High mobility group box 1 (HMGB1), a chromatin-binding nuclear protein, plays a critical role in sepsis by acting as a key “late-phase” inflammatory mediator. Integrin CD11b is essential for inflammatory cell activation and migration, thus mediating inflammatory responses. However, it is unclear whether CD11b participates in the development of sepsis. In this study, we report that CD11b contributes to LPS-induced endotoxin shock and microbial sepsis, as antagonism of CD11b with the CD11b blocking Ab or CD11b inhibitor Gu-4 protects mice against LPS- and microbial sepsis-related lethality, which is associated with significantly diminished serum HMGB1 levels. Consistent with this, CD11b-deficient mice were more resistant to microbial sepsis with a much lower serum HMGB1 level compared with wild-type mice. Pharmacological blockage and genetic knockdown/knockout of CD11b in murine macrophages hampered LPS-stimulated HMGB1 nucleocytoplasmic translocation and extracellular release. Furthermore, silencing CD11b interrupted the interaction of HMGB1 with either a nuclear export factor chromosome region maintenance 1 or classical protein kinase C and inhibited classical protein kinase C–induced HMGB1 phosphorylation, the potential underlying mechanism(s) responsible for CD11b blockage-induced suppression of HMGB1 nucleocytoplasmic translocation and subsequent extracellular release. Thus, our results highlight that CD11b contributes to the development of sepsis, predominantly by facilitating nucleocytoplasmic translocation and active release of HMGB1.

Sepsis, a systemic inflammatory response mediated by microbial infection or injury, is still one of the most important causes of death in intensive care units (1, 2). In recent years, growing evidence has demonstrated that high mobility group box 1 (HMGB1) plays a pivotal role in the pathogenesis of sepsis and septic shock by acting as a key “late-phase” inflammatory mediator (3, 4). Because of the wider window of manipulation opportunities provided by HMGB1 as compared with early proinflammatory cytokines such as TNF-α, IL-1β, and IL-6, HMGB1 has become a new therapeutic target for inflammation (5, 6).

HMGB1 is a nonhistone nuclear protein and displays both intracellular and extracellular activities (7). It has been well documented that innate immune cells including monocytes/macrophages and polymorphonuclear neutrophils (PMNs) can actively secrete HMGB1 in response to either LPS or TNF-α stimulation; however, the underlying mechanisms remain largely unknown (8). The intracellular location of HMGB1 can be changed upon inflammatory stimuli, and furthermore, translocation of HMGB1 form the nucleus into the cytoplasm is a critical step for subsequent HMGB1 active release (9). Several studies have shown that modifications of HMGB1 through phosphorylation or acetylation are important events for HMGB1 extracellular secretion (10, 11). When HMGB1 is phosphorylated within its nuclear localization sequence, it translocates from the nucleus into the secretory lysosome of the cytoplasm (11). Oh et al. (12) reported that in response to LPS stimulation the classical protein kinase C (cPKC) translocated into the nucleus to phosphorylate HMGB1 directly, causing extracellular secretion of HMGB1 via a calcium-dependent mechanism. Acetylation of HMGB1 may also alter its charge, diminishing nuclear localization and resulting in the eventual secretion of HMGB1 into the extracellular space (10). Chromosome region maintenance 1 (CRM1) has been found to be involved in HMGB1 nucleocytoplasmic translocation following HMGB1 modifications by the upstream signal molecules (13, 14). Once released, HMGB1 acts as a chemokine and/or cytokine to exacerbate the inflammatory response by directly binding to TLR2, TLR4, receptor advanced glycation end products, and syndecan (6, 15). Moreover, HMGB1 functions as an adjuvant by binding to and transferring LPS to CD14 to amplify LPS-mediated inflammation during endotoxin shock and Gram-negative sepsis (1618). Therefore, tight control of HMGB1 release has a potential therapeutic advantage for the treatment of inflammation and sepsis.

Integrin-mediated immune cell migration and cell–cell interactions are essential in immune responses (19, 20). In addition to mediating cell adhesion, β2 integrins play important roles in immune response and inflammation (20). β2 Integrins contain a common CD18 β subunit with different α subunits, for example, CD11a (LFA-1), CD11b (Mac-1, CR3), or CD11c (CR4). Mac-1 (CD11b/CD18, αMβ2) plays an important role in innate immunity by modulating inflammatory cell-associated pathogen recognition, phagocytosis, and cell survival (21, 22). CD11b is extensively expressed in most immune cells, including PMNs, monocytes/macrophages, dendritic cells, and NK cells, which is critical for cell activation, chemotaxis, phagocytosis, and cytotoxicity (23, 24). We demonstrated previously that a lactosyl derivative, N-[2-(1,3-dilactosyl)-propanyl]-2-amino-pentandiamide (Gu-4) functioned as a specific antagonist of CD11b and ameliorated LPS-induced acute lung injury (25, 26). Using a rat model of cecal ligation and puncture (CLP)–induced polymicrobial sepsis, we further found that administration of Gu-4, a specific inhibitor of CD11b, rescued animals from polymicrobial sepsis-associated lethality, and this protection was positively correlated with a significantly reduced serum HMGB1 level (27).

