Basic Science Aspects
A Protective Role for Inflammasome Activation Following Injury
Osuka, Akinori; Hanschen, Marc; Stoecklein, Veit; Lederer, James A.
Department of Surgery (Immunology), Brigham and Women’s Hospital/Harvard Medical School, Boston, Massachusetts
Address reprint requests to James A. Lederer, PhD, Department of Surgery (Immunology), Brigham and Women’s Hospital/Harvard Medical School, 75 Francis Street, Boston, MA 02115. E-mail: firstname.lastname@example.org.
This study was supported by grant funding from the National Institutes of Health R01GM035633-23, RO1GM57664-09, and R33AI080565-03.
This study was submitted as 2011 Shock Society New Investigator Award Nominated Paper, Norfolk, Virginia.
The authors have no potential conflict of interest to report.
Received June 7, 2011
Accepted August 16, 2011
ABSTRACT: The inflammasome is activated in response to pathogen or endogenous danger signals and acts as an initiator and mediator of inflammatory reactions. In this study, we wished to identify whether the inflammasome is activated in vivo by injury. And if so, we wanted to characterize the kinetics, the immune cell distribution, and the functional impact of inflammasome activation on the injury response. Because caspase-1 activation is the final product of the inflammasome pathway, we used cleaved caspase-1 p10 and p20 as a measure for inflammasome activation in cells. We first developed a procedure to stain for caspase-1 p10 and p20 by flow cytometry (FACS) in lipopolysaccharide + adenosine triphosphate–stimulated spleen cells. This method for measuring caspase-1 activation was validated using FLICA (fluorochrome inhibitor of caspase), a fluorescently tagged specific binding reagent for activated caspase-1. Once validated by in vitro studies, we measured caspase-1 activation by FACS in immune cell subsets prepared from the lymph nodes and spleens of sham- or burn-injured mice at different time points. Lastly, the functional significance of inflammasome activation following burn injury was tested in mice treated with the specific caspase-1 inhibitor, AC-YVAD-CMK. The results of in vitro studies indicated that adenosine triphosphate and lipopolysaccharide stimulation induced significant caspase-1 activation in dendritic cells, macrophages, and natural killer (NK) cells. This approach also revealed caspase-1 activation in CD4 and CD8 T cells as well as B cells. We then measured caspase-1 activation in cells prepared from the lymph nodes and spleens of sham- or burn-injured mice. Significant caspase-1 activation was detected in macrophages and dendritic cells by 4 h after injury and peaked by day 1 after injury. FLICA staining confirmed that caspase-1 activation occurred in these cells at 1 day after injury. We also found significant injury-induced caspase-1 activation in NK cells, CD4 T cells, and B cells, but CD8 T cells did not demonstrate caspase-1 activation. Surprisingly, we found that blocking caspase-1 activation with AC-YVAD-CMK in vivo caused significantly higher mortality in burn-injured mice (P < 0.01). Taken together, these findings document that injury induces inflammasome activation in many immune cell subsets, but primarily in macrophages, and that inflammasome activation plays a protective role in the host response to severe injury.
ABBREVIATIONS: MFI — mean fluorescence intensity
LPS — lipopolysaccharide
ATP — adenosine triphosphate
Severe injuries induce an early and sustained inflammatory immune response that is detectable in more than 90% of patients following trauma (1). This postinjury hyperinflammatory state, the systemic inflammatory response syndrome (SIRS), depends on the severity of the initial insult and can lead to immunological complications or death by shock and multiorgan failure (1, 2). Although it is obvious that injury is the primary trigger of the host immune response to trauma, the molecular pathways responsible for initiating the host response to injury are not well defined. Given the observation that injury primes the innate immune system for both early and prolonged inflammatory phenotype, we hypothesized that activation of the inflammasome pathway might be a central feature of the injury response in mice.
