Sepsis is a severe pathologic process caused by overwhelming infection of the bloodstream by toxin-producing bacteria; the mortality rate is 20% to 40% (1). Previous studies have demonstrated a critical role for tumor necrosis factor (TNF)-α and interleukin (IL)-1 in the pathogenesis of sepsis, as well as for other cytokines such as IL-12, IL-18, and interferon (IFN)-γ (2). However, the usefulness of anti-TNF-α and IL-1 receptor-antagonist therapies in human sepsis has been disappointing (2). Although there may be multifactors for this failure, a major difficulty in developing therapeutics that target at cytokines is that they are released early in the development of sepsis or systemic inflammatory response syndrome (SIRS). This fact has led to the search for delayed mediators of sepsis that contribute to tissue damage.
Recently, high-mobility group box 1 protein (HMGB1) has been identified as a novel inflammatory cytokine and a late mediator of endotoxin lethality (3, 4). HMGB1 is a member of the HMGB family, and its localization in most cells is in the nucleus (5). It contains two homologous DNA-binding domains, HMGB boxes A and B, and an acidic C-terminal tail (6). HMGB1 is secreted by activated monocytes/macrophages and is released passively by necrotic and damaged cells (3, 7). However, HMGB1 is not released by apoptotic cells even after subsequent secondary necrosis (7). Monocytes/macrophages and pituicytes stimulated by lipopolysaccharide (LPS), TNF-α, IL-1, or IFN-γ release HMGB1 as a late mediator (3, 8, 9). In mice, injection of HMGB1 causes toxic shock (3). Anti-HMGB1 antibodies confer significant protection against delayed endotoxin lethality, even when antibody dosing is initiated after the acute-phase cytokine responses have peaked and resolved (3). HMGB1 is also shown to cause acute lung inflammation (10). Moreover, increased levels of HMGB1 have been implicated in patients with sepsis and other major inflammatory diseases, including rheumatoid arthritis (3, 11, 12). Secreted HMGB1 also stimulates the synthesis and release of proinflammatory cytokines by monocytes and human microvascular endothelial cells (13, 14). HMGB1 triggers activation of the members of the mitogen-activated protein kinase (MAPK) pathway and subsequent activation of NF-κB (14-17). In brief, these studies indicate that HMGB1 plays an important role in the pathogenesis of sepsis. However, it is not entirely clear on the expression, subcellular distribution, and kinetic changes of HMGB1 when murine macrophages are exposed to LPS in vitro.
Heat shock response (HSR), or stress response, is the most primitive and most conserved form of cell response to stress (18, 19). It is nearly universal in all prokaryotes and eukaryotes. The cellular response to stress is represented at the molecular level by the induction of heat shock protein (HSP) synthesis (18). The HSP70 family is the most abundant HSP and mainly includes the constitutive cytosolic Hsc70 (or Hsp73) and the stress-induced cytosolic Hsp70 (or Hsp72). HSPs are widely believed to function as molecular chaperones involved in repair, transport, folding, and unfolding of various intracellular proteins (20). HSPs also play important roles in antigen presentation, cross-presentation, and tumor immunity (21). Recently, it has been found that HSR has anti-inflammatory effects. HSR reduces mortality in experimental models of septic shock, endotoxemia, and adult respiratory distress syndrome (22, 23), and can regulate expression of proinflammatory and anti-inflammatory genes such as TNF-α, IL-1, IL-12, IL-10, and IL-18 (24-26). To date, there has not been a report involving the effect of heat shock pretreatment on the expression and release of HMGB1 in sepsis or endotoxemia.
Therefore, our present study was to fully characterize the kinetic changes of HMGB1 in vitro in response to LPS and to explore the influence of heat shock response on the expression, release, and translocation of HMGB1 induced by LPS.
MATERIALS AND METHODS
BALB/c murine macrophage-like RAW 264.7 cells, obtained from the Shanghai Type Culture Collection (Shanghai, China), were cultured in RPMI medium 1640 (Life Technologies, Rockville, MD) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM glutamine, and antibiotic-antimycotic mix in a humidified incubator with 5% CO2 and 95% air. At 70% confluency, RAW 264.7 cells were removed mechanically and were resuspended in serum-free Opti-MEM I medium (Life Technologies). After preincubation for 2 h, RAW 264.7 cells (5 × 106 cells) were treated with LPS (Esherichia coli 0111:B4; Sigma, St. Louis, MO).