In the present study, we sought to clarify whether CD11b contributes to the development of microbial sepsis by modulating HMGB1 active release. We found that antagonism of CD11b protected mice against endotoxin shock and microbial sepsis, which is associated with significantly diminished serum HMGB1 levels. Consistently, CD11b deficiency rendered mice more resistant to microbial sepsis with substantially blunted circulating HMGB1. Pharmacological blockage and genetic knockdown/knockout of CD11b impaired LPS-stimulated interaction of HMGB1 with CRM1 and cPKC, thus suppressing HMGB1 nucleocytoplasmic translocation and its extracellular release. Our results suggest that CD11b participates in the process of microbial sepsis by promoting HMGB1 release via modulation of CRM1- and cPKC-promoted HMGB1 nucleocytoplasmic translocation.

Pyogen-free, 8- to 10-wk-old male wild-type C57BL/6 and CD11b-deficient (CD11b−/−, B6.129S4-Itgamtm1Myd/J) mice were obtained from the Model Animal Research Center of Nanjing University (Nanjing, China) and The Jackson Laboratory (Bar Harbor, ME), respectively. Mice were housed in barrier cages under controlled environmental conditions (12-h light/12-h dark cycle, 55 ± 5% humidity, 23°C) and had free access to standard laboratory chow and water. Animals were fasted 12 h before experiments and allowed water ad libitum. All animal procedures were conducted with the ethical approval granted from the Institutional Animal Care and Use Committee at Nanjing Normal University and complied with the requirements of provisions and general recommendation of Chinese experimental animal administration legislation. The methods applied in this study were carried out in accordance with the approved guidelines.

Endotoxin shock was induced by i.p. injection of a lethal dose of LPS (35 mg/kg). Polymicrobial sepsis was induced by CLP as described previously (26, 28). Briefly, age- and weight-matched wild-type and CD11b-deficient mice were anesthetized by i.m. injection of 150 μl of a ketamine/xylazine admixture (150 μl ketamine plus 150 μl xylazine made up to 1 ml with 0.9% saline). A midline laparotomy was performed at which the cecum was delivered and ligated at the base just distal to the ileocecal juncture with a 2/0 mersilk tie. A single through puncture was then made distal to the ligature with a 17-gauge needle. The cecum was returned to the peritoneal cavity and the abdomen was closed with 6/0 prolene sutures. The wild-type mice also received an i.p. injection of PBS as the control, CD11b blocking Ab (2.0 mg/kg), and Gu-4 (0.9 mg/kg) 30 min before LPS or CLP challenge. Survival was monitored for at least 7 d.

The TLR4 agonist LPS from Escherichia coli serotype O111:B4 was obtained from Sigma-Aldrich (St. Louis, MO). Abs that recognize CD11b, HMGB1, and CRM1 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), R&D Systems (Minneapolis, MN), and BD Biosciences (San Jose, CA), respectively. Abs that recognize cPKC, phospho-PKC (T497), β-actin, and Lamin B were purchased from Cell Signaling Technology (Beverly, MA). Anti–phospho-Ser/Thr Ab was obtained from Millipore (Billerica, MA). LEAF-purified anti-mouse/human CD11a and CD11b blocking Abs were purchased from BioLegend (San Diego, CA). Texas Red- and FITC-conjugated secondary Abs were purchased from Invitrogen (Carlsbad, CA). Short hairpin RNA (shRNA) specifically targeting CD11b and its scrambled shRNA (scrRNA) were obtained from SuperArray Bioscience (Frederick, MD). The PKC inhibitor Gö6983 was purchased from Selleck Chemicals (Houston, TX). All culture medium and reagents used for cell cultures were purchased from Invitrogen. All other chemicals, unless indicated, were from Sigma-Aldrich.

Peritoneal macrophages were collected from wild-type and CD11b-deficient mice by peritoneal lavage and incubated with DMEM containing 15% heat-inactivated FCS in 24-well plates (Falcon, Lincoln Park, NJ) for 90 min to remove nonadherent cells, as described previously (29, 30). Bone marrow-derived macrophages (BMMs) were isolated from the femurs of wild-type and CD11b-deficient mice and cultured in DMEM containing 20% heat-inactivated FBS, penicillin (100 U/ml), and streptomycin sulfate (100 μg/ml), and supplemented with 10 ng/ml recombinant mouse M-CSF (R&D Systems) for 7 d at 37°C in a humidified 5% CO2 atmosphere, as described previously (30). The purity of both peritoneal macrophages and BMMs was >95%, as confirmed by FACScan analysis of the positive F4/80 Ag staining with an anti-F4/80 Ab (Serotec, Oxford, U.K.).