The inflammasome is a multiprotein complex of signaling proteins that assembles following cellular responses to pathogen or host-derived danger signals, e.g., bacterial lipopolysaccharide (LPS) and extracelluar adenosine triphosphate (ATP) (3). Other signals that activate the inflammasome include uric acid crystals, silica, and aluminum or asbestos particles (4, 5). Some common protein elements of the inflammasome include ASC (apoptosis-associated speck-like protein containing a caspase recruitment domain) and caspase-1; however, distinct inflammasome-activating pathways trigger differing Nod-like receptor, including NALP1, NALP3, or IPAF (3). Thus, inflammasome assembly and activating pathways can be unique, depending on the specific initiating signal. Importantly, caspase-1 activation is a pivotal element of all known inflammasome activation pathways. Following inflammasome pathway activation, pro–caspase-1 is processed into two cleaved products called p10 and p20. Cleaved caspase-1 p10 and p20 then assemble to form interleukin 1 (IL-1)–converting enzyme to process the IL-1 family cytokines, IL-1β, IL-18, and IL-33, to active cytokines (6). Because the inflammasome is activated by pattern recognition receptors for pathogens or danger signals, it is believed that the inflammasome represents a common innate immune system recognition pathway for pathogen- or tissue-driven inflammatory responses (3, 7).
We initiated this study to find out if a severe injury caused by full-thickness 25% total body surface area burn might activate the inflammasome pathway in mice. To accomplish this, we first developed a flow cytometry method for measuring caspase-1 activation in cells by measuring the cleaved caspase-1 products, p10 and p20, in cells stimulated with LPS, ATP, or LPS + ATP. After establishing this new method, burn-injured mice were tested for caspase-1 activation in spleen and lymph node immune cell subsets at early (1, 2, or 4 h) and later (days 1, 3, or 7) time points after burn injury. We discovered that burn injury does indeed signal inflammasome activation by 1 day after injury. Moreover, the kinetics of injury-induced inflammasome activation correlated with detectable increases in circulating IL-1β. Inflammasome activation following burn injury returned to normal levels by 3 days after injury, suggesting that injury does not lead to sustained inflammasome activation. To measure the functional importance of injury-induced inflammasome activation, mice were treated with the irreversible caspase-1 inhibitor, AC-YVAD-CMK (YVAD), to measure phenotypic changes in the response to burn injury. We unexpectedly discovered that inhibiting caspase-1 activity in vivo caused significant mortality in burn-injured mice. This suggests that injury-induced inflammasome activation may be a protective mechanism for survival from severe injury. Collectively, the data reported here contribute new and fundamental information about the biology of inflammasome pathway activation following injury in a mouse model. The methods and information gained from this work could be applied to studies to investigate if trauma patients show inflammasome activation and whether the inflammasome activation phenotype or genotype affects the morbidity or mortality associated with trauma and its complications.
MATERIALS AND METHODS
Male C57BL/6J mice were purchased from The Jackson Laboratories (Bar Harbor, Me). All mice were maintained in our full-barrier animal facility under controlled temperature, humidity, and 12-h light-dark regimen. Mice were acclimatized for at least 1 week before use with a diet of standard chow and water ad libitum. Experiments were performed on mice aged 6 to 8 weeks. All animal protocols were approved by the Harvard Medical School Standing Committee on Animal Research and were found to be in accordance with the guidelines of the US Department of Agriculture and the National Institutes of Health (Bethesda, Md).
Cells were prepared in culture medium (C5); RPMI 1640 was supplemented with 5% heat-inactivated FCS, 1 mM glutamine, 10 mM HEPES, 100 μM nonessential amino acids, penicillin/streptomycin/fungizone, and 2.5 × 10−5 M 2-mercaptoethanol, all purchased from Gibco-Invitrogen (Grand Island, NY). Cells were fixed and permeabilized for FACS stains using phosphate-buffered saline with paraformaldehyde (PFA) and methanol purchased from Sigma Chemical Company (St Louis, Mo). Nonspecific binding was prevented by using mouse TruStain FcX reagent from BioLegend (San Diego, Calif). Antibodies specific for mouse CD3, CD4, CD8, CD11c, CD19, and F4/80 for FACS stains were purchased from BioLegend. Primary antibodies to stain intracellular cleaved caspase-1 (p10 and p20) were purchased from Santa Cruz Biotechnology (Santa Cruz, Calif). The detection of goat antibodies was performed by secondary incubation with PE-conjugated F(ab′)2, fragment of donkey anti–goat IgG, purchased from Santa Cruz Biotechnology. Ac-Tyr-Val-Ala-Asp-chloromethylketone (AC-YVAD-CMK) is a cell-permeable, irreversible inhibitor of caspase-1 and was purchased from Bachem Americas (Torrance, Calif). Fluorochrome inhibitor of caspase (FLICA) was purchased from Immunochemistry Technologies (Bloomington, Minn). Fluorescein isothiocyanate–conjugated annexin V is an apoptosis marker and was purchased from BD Biosciences (San Jose, Calif).
