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,
Departments of
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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
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, 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 polymicrobial sepsis (cecal ligation and puncture (CLP))4 (9, 10, 11, 12, 13, 14, 15, 16, 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, 25, 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, 29, 30, 31, 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, 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, 35, 36, 37, 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, 40, 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, 43, 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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Materials and Methods |
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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 x 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 µl/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-µm 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 x 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 x 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, PE-conjugate hamster anti-mouse CD11c mAb, 7-aminoactinomycin D (7-AAD), 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 1x 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 µl of 1x 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 x 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 x 104 CD11c+/7-AAD– events were collected per tube. For annexin V-stained samples, 1 x 104 CD11c+ events were collected. A total of 1 x 105 events was collected for Th cell analysis, with >1 x 104 CD3+/CD4+/7-AAD– events being collected.
Flow cytometry 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.
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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-µm 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 (x100) was evaluated in each sample.
Data analysis
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).
Statistical analysis
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.
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Results |
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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).
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There was also a significant increase (p < 0.05) in the percentage of apoptotic and dead DCs in the mesenteric and inguinal nodes 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 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).
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No significant differences were observed in the percentage of mature DCs (MHCIIhigh and CD86high) in the lymph nodes of CLP- vs sham-operated mice (Table I). This was true in the local and distant lymph nodes at 6, 12, and 24 h postoperation.
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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.
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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 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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Significant decreases were seen in the CD3+CD4+ T cell population 6–12 h post-CLP, while losses in the DC population were not observed until 12–24 h after induction of polymicrobial sepsis (Fig. 8).
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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 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 decrease in the cell population rather than an increase in another cell population.
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Discussion |
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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 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, 51, 52, 53, 54).
DCs may drive apoptosis in selected lymphocyte populations (36, 39, 40, 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, 64, 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 mouse used. Different inbred strains of mice have different responses and outcomes to sepsis (68, 69, 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.
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Footnotes |
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1 This work was supported in part by Grants R37 GM-40561-15, and R01 GM-63041-03, 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 Sciences-funded 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: moldawer{at}surgery.ufl.edu
4 Abbreviations used in this paper: CLP, cecal ligation and puncture; 7-AAD, 7-aminoactinomycin D; DC, dendritic cell.
Received for publication January 21, 2004. Accepted for publication June 29, 2004.
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