Isolated peritoneal macrophages plated into 24-well plates (2 × 105 cells per well) were stimulated with LPS (200 ng/ml) for various time periods or pretreated with PBS as the control, CD11a blocking Ab (4 μg/ml), CD11b blocking Ab (4 μg/ml), or Gu-4 (40 nM) for 1 h and further stimulated with LPS (200 ng/ml) for 18 h. Cell-free supernatants were collected and stored at −80°C until analysis. For in vivo murine models of endotoxic shock and CLP-induced polymicrobial sepsis, blood samples were collected at different time points after septic challenges. HMGB1 and TNF-α concentrations in the supernatants or in the serum were assessed by ELISA obtained from Shanghai Westang Bio-Tech (Shanghai, China) and eBioscience (San Diego, CA), respectively.

Cell viability was assessed by MTT assay. Briefly, isolated peritoneal macrophages plated into 96-well plates (Falcon) at 2 × 104 cells per well were pretreated with PBS as the control, CD11a blocking Ab (4 μg/ml), CD11b blocking Ab (4 μg/ml), or Gu-4 (40 nM) for 1 h and further stimulated with LPS (200 ng/ml) for 18 h. Cells were then incubated with MTT at 37°C for 4 h by adding 10 μl of a 5 ng/ml MTT solution into each well. After removal of the cell culture supernatant, 200 μl of DMSO was added into each well to dissolve the crystals. The absorbance of each well was measured using a multimode microplate reader (BioTek, Winooski, VT) at 570 nm wavelength and the OD value was recoded.

Peritoneal macrophages isolated from wild-type mice were transfected with CD11b-specific shRNA using FuGENE HD reagent (Roche, Mannheim, Germany) according to the manufacturer’s protocol (SuperArray Bioscience), and its scrRNA was used as the nonsilencing control. Total protein was extracted 48 h after transfection and the efficiency of CD11b interference was analyzed by Western blotting.

Peritoneal macrophages isolated from wild-type mice and CD11b-deficient mice were incubated with LPS (200 ng/ml) for various time periods and stained with the PE-conjugated anti-CD11b mAb (eBioscience). The PE-conjugated isotype-matched mAb (eBioscience) was used as the control. FACScan analysis was performed from at least 10,000 events for detecting the surface expression of CD11b on peritoneal macrophages in response to LPS stimulation on a Coulter FC500 flow cytometer (Beckman Coulter, Krefeld, Germany) using Expo32 software (Beckman Coulter).

Peritoneal macrophages isolated from wild-type and CD11b-deficient mice plated onto 35-mm glass-bottom dishes (Corning, New York, NY) at 5 × 104 cells per dish were stimulated with LPS (200 ng/ml) in the presence or absence of CD11b blocking Ab (4 μg/ml) and the PKC inhibitor Gö6983 (5 μM) for 8 h. Cells were then washed five times with cold PBS, fixed with 4% paraformaldehyde, and permeabilized with 0.1% Triton X-100 for 10 min. After blocking with 3% BSA for 2 h, cells were incubated with the primary anti-HMGB1 and anti-cPKC Abs overnight at 4°C, and further incubated with the FITC- and Texas Red–conjugated secondary Abs. Cell nuclei were stained with DAPI (Invitrogen). Images and immunostaining were taken and analyzed using an A1 confocal laser scanning microscope (Nikon, Tokyo, Japan).

Isolated BMMs were collected at various time periods after different treatments, washed with ice-cold PBS, and lysed on ice in cell lysis buffer (Cell Signaling Technology), supplemented with 1 mM PMSF and cOmplete protease inhibitor mixture (Roche, Indianapolis, IN). The resultant lysates were centrifuged, and nuclear and cytoplasmic extracts were further concentrated using an NE-PER nuclear and cytoplasmic extraction kit (Pierce, Rockford, IL). Protein concentration was determined using a micro bicinchoninic acid protein assay (Pierce). Equal amounts of protein extracts were separated by SDS-PAGE and transblotted onto polyvinylidene difluoride membranes (Schleicher & Schuell, Dassel, Germany). The membrane was blocked for 1 h at room temperature with PBS containing 0.05% Tween 20 and 5% nonfat milk and probed overnight at 4°C with primary Abs. Blots were then incubated with IRDye 800 fluorophore-conjugated secondary Abs (Rockland Immunochemicals, Gilbertsville, PA) and visualized with the LI-COR Odyssey infrared imaging system (LI-COR Biosciences, Lincoln, NE). For immunoprecipitation, equal amounts of the extracted protein were incubated with the indicated Abs overnight at 4°C on a rotator. Thereafter, 50 μl of a 50% slurry of prewashed protein A/G Plus–agarose beads (Pierce) was added to each sample of immunocomplexes and incubated at 4°C for an additional 2-h period. The samples were spun briefly and washed three times in the lysis buffer. The immunoprecipitates were separated by SDS-PAGE, transblotted onto polyvinylidene difluoride membranes, and probed with appropriate Abs. Blots were further incubated with IRDye 800 fluorophore-conjugated secondary Abs (Rockland Immunochemicals) and visualized with the LI-COR Odyssey infrared imaging system (LI-COR Biosciences).