Mouse injury model
The mouse burn injury protocol was performed as described previously (8). Briefly, mice were anethestized by i.p. injection with ketamine (125 mg/kg; Fort Dodge Animal Health, Fort Dodge, Iowa) and xylazine (10 mg/kg; Lloyd Laboratories, Shenandoah, Iowa). The dorsal fur was shaved, and mice were placed in a plastic mold that exposes only 25% of their total body surface area. Injury was induced by immersing the exposed part of the dorsum to 90°C for 9 s in a water bath. This approach causes full-thickness and well-demarcated anesthetic burn injury; thus, mice were not given analgesics after the injury. Sham-treated mice underwent the same procedure, but were exposed to water at 24°C for 9 s. All animals were resuscitated with an i.p. injection of 1 mL of 0.9% pyrogen-free saline. The mortality from burn injury was less than 5%.
Lymph node and spleen cell preparation
At 1 h, 2 h, 4 h, 1 day, 3 days, or 7 days after injury, mice were killed by CO2 asphyxiation and the draining lymph nodes (axillary, brachial, inguinal), and the spleens were harvested. The lymph nodes and spleens were immediately minced using a sterile wire mesh, and cell suspensions were prepared in C5 medium. Red blood cells in splenic cells preparations were lysed using a red blood cell lysis buffer, and cell preparations were washed twice by centrifugation (200g, 10 min) in C5 medium and then strained to remove debris. Cells were plated in tissue culture–treated, round-bottom, 96-well plates (Corning Inc, Corning, NY) at 5 × 105 cells/well and immediately fixed by adding 0.15% PFA buffer.
After 10-min fixation in 0.15% PFA, cells were centrifuged to pellet and then permeabilized by adding ice-cold methanol. Cells were washed by centrifugation and then incubated with 1:50 dilution of mouse TruStain FcX reagent. Fixed and permeabilized cells were stained with APC-labeled antibodies specific for CD11c, CD19, F4/80, or NK1-1. CD4 and CD8 T cells were identified by FITC-labeled anti-CD3 antibody with either APC-labeled CD4 or CD8 antibodies. Apoptotic cells were identified by FITC-labeled annexin V. Annexin V has high affinity for phospholipid phosphatidylserine, which is translocated from the inner to the outer leaflet of the plasma membrane in apoptotic cells (9). Cells were incubated overnight with goat anti–mouse primary antibodies specific for cleaved caspase-1 p10 or cleaved caspase-1 p20. After washing by centrifugation, PE-labeled secondary donkey anti–goat IgG (F(ab′)2 fragment) was added as a last step in the staining procedure. Negative controls for cleaved caspase-1 detection were generated by performing the staining procedure without primary Ab. Stained samples were fixed in 0.3% PFA buffer, washed, and reconstituted in phosphate-buffered saline for flow cytometry using a Miltenyi MacsQuant instrument (Miltenyi Biotec, Auburn, Calif).
Goat anti–mouse p10- or p20-specific antibodies used in this study were tested in vitro to optimize staining conditions before use for in vivo studies. For these tests, whole spleen cell suspensions were cultured at 37°C, 5% CO2 at 5 × 105 cells per well in a round-bottom, 96-well plate. After 1-h preincubation, LPS at 100 ng/mL and ATP at 5 mM were added. Cells were processed for intracellular p10 and p20 caspase-1 staining and analyzed by flow cytometry as described above.
Measurement of caspase-1 activity
Cells were stimulated for 1 h with ATP and LPS before the addition of FAM-YVAD-FMK (5-carboxyfluorescein–Tyr-Val-Ala-Asp–fluoromethylketone, FLICA) according to the manufacturer’s instructions (Immunochemistry Technologies). FLICA+ cells were detected and measured by FACS.
Culture supernatants were harvested at 48 h after spleen cell stimulation and tested for IL-1α, IL-1β, IL-2, IL-4, IL-6, IL-10, IL-12p40, IL-12p70, IL-13, IL-18, IL-33, interferon γ (IFN-γ), and tumor necrosis factor α (TNF-α) by our custom-made Luminex multiplex cytokine-detection bead assay using a Luminex 200 instrument (Luminex Corp, Austin, Tex). Blood samples were harvested by cardiac puncture in EDTA anticoagulant solution (0.1 mL of 169 mM EDTA) from sham or burn mice and were centrifuged at 1,000g for 10 min to prepare plasma. Plasma samples were also tested for cytokine levels using our Luminex bead assay platform. Our cytokine assays routinely attain sensitivity ranges of 1 to 25,000 pg/mL.