Heat shock treatment
RAW 264.7 cells (5 × 106 cells) were sealed in screw cap flasks containing an atmosphere of 5% CO2/95% air. These flasks were then immersed completely in a water bath with a measured temperature of 42.5°C. By using this protocol, the medium within the flask reached 42.5°C within 5 min of immersion. After 1 h of immersion, cells were left at 37°C for 12 h and were then treated with LPS.
Preparation of cellular extracts
At the appropriate time after LPS treatment, cells were harvested and washed twice with cold phosphate-buffered saline (PBS); nuclear and cytoplasm extracts were prepared according to the method of Edgar et al. (27). Briefly, the cell pellet was resuspended in 400 μL of cold buffer A (10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM dithiothreitol [DTT], and 0.5 mM phenylmethylsulfonyl fluoride [PMSF]). The cells were allowed to swell on ice for 15 min, after which 25 μL of a 10% solution of NP-40 was added and the tube was vigorously vortexed for 10 s. The homogenate was centrifuged for 30 s in a microfuge. The supernatant contained cytoplasm. The nuclear pellet was resuspended in 50 μL of ice-cold buffer B (20 mM HEPES, pH 7.9, 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, and 1 mM PMSF) and the tube was vigorously rocked at 4°C for 15 min on a shaking platform. The nuclear extract was centrifuged for 5 min in a microfuge at 4°C, and the supernatant was frozen in aliquots at −80°C. The protein content of the different fractions was determined by the Bradford assay method.
Determination of HMGB1 in the culture medium
After corresponding treatment, cell culture-conditioned medium was filtered through Millex-GP (Millipore, Bedford, MA) to clear the samples from cell debris and macromolecular complexes. Samples were then concentrated 40-fold with Amicon Ultra-4-10000 NMWL (Millipore). Centrifugation conditions were fixed angle (35 degrees)and 7500g for 20 min at 4°C.
Western blot analysis
Protein from the subcellular fractions or concentrated supernatants or cell lysates was added to 2× SDS sample buffer (100 mM Tris-HCl, pH 6.8, 4% [w/v] SDS, 20% glycerol, 200 mM DTT, and 0.1% [w/v] bromphenol blue), boiled in a water bath for 5 min, analyzed in 10% SDS-PAGE, and transferred to a nitrocellulose membrane (Promega, Madison, WI). The membrane was routinely stained with Ponceau S solution (2 μg/mL; Sigma) for 5 min to determine transfer efficiency and protein loading levels per track (data not included). Membranes were then washed free of Ponceau S in phosphate-buffered saline (PBS), blocked with blocking buffer at room temperature for 6 h, and then incubated 2 h at 25°C with primary antibody (rabbit anti-HMGB1 polyclonal antibody [1:1000; BD Biosciences, Mountain View, CA], mouse anti-Hsp72 monoclonal antibody [1:1000; StressGen, Victoria, British Columbia, Canada], rat anti-β-tubulin monoclonal antibody [1:1000; Sigma], and mouse antiproliferating cellular nuclear antigen [PCNA] monoclonal antibody [1:1000; Upstate Biotechnology, Lake Placid, NY]), followed by peroxidase-conjugated secondary antibody immunoglobulin (Ig) G (anti-rabbit, anti-rat, or anti-mouse [1:1000; Boster Biotech, Wuhan, China]) for 1 h at 25°C. The signals were visualized by DAB detection (Boster Biotech) according to the manufacturer's instruction and the bands of protein were scanned and counts quantitated with the Band Leader software (Shanghai, China). Anti-β-tubulin antibody (28) and anti-PCNA (29) served as loading control.
After LPS stimulation, total RNA was extracted with Trizol reagent (Gibco, Grand Island, NY). Total RNA (1 μg) was then reversely transcribed into cDNA in a 20-μL volume including 20 U AMV (TaKaRa, Otsu, Japan). The internal mRNA standard was housekeeping gene GAPDH. Primers were as follows: HMGB1, sense: 5′-ATGGGCAAAGGAGATCCTA-3′, antisense: 5′-ATTCATCATCATCATCTTCT-3′; GAPDH, sense: 5′-AAGCCCATCACCATCTTCCA-3′, antisense: 5′-CCTGCTTCACCACCTTCTTG-3′. Each 25-μL PCR volume contained 1 μL of RT mix template, 1 U of Taq DNA polymerase (TaKaRa), and 10 pM each primer (sense and antisense). The HMGB1 cycling condition was 30 s of denaturation at 95°C, 30 s of annealing at 58°C, and 30 s of elongation at 72°C over 25 cycles for all reactions. The GAPDH cycling condition was 30 s of denaturation at 95°C, 30 s of annealing at 59°C, and 30 s of elongation at 72°C over 22 cycles for all reactions.