All data are presented as means ± SD. Statistical analysis was performed using the log-rank test with Kaplan–Meier analysis for survival and one-way ANOVA for all others. Statistical calculations were carried out using SPSS 13.0 version (SPSS, Chicago, IL). Differences were judged statistically significant when the p value was <0.05.

We first examined whether CD11b participates in LPS-induced endotoxin shock. Wild-type mice were pretreated with PBS, CD11b blocking Ab, or Gu-4 for 30 min and further challenged with a lethal dose of LPS (35 mg/kg). Survival was monitored for at least 7 d. Antagonism of CD11b with its specific blocking Ab protected mice from LPS-induced lethality, with a significant decrease in mortality from 100% in PBS-treated mice to 50% in CD11b blocking Ab-treated mice (p = 0.0075) (Fig. 1A). A similar protection was also observed in Gu-4–treated mice, with significantly improved survival advantage (p = 0.0143 versus PBS-treated mice) (Fig. 1A). We further analyzed serum levels of HMGB1 and TNF-α at various time points after LPS challenge and found that pretreatment with either CD11b blocking Ab or Gu-4 substantially attenuated serum HMGB1 levels at 12 and 24 h after LPS challenge (p < 0.05 versus PBS-treated mice) (Fig. 1B), but had no inhibitory effect on serum TNF-α levels (Fig. 1C).

FIGURE 1.

Antagonism of CD11b protects mice against LPS-induced endotoxin shock and CLP-induced polymicrobial sepsis with attenuated serum HMGB1. (A) Kaplan–Meier survival curve shows improved survival in CD11b blocking Ab-treated mice (n = 10) (p = 0.0075) and Gu-4–treated mice (n = 10) (p = 0.0143) compared with PBS-treated mice (n = 10) after LPS challenge. (B and C) Serum levels of HMGB1 (B) and TNF-α (C) after LPS challenge. (D) Kaplan–Meier survival curve shows improved survival in CD11b blocking Ab-treated mice (n = 10) (p = 0.0385) and Gu-4–treated mice (n = 10) (p = 0.0395) compared with PBS-treated mice (n = 10) after CLP challenge. (E and F) Serum levels of HMGB1 (E) and TNF-α (F) after CLP challenge. (G) Kaplan–Meier survival curve shows that CD11b-defient mice (n = 10) were more resistant to CLP-induced sepsis than were their wild-type (WT) littermates (n = 10) (p = 0.0349) with the significantly less serum level of HMGB1 after CLP challenge (H). Data shown in (B), (C), (E), (F), and (H) are mean ± SD of three mice per time point. *p < 0.05, **p < 0.01 versus 0 h. p < 0.05 versus PBS-treated mice or WT mice.

FIGURE 1.

Antagonism of CD11b protects mice against LPS-induced endotoxin shock and CLP-induced polymicrobial sepsis with attenuated serum HMGB1. (A) Kaplan–Meier survival curve shows improved survival in CD11b blocking Ab-treated mice (n = 10) (p = 0.0075) and Gu-4–treated mice (n = 10) (p = 0.0143) compared with PBS-treated mice (n = 10) after LPS challenge. (B and C) Serum levels of HMGB1 (B) and TNF-α (C) after LPS challenge. (D) Kaplan–Meier survival curve shows improved survival in CD11b blocking Ab-treated mice (n = 10) (p = 0.0385) and Gu-4–treated mice (n = 10) (p = 0.0395) compared with PBS-treated mice (n = 10) after CLP challenge. (E and F) Serum levels of HMGB1 (E) and TNF-α (F) after CLP challenge. (G) Kaplan–Meier survival curve shows that CD11b-defient mice (n = 10) were more resistant to CLP-induced sepsis than were their wild-type (WT) littermates (n = 10) (p = 0.0349) with the significantly less serum level of HMGB1 after CLP challenge (H). Data shown in (B), (C), (E), (F), and (H) are mean ± SD of three mice per time point. *p < 0.05, **p < 0.01 versus 0 h. p < 0.05 versus PBS-treated mice or WT mice.