The results presented in the study were analyzed by two-way analysis of variance (ANOVA) and Tukey multiple-comparisons tests or Mann-Whitney U test using Prism 5.0 software (GraphPad Software, Inc, San Diego, Calif). Survival differences were evaluated by the log-rank test. P < 0.05 was considered significant. Data are plotted as mean (SEM) values.
Optimization of intracellular cleaved caspase-1 detection by flow cytometry
Because caspase-1 cleavage is a central component of inflammasome activation, we needed to develop a procedure to accurately measure and quantify caspase-1 activation in cells. Therefore, we developed and optimized a flow cytometry–based method for detecting the cleaved caspase-1 end products, p10 and p20, in cells. We used an in vitro approach to develop this method. Spleen cells were prepared from mice and stimulated with LPS, ATP, or LPS + ATP—stimuli that are strong inducers of caspase-1 cleavage. After 1-h stimulation, cells were fixed and stained for intracellular p10 and p20 levels as well as cell surface markers for specific immune cell subsets. The FACS histograms shown in Figure 1A illustrate the level of p10 and p20 staining in total splenocytes, CD11c+ dendritic cells (DCs), and F4/80+ macrophages. The plots include unstained controls, unstimulated cells, and cells stimulated with ATP + LPS for 1 h. These representative histograms illustrate induced changes in p10 expression as measured by an increase in fluorescence intensity in unstimulated versus stimulated spleen cells and FACS-gated immune cell subsets—macrophages and DCs. Moreover, they demonstrate our ability to detect caspase-1 cleavage and activation in cells that are stimulated to activate the inflammasome.
To validate our FACS method, we measured caspase-1 activation by an alternative method using a cell-permeable fluorescent probe called FLICA, which specifically binds to activated caspase-1 (10). Spleen cells were activated with LPS and ATP for 1 h in the presence of FLICA. Cells were then stained to identify macrophages or DCs and analyzed by FACS. The FACS dot plots shown in Figure 1 B illustrate that in vitro stimulation with LPS and ATP caused detectable caspase-1 activation by the FLICA method. Moreover, the FLICA stain data demonstrate a constitutive level of activated caspase-1 in macrophages and DCs that is increased in response to LPS and ATP stimulation that is similar to what we observed by the FACS staining method. These data further validate the accuracy of our FACS method to detect activated caspase-1 in cells.
Using this FACS staining approach, we assessed the kinetics of caspase-1 cleavage in LPS and ATP stimulated spleen cells. Spleen cells were stimulated with LPS, ATP, or LPS + ATP for 15 min, 30 min, 1 h, 2 h, or 4 h and then stained for p10 and p20 (Fig. 2A). As shown, caspase-1 activation occurred when cells were stimulated with ATP, but not LPS alone. However, the combination of LPS and ATP caused sustained caspase-1 activation that was still detectable at 4 h after stimulation, whereas ATP stimulation alone showed only early caspase-1 activation. The data plotted in Figure 2B show caspase-1 p10 or p20 levels in macrophages, DCs, natural killer (NK) cells, CD4 T cells (CD3+/CD4+ cells), CD8 T cells (CD3+/CD8+ cells), and B cells (CD19 + cells) at 1 h after LPS + ATP. Surprisingly, we found evidence for significant caspase-1 activation in all of these immune cell subsets. However, p20 staining was not detected in CD4 T and B cells.
After finding in vitro conditions that induced significant caspase-1 activation, we wished to determine if activating caspase-1 in cells by ATP or LPS + ATP stimulation could induce caspase-1–dependent cytokine production. Spleen cells were stimulated with LPS, ATP, or LPS + ATP, and supernatants were tested for cytokine levels by multiplex cytokine assays. The data plotted in Figure 3 illustrate that these different stimuli induced different levels and types of cytokines. As anticipated, LPS stimulation alone induced significant levels of TNF-α and IL-6. In contrast, IL-1β was not induced by LPS but was significantly induced by ATP alone and combined stimulation with LPS + ATP. Surprisingly, the addition of ATP markedly suppressed TNF-α and IL-6 production, indicating that extracellular ATP stimulation suppresses TLR4-induced cytokines. Most important, the cytokine profiling dataconfirm that the experimental conditions that induced caspase-1 activation in cells also specifically induced high-level production of the caspase-1–dependent cytokines, IL-1β and IL-18.