RAW 264.7 cells (2 × 106 cells), cultured on glass coverslips, were fixed in 4% formaldehyde for 30 min at room temperature before detergent extraction with 0.1% Triton X-100 for 10 min at 4°C. Coverslips were saturated with PBS containing 2% bovine serum albumin (BSA) for 1 h at room temperature and were processed for immunofluorescence with rabbit anti-HMGB1 polyclonal antibody (1:50; BD Biosciences) followed by Cy3-conjugated sheep anti-rabbit Ig (1:100; Sigma), and Hochest 33258 (1 μg/mL; Sigma). Between all incubation steps, cells were washed three times for 3 min with PBS containing 0.2% BSA. Coverslips were mounted on slides using Movio (Sigma). Fluorescence signals were analyzed by fluorescent microscopy (Olympus, Tokyo, Japan).
Evaluation of cell viability
The viability of RAW 264.7 cells was assessed before experiments or after incubation with the various stimuli. Less than 5% of cells incubated with LPS (100-1000 ng/mL), heat shock (at 42.5°C for 1 h), or heat shock pretreatment exhibited an altered morphology or uptake of trypan blue, indicating that membrane integrity was not disrupted (data not included).
Data in the figures and text are expressed as means ± SEM. Significance of differences between groups was determined by two-tailed Student's t test or Fisher's least significant difference test, as indicated. P < 0.05 is considered significant.
HMGB1 protein and mRNA expression after LPS treatment
To evaluate whether LPS affected the production of HMGB1, we determined the kinetics of HMGB1 protein and mRNA expression. Incubation with LPS (500 ng/mL) did not significantly influence total intracellular HMGB1 protein levels (Fig. 1A) by Western blot analysis. However, HMGB1 mRNA levels were affected by LPS treatment. Figure 1B shows a representative result from three separate experiments. A distinct decrease of HMGB1 mRNA expression was observed at 18 and 24 h after LPS (500 ng/mL) stimulation in RAW 264.7 cells compared with untreated controls (Fig. 1B).
HMGB1 relocalizes from the nucleus to the cytoplasm in macrophage activated by LPS
The intracellular localization of HMGB1 in RAW 264.7 cells freshly isolated or treated with LPS was investigated by immunocytochemical analysis. Cells were stained with anti-HMGB1 antibodies. Nonstimulated macrophages (Fig. 2A) displayed a strong staining for HMGB1 in the nucleus. Twenty hours after stimulation with LPS (500 ng/mL; Fig. 2C), HMGB1 appeared to move from the nucleus, which became weakly stained, to the cytoplasm, which displayed a strong staining.
To investigate the distribution of HMGB1, normal and LPS-activated macrophages were subcellular fractionated and the distribution of HMGB1 was analyzed by Western blot. As shown in Figure 3, a translocation of HMGB1 from the nucleus to the cytoplasm was observed at 20 h after LPS (500 ng/mL) administration despite of different concentrations. In normal macrophages, most of the HMGB1 was found in the nucleus, with a small amount of HMGB1 in the soluble cytoplasmic fraction. In LPS-activated macrophages, less HMGB1 was detected in the nuclear fraction and more HMGB1 was detected in the cytoplasm compared with control.
Release of HMGB1 by LPS-activated macrophages is a late event
To evaluate the potential role of LPS in stimulating HMGB1 release, cultured RAW 264.7 cells were stimulated by the addition of LPS, and the levels of HMGB1 in the culture medium were subsequently measured by Western blot analysis. HMGB1 could not be detected in the culture medium in the absence of LPS stimuli, but HMGB1 release started to be detected at 8 h and was still increasing at 12 and 24 h after LPS (500 ng/mL) treatment (Fig. 4). A wide concentration range of LPS (100, 500, and 1000 ng/mL) induced significant release of HMGB1 in RAW 264.7 cells. HMGB1 was released from LPS-activated RAW 264.7 cells in a time- and dose-dependent manner (Fig. 4, A and B).
