuschak, T. G. Buchman, and I. E. Karl. 1999. Apoptotic cell death in patients with sepsis, shock, and multiple organ dysfunction. Crit. Care Med. 27:1230. 20. Hotchkiss, R. S., K. W. Tinsley, P. E. Swanson, R. E. Schmieg, Jr., J. J. Hui, K. C. Chang, D. F. Osborne, B. D. Freeman, J. P. Cobb, T. G. Buchman, and I. E. Karl. 2001. Sepsis-induced apoptosis causes progressive profound depletion of B and CD4⫹ T lymphocytes in humans. J. Immunol. 166:6952. 21. Hotchkiss, R. S., K. C. Chang, P. E. Swanson, K. W. Tinsley, J. J. Hui, P. Klender, S. Xanthoudakis, S. Roy, C. Black, E. Grimm, et al. 2000. Caspase inhibitors improve survival in sepsis: a critical role of the lymphocyte. Nat. Immunol. 1:496.

22. Hotchkiss, R. S., P. E. Swanson, C. M. Knudson, K. C. Chang, J. P. Cobb, D. F. Osborne, K. M. Zollner, T. G. Buchman, S. J. Korsmeyer, and I. E. Karl. 1999. Overexpression of Bcl-2 in transgenic mice decreases apoptosis and improves survival in sepsis. J. Immunol. 162:4148. 23. Haendeler, J., U. K. Messmer, B. Brune, E. Neugebauer, and S. Dimmeler. 1996. Endotoxic shock leads to apoptosis in vivo and reduces Bcl-2. Shock 6:405. 24. Steinman, R. M., and Z. A. Cohn. 1973. Identification of a novel cell type in peripheral lymphoid organs of mice. I. Morphology, quantitation, tissue distribution. J. Exp. Med. 137:1142. 25. Steinman, R. M., and Z. A. Cohn. 1974. Identification of a novel cell type in peripheral lymphoid organs of mice. II. Functional properties in vitro. J. Exp. Med. 139:380. 26. Steinman, R. M., J. C. Adams, and Z. A. Cohn. 1975. Identification of a novel cell type in peripheral lymphoid organs of mice. IV. Identification and distribution in mouse spleen. J. Exp. Med. 141:804. 27. Rescigno, M., F. Granucci, and P. Ricciardi-Castagnoli. 1999. Dendritic cells at the end of the millennium. Immunol. Cell Biol. 77:404. 28. Inaba, K., and R. M. Steinman. 1985. Protein-specific helper T-lymphocyte formation initiated by dendritic cells. Science 229:475. 29. Schuler, G., and R. M. Steinman. 1985. Murine epidermal Langerhans cells mature into potent immunostimulatory dendritic cells in vitro. J. Exp. Med. 161:526. 30. Inaba, K., G. Schuler, M. D. Witmer, J. Valinksy, B. Atassi, and R. M. Steinman. 1986. Immunologic properties of purified epidermal Langerhans cells: distinct requirements for stimulation of unprimed and sensitized T lymphocytes. J. Exp. Med. 164:605. 31. Lipscomb, M. F., and B. J. Masten. 2002. Dendritic cells: immune regulators in health and disease. Physiol. Rev. 82:97. 32. Moser, M. 2003. Dendritic cells in immunity and tolerance: do they display opposite functions? Immunity 19:5. 33. Reis e Sousa, C., S. D. Diebold, A. D. Edwards, N. Rogers, O. Schulz, and R. Sporri. 2003. Regulation of dendritic cell function by microbial stimuli. Pathol. Biol. 51:67. 34. Traver, D., K. Akashi, M. Manz, M. Merad, T. Miyamoto, E. G. Engleman, and I. L. Weissman. 2000. Development of CD8␣-positive dendritic cells from a common myeloid progenitor. Science 290:2152. 35. Kronin, V., L. Wu, S. Gong, M. C. Nussenzweig, and K. Shortman. 2000. DEC205 as a marker of dendritic cells with regulatory effects on CD8 T cell responses. Int. Immunol. 12:731. 36. McLellan, A. D., and E. Kampgen. 