Time-course study for caspase-1 activation in innate immune cell subsets after burn injury
Following sham or burn injury, cells were prepared from the draining lymph nodes or spleens and FACS stained for CD11c, F4/80, NK 1.1, and cleaved caspase-1 p10 or p20 at 1 h, 2 h, 4 h and at 1, 3, 7 days after injury. We observed an increase in p10 in lymph node macrophages by 2 h after injury, whereas significant increases in p10 in splenic macrophages and DCs were seen at 1 day after injury (Fig. 4A). Also, this injury-induced increase in p10 levels in splenic macrophages and DCs peaked by day 1 after injury and returned to normal levels by 3 days after injury. Macrophages appeared to express the highest levels of p10 after injury as compared with DCs and NK cells. Interestingly, burn injury also caused an increase in p20 levels in macrophages, DCs, and NK cells, but the change was not statistically significant. To confirm and extend our findings, we measured caspase-1 activation in these spleen cell subsets with FLICA after sham versus burn injury. As shown in Figure 4B, we detected an increase in FLICA+ macrophages, DCs, and NK cells prepared from the spleens of mice at 1 day after burn as compared with sham injury. The FLICA stains support the FACS results but also show burn-induced caspase-1 activation in NK cells.
Next, we wished to determine which immune cell subsets show injury-induced caspase-1 activation. To accomplish this, we identified DCs, macrophages, NK cells, CD4 and CD8 T cells (CD3+), and B cells by cell-surface staining and measured p10 or p20 levels by intracellular staining. As shown in Figure 5, these data indicate that burn injury induced significant caspase-1 activation in all but CD8+ T cells. Again, innate immune cell populations showed higher caspase-1 activation than CD4 T or B cells. In addition, we examined whether cells showing caspase-1 activation were undergoing apoptosis by measuring annexin V binding on p10 or p20 expressing immune cell subsets. We observed significant increases in annexin V binding on most immune cell subsets prepared from the spleens of mice at 1 day after burn injury—the exception was CD8 T cells.. However, annexin V–positive cells did not show higher levels of p10 or p20 expression (Table 1). This finding suggests that the cells showing increased caspase-1 activation following burn injury are not undergoing apoptosis.
Finally, we wanted to sequentially measure circulating cytokine levels in sham and burn mice to determine if the kinetics of caspase-1 activation might coincide with the production of IL-1β, IL-18, and IL-33. We prepared plasma samples from mice at 1, 2, and 4 h and at 1, 3, 7 days after sham or burn injury, and these samples were screened for cytokine levels by multiplex cytokine assays. The multiplex assay approach allowed us to screen for many different cytokines, but only IL-1β, IL-6, and TNF-α were found to be induced by burn injury at these time points (Fig. 6). Interestingly, we did not detect significant levels of these cytokines until 1 day after injury. The levels of IL-1β and IL-6 returned to normal by 7 days after burn injury, but TNF-α levels remained high. These data indicate that injury-induced IL-1β coincides with the kinetics of burn-induced caspase-1 activation and with burn-induced increases in other proinflammatory mediators.
Functional significance of inflammasome activation following burn injury
To determine the functional significance of inflammasome activation after burn injury, we treated mice with an irreversible caspase-1–specific inhibitor called AC-YVAD-CMK (YVAD). This inhibitor blocks inflammasome assembly by binding specifically to caspase-1 subunits (11). Dose-response studies were performed by treating mice with 1, 5, or 10 mg/kg at 2 h before burn injury and measuring caspase-1 activation by FACS at 1 day after injury. As shown in Figure 7A, treating mice with 10 mg/kg YVAD potently blocked caspase-1 activation as judged by p10 and p20 staining in spleen cells, whereas lesser doses did not effectively block caspase-1 activation. After determining an effective dose for YVAD, we tested the impact of blocking inflammasome activation on the burn injury response in mice. Unexpectedly, we found that YVAD treatment caused significantly higher mortality in burn-injured mice (P < 0.01) (Fig. 7B). To confirm that inhibiting caspase-1 activation caused higher mortality in burn-injured mice, we performed an additional YVAD dose study comparing 1-, 5-, and 10-mg/kg treatments. We observed that 10 mg/kg–treated mice showed high mortality, whereas mice treated with lower and less effective doses of 1 or 5 mg/kg showed no burn injury–induced mortality (Fig. 7C). Taken together, these results support the conclusion that injury-induced inflammasome activation is protective.