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We next examined whether CD11b also contributes to bacterial sepsis. Wild-type mice pretreated with PBS, CD11b blocking Ab, or Gu-4 were subjected to CLP-induced polymicrobial infection, a more clinically relevant model of sepsis. Similar to the significantly improved survival seen in LPS-induced endotoxin shock, antagonism of CD11b with CD11b blocking Ab or Gu-4 afforded protection against CLP-associated lethality, with substantially enhanced survival rates (p = 0.0385 and p = 0.0395 versus PBS-treated mice) (Fig. 1D). Of note, administration of CD11b blocking Ab or Gu-4 resulted in significant reductions in serum levels of HMGB1 at 12 and 24 h after CLP (p < 0.05 versus PBS-treated mice) (Fig. 1E), but not serum TNF-α (Fig. 1F) and other inflammatory cytokines, including serum IL-6, IL-10, and IL-12p70 (Supplemental Fig. 1). To further verify whether antagonism of CD11b exerts a protective benefit upon septic challenges, we used wild-type and CD11b-deficient mice challenged with CLP. As shown in Fig. 1G, CD11b-deficient mice were more resistant to CLP-induced polymicrobial sepsis with an overall survival at 50% compared with a 10% survival rate in their wild-type littermates (p = 0.0349), with much significantly lower serum HMGB1 levels at 12, 24, and 48 h after CLP (p < 0.05 versus wild-type mice) (Fig. 1H). These results indicate that antagonism of CD11b protects mice against LPS- and CLP-associated septic death, possibly via the inhibition of HMGB1 release.

We next determined whether CD11b plays a role in HMGB1 release from LPS-stimulated macrophages by pharmacological blockage and genetic knockdown or knockout of CD11b. Stimulation with LPS caused a time-dependent HMGB1 release from murine macrophages (Fig. 2A). Consistent with its serum levels seen in LPS- and CLP-challenged mice, HMGB1 was released in a remarkably delayed manner with no significant changes at 0–4 h after LPS stimulation; however, significant increases in HMGB1 release were observed, starting from 8 h after LPS stimulation (p < 0.05 and p < 0.01 versus HMGB1 at 0 h) (Fig. 2A), suggesting that LPS-stimulated HMGB1 release is a late-phase event. In contrast, expression of CD11b on murine macrophages reached the peak level at 1 h after LPS stimulation (p < 0.01 versus CD11b at 0 h), and then gradually returned to its constitutive level (Fig. 2B). Pretreatment of macrophages with CD11b blocking Ab or Gu-4, but not CD11a blocking Ab, significantly attenuated LPS-stimulated HMGB1 release (p < 0.05 versus PBS-treated macrophages) (Fig. 2C). To ascertain whether the released HMGB1 was possibly derived from necrotic or damaged cells via a passive way, cell viability assessed among different treatments excluded the possibility of these two types of HMGB1 passive release (Fig. 2D). These results indicate that pharmacological blockage of CD11b efficiently suppresses HMGB1 active release from LPS-stimulated macrophages.

FIGURE 2.

Pharmacological blockage of CD11b attenuates LPS-stimulated HMGB1 release. Murine peritoneal macrophages were stimulated with 200 ng/ml LPS for various time periods. LPS-stimulated HMGB1 release in the culture supernatants (A) and CD11b expression on macrophages (B) were assessed by ELISA and FACScan analysis, respectively. HMGB1 release from macrophages (C) and cell viability (D) at 18 h after LPS stimulation in the presence or absence of CD11a blocking Ab (4 μg/ml), CD11b blocking Ab (4 μg/ml), and Gu-4 (40 nM) were assessed by ELISA and MTT assay. Data shown are mean ± SD of triplicate samples and are representative of at least three independent experiments. *p < 0.05, **p < 0.01 versus 0 h or cells incubated with culture medium (CM). p < 0.05 versus PBS-treated, LPS-stimulated cells.

FIGURE 2.

Pharmacological blockage of CD11b attenuates LPS-stimulated HMGB1 release. Murine peritoneal macrophages were stimulated with 200 ng/ml LPS for various time periods. LPS-stimulated HMGB1 release in the culture supernatants (A) and CD11b expression on macrophages (B) were assessed by ELISA and FACScan analysis, respectively. HMGB1 release from macrophages (C) and cell viability (D) at 18 h after LPS stimulation in the presence or absence of CD11a blocking Ab (4 μg/ml), CD11b blocking Ab (4 μg/ml), and Gu-4 (40 nM) were assessed by ELISA and MTT assay. Data shown are mean ± SD of triplicate samples and are representative of at least three independent experiments. *p < 0.05, **p < 0.01 versus 0 h or cells incubated with culture medium (CM). p < 0.05 versus PBS-treated, LPS-stimulated cells.