2000. Functions of myeloid and lymphoid dendritic cells. Immunol. Lett. 72:101. 37. Liu, K., T. Iyoda, M. Saternus, Y. Kimura, K. Inaba, and R. M. Steinman. 2002. Immune tolerance after delivery of dying cells to dendritic cells in situ. J. Exp. Med. 196:1091. 38. Belz, G. T., G. M. Behrens, C. M. Smith, J. F. Miller, C. Jones, K. Lejon, C. G. Fathman, S. N. Mueller, K. Shortman, F. R. Carbone, and W. R. Heath. 2002. The CD8␣⫹ dendritic cell is responsible for inducing peripheral self-tolerance to tissue-associated antigens. J. Exp. Med. 196:1099. 39. Tinsley, K. W., M. H. Grayson, P. E. Swanson, A. M. Drewry, K. C. Chang, I. E. Karl, and R. S. Hotchkiss. 2003. Sepsis induces apoptosis and profound depletion of splenic interdigitating and follicular dendritic cells. J. Immunol. 171:909. 40. Ronchese, F., and I. F. Hermans. 2001. Killing of dendritic cells: a life cut short or a purposeful death? J. Exp. Med. 194:F23. 41. Hur, D. Y., D. J. Kim, S. Kim, Y. I. Kim, D. Cho, D. S. Lee, Y. Hwang, K. Bae, K. Y. Chang, and W. J. Lee. 2000. Role of follicular dendritic cells in the apoptosis of germinal center B cells. Immunol. Lett. 72:107. 42. Reis e Sousa, C., A. Sher, and P. Kaye. 1999. The role of dendritic cells in the induction and regulation of immunity to microbial infection. Curr. Opin. Immunol. 11:392. 43. De Smedt, T., B. Pajak, E. Muraille, L. Lespagnard, E. Heinen, P. De Baetselier, J. Urbain, O. Leo, and M. Moser. 1996. Regulation of dendritic cell numbers and maturation by lipopolysaccharide in vivo. J. Exp. Med. 184:1413. 44. De Smedt, T., B. Pajak, G. G. Klaus, R. J. Noelle, J. Urbain, O. Leo, and M. Moser. 1998. Antigen-specific T lymphocytes regulate lipopolysaccharideinduced apoptosis of dendritic cells in vivo. J. Immunol. 161:4476. 45. Hotchkiss, R. S., K. W. Tinsley, P. E. Swanson, M. H. Grayson, D. F. Osborne, T. H. Wagner, J. P. Cobb, C. Coopersmith, and I. E. Karl. 2002. Depletion of dendritic cells, but not macrophages, in patients with sepsis. J. Immunol. 168:2493. 46. Baker, C. C., I. H. Chaudry, H. O. Gaines, and A. E. Baue. 1983. Evaluation of factors affecting mortality rate after sepsis in a murine cecal ligation and puncture model. Surgery 94:331. 47. Oberholzer, A., C. Oberholzer, K. S. Bahjat, R. Ungaro, C. L. Tannahill, M. Murday, F. R. Bahjat, Z. Abouhamze, V. Tsai, D. LaFace, et al. 2002. Increased survival in sepsis by in vivo adenovirus-induced expression of IL-10 in dendritic cells. J. Immunol. 168:3412. 48. Fukuzuka, K., J. J. Rosenberg, G. C. Gaines, C. K. Edwards III, M. Clare-Salzler, S. L. MacKay, L. L. Moldawer, E. M. Copeland III, and D. W. Mozingo. 1999. Caspase-3-dependent organ apoptosis early after burn injury. Ann. Surg. 229:851. 49. Rescigno, M., S. Citterio, C. Thery, M. Rittig, D. Medaglini, G. Pozzi, S. Amigorena, and P. Ricciardi-Castagnoli. 1998. Bacteria-induced neo-biosynthesis, stabilization, and surface expression of functional class I molecules in mouse dendritic cells. Proc. Natl. Acad. Sci. USA 95:5229. 50. Cooper, M. A., T. A. Fehniger, A. Fuchs, M. Colonna, and M. A. Caligiuri. 2004. NK cell and DC interactions. Trends Immunol. 25:47.


References

SEPSIS AND THE DC


62.

63.

64.

65. 66.

67.

68.

69.