Given the high mortality in YVAD-treated burn mice, we wanted to determine why YVAD-treated burn mice died. Blood was collected from YVAD-treated mice at 1 day after burn injury to test for evidence of bacteremia by performing blood cultures. No bacteria were detected in blood samples prepared at 1 or 2 days after burn injury in YVAD-treated or untreated mice (data not shown). In addition, we treated groups of YVAD-treated burn mice with oral antibiotics (sulfamethoxazole/trimethoprim) and found no significant difference in survival between untreated and antibiotic-treated mice (data not shown, n = 16 mice per group). Lastly, we measured plasma cytokine levels at 1 day after burn injury in untreated versus YVAD-treated mice. As suspected, IL-1β was significantly suppressed by YVAD treatment (Fig. 8). However, IL-6 and IL-33 were significantly higher in plasma prepared from YVAD-treated as compared with untreated burn mice. Moreover, we found significantly lower levels of counter-inflammatory–type cytokines, IL-4 and IL-13, in plasma from YVAD-treated burn mice as compared with untreated burn mice. These findings suggest that blocking inflammasome activation by YVAD treatment may have caused higher inflammation and organ damage in burn-injured mice by increasing inflammatory and reducing counter-inflammatory mediators. The combination of increased inflammation with no evidence of bacteremia suggests that the inflammasome plays a key role in controlling the immune response to severe injury.
It is generally accepted that burn injury and trauma prime the innate immune system to enhance inflammatory responses to pathogen-associated molecular signals; however, the molecular pathways responsible for injury-induced immune system activation are not well defined (12, 13). Research addressing how the immune system responds to injured and dead cells has identified an expanding list of molecules that signal inflammatory-type responses by receptor-mediated signals (4, 14–16). These endogenous molecules are collectively referred to as danger molecules or damage-associated molecular patterns because they are generated when there is cell death or damage caused by pathogens or injury (17, 18). A central feature of danger signals is that they evoke an inflammatory-type response to initiate both innate and adaptive immune responses (19). The inflammasome was identified as a central feature of the danger response because most of the extracellular or intracellular signals that activate the inflammasome are considered danger molecules or alarmins (14, 20–22). This study was initiated to test the hypothesis that the inflammasome is activated in immune cells in an in vivo injury model because little is known about the in vivo nature of injury-induced inflammasome activation. Moreover, we wished to test the hypothesis that sustained inflammasome activation following burn injury in mice might contribute the development of SIRS and the two-hit response that occurs at later time points after injury. Our findings presented here establish that burn injury does induce inflammasome activation, but that its activation is shorter lived than anticipated—peak inflammasome activation occurred by 1 day after injury and returning to normal levels at 3 and 7 days after injury. This observation is novel and indicates that injury induces early but not sustained inflammasome activation. Thus, the inflammasome appears to be involved in initiating the host response to injury, but may not contribute to the SIRS and two-hit response phenotype that occurs after severe injury.
As part of this study, we developed a new FACS-based method to detect caspase-1 activation in cells. To our knowledge, this is the first study to characterize caspase-1 activation in cells by measuring changes in p10 and p20 levels by flowcytometry. A major advantage of using FACS to measure caspase-1 activation in cells versus Western immunoblot is that it allows for identifying the immune cell subsets showing caspase-1 or inflammasome activation. We used in vitro experiments to develop and validate our FACS-based method because it allowed us to stimulate inflammasome activation under conditions that are well known to activate the inflammasome. Preliminary studies were performed to optimize the cell fixation conditions, staining antibody concentrations, and stain incubation times. We found that light fixation with 0.15% PFA, followed by methanol fixation provided the best fixation conditions for these stains. Interestingly, we also found that overnight incubation with p10 or p20 antibodies was needed to generate the best staining signal. As illustrated in Figure 1, spleen cells stimulated with LPS + ATP showed detectable shifts in MFI for p10 and p20 by 1 h after stimulation. FACS staining was then used to demonstrate that caspase-1 activation by LPS + ATP stimulation is not restricted to macrophages. Interestingly, we observed significant induction of either cleaved caspase p10 or p20 in DCs, NK cells, CD4 T cells, CD8 T cells, and B cells at 1 h after LPS + ATP stimulation. Macrophages, DCs, and NK cells showed higher level constitutive p10 and p20 than T or B cells. The observed caspase-1 activation in NK cells, T cells, and B cells is novel and suggests that caspase-1 activation occurs in a wider assortment of immune cell types than previously appreciated. Burn injury also induced caspase-1 activation in these similar immune cell subsets. Again, the development of a FACS-based method to detect the p10 and p20 subunits of caspase-1 made it possible to discover that caspase-1 activation can occur in many different immune cell subsets.