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To clarify whether CD11b is primarily involved in LPS-stimulated HMGB1 release, we first knocked down CD11b in murine macrophages by transfection of CD11b-specific shRNA. Among the four clones of CD11b-specific shRNA (shCD11b) tested, the shCD11b clone 2 strongly silenced protein expression of CD11b in murine macrophages compared with macrophages transfected with scrRNA (Fig. 3A, 3B) and was chosen to use in the consecutive experiment. Next, we assessed whether knockdown of CD11b potentially affects LPS-stimulated HMGB1 release. As shown in Fig. 3C, HMGB1 release in response to LPS stimulation was markedly attenuated in CD11b-silenced macrophages by transfection with shCD11b (p < 0.05 versus LPS-stimulated macrophages transfected with scrRNA). To further confirm the importance of CD11b in LPS-stimulated HMGB1 release, we used peritoneal macrophages isolated from wild-type and CD11b-deficient mice. FACScan analysis revealed the constitutive expression of CD11b on wild-type macrophages, but no detectable CD11b expression was found on CD11b-deficient macrophages (Fig. 3D). Consistently, CD11b-deficient macrophages displayed significantly reduced HMGB1 release upon LPS stimulation (p < 0.01 versus wild-type macrophages) (Fig. 3E), thus providing the direct evidence that CD11b is required for LPS-stimulated HMGB1 release.

FIGURE 3.

Genetic silencing or depleting CD11b diminishes LPS-stimulated HMGB1 release. (A and B) Murine macrophages were transfected with four clones of shCD11b or scrRNA, and CD11b knockdown was confirmed by Western blot analysis 48 h after transfection. (C) Murine macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 18 h, and HMGB1 concentrations in culture supernatants were assessed by ELISA. (D) Surface expression of CD11b on peritoneal macrophages isolated from wild-type (WT) and CD11b-deficient mice was assessed by FACScan analysis. (E) Peritoneal macrophages isolated from WT and CD11b-deficient mice were incubated with PBS or stimulated with 200 ng/ml LPS for 18 h, and HMGB1 concentrations in culture supernatants were assessed by ELISA. Data shown are mean ± SD of triplicate samples and are representative of at least three independent experiments. **p < 0.01 versus cells incubated with PBS. p < 0.05, ≠≠p < 0.01 versus LPS-stimulated, scrRNA-transfected cells or WT macrophages.

FIGURE 3.

Genetic silencing or depleting CD11b diminishes LPS-stimulated HMGB1 release. (A and B) Murine macrophages were transfected with four clones of shCD11b or scrRNA, and CD11b knockdown was confirmed by Western blot analysis 48 h after transfection. (C) Murine macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 18 h, and HMGB1 concentrations in culture supernatants were assessed by ELISA. (D) Surface expression of CD11b on peritoneal macrophages isolated from wild-type (WT) and CD11b-deficient mice was assessed by FACScan analysis. (E) Peritoneal macrophages isolated from WT and CD11b-deficient mice were incubated with PBS or stimulated with 200 ng/ml LPS for 18 h, and HMGB1 concentrations in culture supernatants were assessed by ELISA. Data shown are mean ± SD of triplicate samples and are representative of at least three independent experiments. **p < 0.01 versus cells incubated with PBS. p < 0.05, ≠≠p < 0.01 versus LPS-stimulated, scrRNA-transfected cells or WT macrophages.

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HMGB1 is normally localized in the nucleus but can be translocated into the cytoplasm in response to inflammatory stimulation, which is the prerequisite for subsequent HMGB1 extracellular secretion (8, 9). Stimulation of murine macrophages with LPS resulted in HMGB1 nucleocytoplasmic translocation in a time-dependent manner as evidenced by the expressing levels of HMGB1 between cytoplasmic and nuclear fractions (Fig. 4A), where the cytoplasmic HMGB1 was accumulated gradually (p < 0.05 and p < 0.01 versus HMGB1 at 0 h) and the nuclear HMGB1 was decreased moderately in response to LPS stimulation (p < 0.05 versus HMGB1 at 0 h) (Fig. 4B). Of note, knockdown of CD11b by transfection of murine macrophages with shCD11b markedly suppressed LPS-induced HMGB1 translocation from the nucleus into the cytoplasm (p < 0.01 versus LPS-stimulated macrophages transfected with scrRNA) (Fig. 4C, 4D). To further confirm the inhibitory effect of blocking CD11b on HMGB1 nucleocytoplasmic translocation, we examined the subcellular localization of HMGB1 in wild-type and CD11b-deficient macrophages by immunofluorescent staining. HMGB1 was predominantly localized in the nucleus of naive wild-type macrophages, but translocated and accumulated noticeably in the cytoplasm upon LPS stimulation (Fig. 4E). In contrast, impaired HMGB1 nucleocytoplasmic translocation was observed in LPS-stimulated CD11b-deficient macrophages and most HMGB1 continued to remain in the nucleus even after LPS stimulation for 8 h (Fig. 4E). These results suggest that antagonism of CD11b-induced suppression of HMGB1 release is partly through interfering with LPS-stimulated nucleocytoplasmic translocation of HMGB1.

FIGURE 4.