Effects of HSR on expression of Hsp72 and HMGB1
In previous studies, exposure of cells to stressful conditions activates the HSR via the induction of HSPs. Among the HSPs, we investigated the kinetics of Hsp72 expression in heat-treated RAW 264.7 cells. In the three independent experiments, after the treatment of cells at 42.5°C for 1 h, Hsp72 protein was measured at different recovery times of 0, 2, 4, 8, 12, and 24 h. As shown in Figure 5A, the expression level of Hsp72 in normal control cells was too low to be detected. In contrast, Hsp72 could be detected at 4 h of recovery after heat shock pretreatment and the maximal level was observed at 12 h (Fig. 5A), consistent with the cells undergoing the HSR. Previous studies revealed that during HSR, different heat shock genes are activated, whereas most other genes are down regulated. Thus, the influence of heat shock per se on expression of HMGB1 was investigated. As shown in Figure 5B, HMGB1 protein levels decreased at 4 h and are significantly suppressed at 8 h, before gradually returning to a normal level at 12 h after heat shock stress.
Heat shock pretreatment inhibited release and translocation of HMGB1 induced by LPS
To determine whether HSR could affect the release of macrophage HMGB1, HMGB1 was measured in conditioned medium of macrophage cultures incubated with LPS for 20 h in the presence or absence of heat shock pretreatment. Heat shock pretreatment significantly inhibited HMGB1 release induced by LPS (500 ng/mL; Fig. 6A). However, in RAW 264.7 cells incubated for 20 h with LPS (500 ng/mL), we found that heat shock prevented LPS-mediated translocation of HMGB1 from the nucleus to the cytoplasm (Fig. 6B).
HMGB1 has recently been identified as a mediator of endotoxin lethality (3, 4, 7). We have shown here that the expression of HMGB1 was not increased in cultured RAW 264.7 cells after LPS (500 ng/mL) treatment (Fig. 1). This result was consistent with other previous data. Wang et al. (3) reported that the levels of HMGB1 mRNA were unaffected between 0 and 16 h in vitro after LPS (1000 ng/mL) treatment. We have extended these previous observations and discovered that a decrease of HMGB1 mRNA expression was observed after 18 h after LPS (500 ng/mL) treatment (Fig. 1B). However, in vivo administration of LPS or thermal injury increased the mRNA expression of HMGB1 (30, 31). These data suggest that there may be a difference in the regulation of HMGB1 after administration of LPS in vitro or in vivo.
Under physiological conditions, HMGB1 binds to chromatin in a stable way (4, 6). After LPS stimulation, HMGB1 appears to migrate from the nucleus to the cytoplasm (Fig. 2) (32). In LPS-activated macrophages, less HMGB1 was detected in the nuclear fraction and more HMGB1 in the cytoplasm compared with control (Fig. 3).
To verify that HMGB1 could also release into extracellular space after LPS administration in cultured macrophages, we detected the levels of HMGB1 in the culture medium in normal and LPS-activated RAW 264.7 cells. The release of HMGB1 started to be detected at 8 h and was still increasing at 12 and 24 h after LPS (500 ng/mL) treatment. These data were in accord with other reports (3, 9). Mice showed increased serum levels of HMGB1 from 8 to 32 h after endotoxin exposure (3). Serum levels of HMGB1 were increased in patients with sepsis or hemorrhagic shock (3, 11). Compared with normal controls, serum HMGB1 levels were significantly elevated in critically ill patients with sepsis, and were even higher in nonsurvivors compared with survivors (3).
HMGB1 was also released in a dose-dependent manner in vitro when RAW 264.7 cells were stimulated with LPS. Cell viability, as judged by trypan blue exclusion, was unaffected by LPS at concentrations ranging from 100 to 1000 ng/mL, indicating that HMGB1 release was not due to cell death (date not shown). HMGB1 could be secreted by activated macrophages and can also be leaked out by necrotic cells (3, 7). Gardella et al. (32) reported that HMGB1 was secreted by monocytes via a nonclassical, vesicle-mediated secretory pathway. Recent studies have shown that monocytes have a nuclear reshuttling mechanism and (de)acetylating activities to regulate the cellular localization of HMGB1 (33). IFN-γ induced HMGB1 release partly through a TNF-dependent and a Janus kinase 2-dependent mechanism (9).