Cytokine multiplex assays were used to measure ATP-, LPS-, or ATP + LPS–induced cytokine production. Lipopolysaccharide stimulation alone induced significant TNF-α and IL-6 production by spleen cells, whereas IL-1β and IL-18 were induced by ATP or ATP + LPS stimulation. This finding confirmed that activating caspase-1 in vitro caused significant production of the caspase-1–dependent cytokines, IL-1β and IL-18. While performing these experiments, we also found that adding ATP to LPS-stimulated spleen cells blocked LPS induced TNF-α and IL-6 production. This result was surprising and suggests that extracellular ATP has potent anti-inflammatory activity that may be independent of its caspase-1–activating activity. The processing or degradation ATP to adenosine and adenosine receptor signaling may be responsible for suppressing TNF-α and IL-6 (23).
The results from time-course injury studies revealed that burn injury induces inflammasome activation as measured by caspase-1 activation. Given that inflammasome activation occurred early after burn injury, we anticipated that it may be involved in initiating the inflammatory response to burn injury. This observation also suggested to us that blocking inflammasome activation in vivo might evoke a beneficial response to injury by reducing inflammation-driven host responses as has been shown in other models of tissue injury (24–26). However, we found that treating mice with the caspase-1–specific inhibitor AC-YVAD-CMK showed a significant increase in injury-induced mortality by 4 days after burn injury. This finding led us to conclude that inflammasome activation and, in particular, caspase-1 activation play an unanticipated protective role in the host response to injury. The in vivo dose of AC-YVAD-CMK used to inhibit caspase-1 activation was found to be an effective dose as judged by inhibition of caspase-1 activation and IL-1β production following burn injury. Furthermore, we observed that suboptimal inhibitory doses of less than 10 mg/kg did not cause increased mortality and did not inhibit caspase-1 activation in burn-injured mice. The results of these dose-response studies further confirm that inflammasome activation by injury promotes better survival from severe injury. Other reports that used AC YVAD-CMK to block caspase-1 in vivo did not describe changes in mortality, but did demonstrate that blocking caspase-1 activation reduced inflammatory responses to tissue injury (24, 27). Thus, our findings are unique among others that used AC-YVAD-CMK to block inflammasome activation in vivo.
We addressed potential causes for increased mortality in burn-injured, YVAD-treated mice. First, we suspected that these mice may die of infection due to reduced antimicrobial immunity. However, we did not detect bacteria in the blood of burn + YVAD–treated mice. In addition, antibiotic treatment did not significantly protect burn + YVAD–treated mice. These findings suggest that bacteremia was not the cause of death. Subsequently, we tested for changes in plasma cytokine levels in burn as compared with AC-YVAD-CMK–treated mice. We discovered that YVAD treatment significantly blocked burn-induced IL-1β as well as IL-4 and IL-13. In contrast, IL-33 and IL-6 levels were significantly higher in YVAD-treated burn mice. These cytokines have been shown to be important mediators of the acute-phase response, tissue damage, and shock (28–30). The cytokine profile data suggest that mice may be dying of cytokine-induced shock or organ damage.
In summary, the studies presented here demonstrate that severe injury caused by burn injury induces early, but not late or sustained inflammasome activation in a variety of immune cell subsets. Thus, the inflammasome does not appear to play a direct role in mediating the SIRS and the two-hit response to injury. Instead, we discovered that injury-induced inflammasome activation is protective. Because we rarely observe greater than 5% direct mortality in our mouse injury model, this suggests that inflammasome-activating signals promote a beneficial response to severe injury. Therefore, it is possible that treatment strategies that target inflammasome activation pathways could be used to protect patients from organ damage and shock caused by severe injuries. For this reason, future work will seek to define the pathways and factors that induce inflammasome activation following injury.
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