Genetic knockdown/knockout of CD11b suppresses LPS-stimulated nucleocytoplasmic translocation of HMGB1. (A and B) Murine macrophages were stimulated with 200 ng/ml LPS for the indicated time periods. HMGB1 expression in cytoplasmic and nuclear fractions was assessed by immunoblotting (A), and the intensity of HMGB1 signal in each band was normalized by β-actin or Lamin B (B). (C and D) Macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 8 h, and HMGB1 expression in cytoplasmic and nuclear fractions was assessed by immunoblotting (C), and the intensity of HMGB1 signal in each band was normalized by β-actin or Lamin B (D). Data shown are mean ± SD of three separate experiments. *p < 0.05, **p < 0.01 versus 0 h or cells incubated with PBS. p < 0.05, ≠≠p < 0.01 versus LPS-stimulated, scrRNA-transfected cells. (E) Peritoneal macrophages isolated from wild-type (WT) and CD11b-deficient mice were stimulated with 200 ng/ml LPS for 8 h. Confocal images were taken after cells were stained with the anti-HMGB1 Ab and FITC-conjugated secondary Ab. Cell nuclei were stained with DAPI. Original magnification ×1000.

FIGURE 4.

Genetic knockdown/knockout of CD11b suppresses LPS-stimulated nucleocytoplasmic translocation of HMGB1. (A and B) Murine macrophages were stimulated with 200 ng/ml LPS for the indicated time periods. HMGB1 expression in cytoplasmic and nuclear fractions was assessed by immunoblotting (A), and the intensity of HMGB1 signal in each band was normalized by β-actin or Lamin B (B). (C and D) Macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 8 h, and HMGB1 expression in cytoplasmic and nuclear fractions was assessed by immunoblotting (C), and the intensity of HMGB1 signal in each band was normalized by β-actin or Lamin B (D). Data shown are mean ± SD of three separate experiments. *p < 0.05, **p < 0.01 versus 0 h or cells incubated with PBS. p < 0.05, ≠≠p < 0.01 versus LPS-stimulated, scrRNA-transfected cells. (E) Peritoneal macrophages isolated from wild-type (WT) and CD11b-deficient mice were stimulated with 200 ng/ml LPS for 8 h. Confocal images were taken after cells were stained with the anti-HMGB1 Ab and FITC-conjugated secondary Ab. Cell nuclei were stained with DAPI. Original magnification ×1000.

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To elucidate the underlying mechanisms responsible for CD11b blockage-induced interruption of HMGB1 nucleocytoplasmic translocation, we evaluated interaction of HMGB1 with CRM1, a nuclear export factor, by coimmunoprecipitation. As shown in Fig. 5A, the amount of CRM1 in the immunoprecipitated complex with an HMGB1-specific Ab was increased in response to LPS stimulation (Fig. 5A), whereas silencing CD11b in murine macrophages by transfection with shCD11b substantially attenuated LPS-induced CRM1–HMGB1 interaction (p < 0.05 versus LPS-stimulated macrophages transfected with scrRNA) (Fig. 5A, 5B). LPS stimulation-induced posttranslational modification of HMGB1 by phosphorylation is important for subsequent HMGB1 cytoplasmic translocation and extracellular release (1416). Therefore, we examined the effect of CD11b knockdown on phosphorylation of HMGB1. In response to LPS stimulation, phosphorylated HMGB1 at the serine residues was markedly increased in the immunocomplex (Fig. 5C); however, knockdown of CD11b strongly inhibited LPS-induced phosphorylation of HMGB1 (p < 0.05 versus LPS-stimulated macrophages transfected with scrRNA) (Fig. 5C, 5D). These results suggest that silencing CD11b impairs HMGB1 cytoplasmic translocation, which is mediated, at least in part, by the interruption of CRM1–HMGB1 interaction and HMGB1 phosphorylation.

FIGURE 5.

Silencing CD11b impairs interaction of HMGB1 with CRM1 and phosphorylation of HMGB1. Naive murine macrophages and macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 6 h. (A and B) Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti-CRM1 Ab (A), and the intensity of immunoprecipitated CRM1 in each band was corrected by total CRM1 in the lysate (B). (C and D) Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti–p-Ser Ab (C), and the intensity of p-HMGB1 in each band was corrected by total HMGB1 in the lysate (D). Data shown are mean ± SD of three separate experiments. *p < 0.05 versus cells incubated with PBS. p < 0.05 versus LPS-stimulated, scrRNA-transfected cells.

FIGURE 5.

Silencing CD11b impairs interaction of HMGB1 with CRM1 and phosphorylation of HMGB1. Naive murine macrophages and macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 6 h. (A and B) Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti-CRM1 Ab (A), and the intensity of immunoprecipitated CRM1 in each band was corrected by total CRM1 in the lysate (B). (C and D) Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti–p-Ser Ab (C), and the intensity of p-HMGB1 in each band was corrected by total HMGB1 in the lysate (D). Data shown are mean ± SD of three separate experiments. *p < 0.05 versus cells incubated with PBS. p < 0.05 versus LPS-stimulated, scrRNA-transfected cells.