70.

splenocytes worsens survival, whereas adoptive transfer of necrotic splenocytes improves survival in sepsis. Proc. Natl. Acad. Sci. USA 100:6724. Di Nicola, M., S. Napoli, A. Anichini, R. Mortarini, L. Romagnoli, M. Magni, P. Matteucci, P. Baldassarri, I. Bersani, C. Carlo-Stella, and A. M. Gianni. 2003. Dendritic cell viability is decreased after phagocytosis of apoptotic tumor cells induced by staurosporine or vaccinia virus infection. Haematologica 88:1396. Ku, C. C., M. Murakami, A. Sakamoto, J. Kappler, and P. Marrack. 2000. Control of homeostasis of CD8⫹ memory T cells by opposing cytokines. Science 288:675. Miller, M. D., and M. S. Krangel. 1992. Biology and biochemistry of the chemokines: a family of chemotactic and inflammatory cytokines. Crit. Rev. Immunol. 12:17. Rollins, B. J. 1997. Chemokines. Blood 90:909. Sen, S., K. Muthu, L. K. He, A. Daud, S. Jones, R. Gamelli, and R. Shankar. 2003. Thermal injury and sepsis deplete precursor dendritic cells and alter their functions. J. Amer. Coll. Surg. 197(Suppl. 3):S39. De, A. K., K. Laudanski, and C. L. Miller-Graziano. 2003. Failure of monocytes of trauma patients to convert to immature dendritic cells is related to preferential macrophage-colony-stimulating factor-driven macrophage differentiation. J. Immunol. 170:6355. Stewart, D., W. B. Fulton, C. Wilson, C. L. Monitto, C. N. Paidas, R. H. Reeves, and A. De Maio. 2002. Genetic contribution to the septic response in a mouse model. Shock 18:342. Van der Poll, T., P. M. Jansen, W. J. Montegut, C. C. Braxton, S. E. Calvano, S. A. Stackpole, S. R. Smith, S. W. Swanson, C. E. Hack, S. F. Lowry, and L. L. Moldawer. 1997. Effects of IL-10 on systemic inflammatory responses during sublethal primate endotoxemia. J. Immunol. 158:1971. Remick, D. G., S. J. Garg, D. E. Newcomb, G. Wollenberg, T. K. Huie, and G. L. Bolgos. 1998. Exogenous interleukin-10 fails to decrease the mortality or morbidity of sepsis. Crit. Care Med. 26:895.


51. Fernandez, N. C., C. Flament, F. Crepineau, E. Angevin, E. Vivier, and L. Zitvogel. 2002. Dendritic cells (DC) promote natural killer (NK) cell functions: dynamics of the human DC/NK cell cross talk. Eur. Cytokine Network 13:17. 52. Scott, M. J., J. J. Hoth, S. A. Gardner, J. C. Peyton, and W. G. Cheadle. 2003. Natural killer cell activation primes macrophages to clear bacterial infection. Am. Surg. 69:679. 53. Godshall, C. J., M. J. Scott, P. T. Burch, J. C. Peyton, and W. G. Cheadle. 2003. Natural killer cells participate in bacterial clearance during septic peritonitis through interactions with macrophages. Shock 19:144. 54. Badgwell, B., R. Parihar, C. Magro, J. Dierksheide, T. Russo, and W. E. Carson III. 2002. Natural killer cells contribute to the lethality of a murine model of Escherichia coli infection. Surgery 132:205. 55. Jameson, S. C., and M. J. Bevan. 1995. T cell receptor antagonists and partial agonists. Immunity 2:1. 56. Sloan-Lancaster, J., B. D. Evavold, and P. M. Allen. 1994. Th2 cell clonal anergy as a consequence of partial activation. J. Exp. Med. 180:1195. 57. Yu, X. Z., S. D. Levin, J. Madrenas, and C. Anasetti. 2004. Lck is required for activation-induced T cell death after TCR ligation with partial agonists. J. Immunol. 172:1437. 58. Langenkamp, A., M. Messi, A. Lanzavecchia, and F. Sallusto. 2000. Kinetics of dendritic cell activation: impact on priming of TH1, TH2 and nonpolarized T cells. Nat. Immunol. 1:311. 59. Wysocka, M., S. Robertson, H. Riemann, J. Caamano, C. Hunter, A. Mackiewicz, L. J. Montaner, G. Trinchieri, and C. L. Karp. 2001. IL-12 suppression during experimental endotoxin tolerance: dendritic cell loss and macrophage hyporesponsiveness. J. Immunol. 166:7504. 60. Vendetti, S., J. G. Chai, J. Dyson, E. Simpson, G. Lombardi, and R. Lechler. 2000. Anergic T cells inhibit the antigen-presenting function of dendritic cells. J. Immunol. 165:1175. 61. Hotchkiss, R. S., K. C. Chang, M. H. Grayson, K. W. Tinsley, B. S. Dunne, C. G. Davis, D. F. Osborne, and I. E. Karl. 2003. Adoptive transfer of apoptotic

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