HSR, a primitive and highly conserved cellular defense mechanism, has broad protective effects against sepsis-induced injury (22-26). In various models of sepsis, induction of HSR protected against sepsis-induced mortality, organ injury, cardiovascular dysfunction, and apoptosis. The mechanisms by which HSR protects against sepsis-induced injury are currently under investigation. One potential mechanism involving the effect of HSR is to inhibit proinflammatory responses. HSR has been demonstrated to inhibit expression of the early cytokines TNF-α and IL-1β (24). Here, we reported that HSR inhibited the late proinflammatory cytokine HMGB1 release. Previous studies revealed that during HSR, different heat shock genes were activated, whereas most other genes became silent (18). Thus, the influence of heat shock per se on the expression of HMGB1 was investigated first. We observed that HMGB1 protein decreased at 4 h, was significantly suppressed at 8 h, and was restored to a normal level at 12 to 24 h after heat shock stress (42.5°C, 1 h). As the major inducible heat shock protein, our data showed that the maximal level Hsp72 was observed at 12 h after heat shock in RAW 264.7 cells. Thus, using this model, we could eliminate the influence of HSR itself on the expression of HMGB1 in the subsequent experiment of HSR pretreatment, which affected release of HMGB1 when cells were recovered for 12 h after heat shock. In subsequent experiments, we found that HMGB1 levels in culture medium in the HSR pretreatment group were significantly less than those in control group after LPS (500 ng/mL) treatment. Cell viability was unaffected after HSR (42.5°C, 1 h) pretreatment (date not shown). To further understand the mechanism by which heat shock inhibited HMGB1 release from cultured macrophages, we detected the levels of LPS-induced nuclear HMGB1 in normal and HSR pretreatment group by Western blot. Our data here show that RAW 264.7 cells that had experienced heat shock pretreatment remarkably prevented LPS-mediated HMGB1 decreases in the nucleus.
To date, the mechanism for the inhibitory effect of heat shock on release of HMGB1 is not clear. Here, we postulated the possible mechanism as follows according to our knowledge. Anti-inflammatory regulation by the stress response was an effective autoprotective mechanism when the host encountered harmful stimuli. Cells and organisms previously submitted to stresses that induced HSR became protected against a second exposure to the same stress, as well as against other types of injury, provided that this happened during the period in which HSPs were significantly expressed (18). We postulate one kind of overexpressed HSP could directly influence the expression and release of HMGB1 through protein-protein interactions. Heat shock-inhibited HMGB1 release may be achieved by interfering with the correlative signaling pathway. Upon activation of inflammatory cells by LPS or the binding of IL-1, TNF, LPS, IFN-γ, or HMGB1 itself to their own receptors, the NF-κB and MAPK (p38, ERK, and JNK) pathways were activated. Together with others, we have discovered that HSR could inhibit cytokine-mediated phosphorylation of JNK and ERK and nuclear translocation of NF-κB (24-26). Thus, the organism after heat shock might regulate inflammatory signaling pathway and make HMGB1 in a stable underacetylated state. Heat shock factor 1 (HSF1) might regulate activation of the HMGB1 promoter. HSF1 is a transcription factor that played a central role in the HSR by regulating expression of HSPs (19). Previously, it has been shown that HSF1 can directly repress activation of the IL-1β and TNF-α promoter (34, 35). Our previous study showed that HSF-1 null mutant mice increased proinflammatory cytokine production in response to endotoxin, and increased endotoxin-mediated lethality compared with wild-type animals (19). Moreover, we found that HMGB1 promoter contains an intact HSF1-binding site (located in the −638 to −648 region. The transcriptional start site was numbered as +1) through predicting transcription factor binding sites in the HMGB1 promoter using a web tool TESS (http://www.cbil.upenn.edu/tess), which indicated that HMGB1 could be regulated by HSF1 (data not shown).
In summary, HMGB1 was released by activated macrophages, induced the release of other proinflammatory mediators, and mediated lethality. The delayed kinetics of HMGB1 release provided a wider therapeutic window for administration of antagonists, and inhibitors to treat human sepsis or SIRS. Here, we demonstrated in vitro that heat shock pretreatment could potently suppress the release and translocation of HMGB1 in response to LPS. Further study of this phenomenon on animal experiments may provide more insight into the molecular mechanisms of HSR anti-HMGB1 effects.
The authors thank Dr. Ivor Benjamin (University of Utah Cardiology Division), Dr. Donald A. Fox and Dr. Qin Chen (University of Houston), and Ms. Zhou Tian for the critical reading of the manuscript. They also thank Meidong Liu for help in preparing reagents.
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