Close modal

Translocation of cPKC from the cytoplasm into the nucleus upon LPS stimulation has been shown to directly phosphorylate HMGB1, leading to HMGB1 extracellular secretion (12). We therefore first detected the interaction of cPKC with HMGB1. As shown in Fig. 6A, the amount of cPKC in the immunoprecipitated complex with an HMGB1-specific Ab was substantially increased in LPS-stimulated murine macrophages. Knockdown of CD11b in macrophages with its specific shCD11b significantly reduced LPS-induced cPKC–HMGB1 interaction (p < 0.05 versus LPS-stimulated macrophages transfected with scrRNA) (Fig. 6A, 6B). We next assessed the cellular distribution of cPKC and its nuclear traslocation upon LPS stimulation in the presence and absence of CD11b blocking Ab by double immunofluorescent staining. cPKC was primarily located in the cytoplasm in naive murine macrophages (Fig. 6C). However, macrophages stimulated with LPS demonstrated translocation of cPKC from the cytoplasm into the nucleus, whereas macrophages stimulated with LPS in the presence of CD11b blocking Ab displayed remarkably weak red colors in the nucleus, indicating that blockage of CD11b impairs LPS-induced transportation of the cytoplasmic cPKC into the nucleus (Fig. 6C). Furthermore, LPS stimulation led to a strong activation of PKC in murine macrophages (p < 0.05 versus naive macrophages), whereas silencing CD11b substantially inhibited LPS-induced phosphorylation of PKC in macrophages transfected with shCD11b (p < 0.05 versus macrophages transfected with scrRNA) (Fig. 6D, 6E). These results indicate that CD11b is involved in LPS-stimulated PKC activation and cPKC–HMGB1 interaction.

FIGURE 6.

Antagonism of CD11b inhibits LPS-stimulated cPKC activation and cPKC–HMGB1 interaction. Naive murine macrophages and macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 6 h (A and B) or 30 min (D and E). (A and B) Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti-cPKC Ab (A), and the intensity of immunoprecipitated cPKC in each band was corrected by total cPKC in the lysate (B). (C and H) Macrophages were incubated with PBS (naive) or stimulated with 200 ng/ml LPS for 8 h in the presence or absence of CD11b blocking Ab (4 μg/ml) (C) and the PKC inhibitor Gö6983 (5 μM) (H). The colocalization of cPKC with HMGB1 (C) and nucleocytoplasmic translocation of HMGB1 (H) were determined after cells were doubly stained with anti-HMGB1 and anti-cPKC Abs, followed by FITC-and Texas Red–conjugated secondary Abs (C), or stained with the anti-HMGB1 Ab, followed by FITC-conjugated secondary Ab (H). Cell nuclei were stained with DAPI. Original magnification ×1000. (D and E) Cell lysates were subjected to immunoblotting with anti-CD11b, anti-cPKC, and anti–p-PKC Abs (D), and the intensity of CD11b, cPKC, and p-PKC signals in each band was normalized by β-actin (E). (F and G) Murine macrophages pretreated with Gö6983 (5 μM) for 60 min were incubated with PBS or stimulated with 200 ng/ml LPS for 6 h. Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti-cPKC Ab (F), and the intensity of immunoprecipitated cPKC in each band was corrected by total cPKC in the lysate (B). Data shown are mean ± SD of three separate experiments. *p < 0.05, **p < 0.01 versus cells incubated with PBS. p < 0.05 versus LPS-stimulated, scrRNA-transfected cells or LPS-stimulated cells.

FIGURE 6.

Antagonism of CD11b inhibits LPS-stimulated cPKC activation and cPKC–HMGB1 interaction. Naive murine macrophages and macrophages transfected with shCD11b or scrRNA were incubated with PBS or stimulated with 200 ng/ml LPS for 6 h (A and B) or 30 min (D and E). (A and B) Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti-cPKC Ab (A), and the intensity of immunoprecipitated cPKC in each band was corrected by total cPKC in the lysate (B). (C and H) Macrophages were incubated with PBS (naive) or stimulated with 200 ng/ml LPS for 8 h in the presence or absence of CD11b blocking Ab (4 μg/ml) (C) and the PKC inhibitor Gö6983 (5 μM) (H). The colocalization of cPKC with HMGB1 (C) and nucleocytoplasmic translocation of HMGB1 (H) were determined after cells were doubly stained with anti-HMGB1 and anti-cPKC Abs, followed by FITC-and Texas Red–conjugated secondary Abs (C), or stained with the anti-HMGB1 Ab, followed by FITC-conjugated secondary Ab (H). Cell nuclei were stained with DAPI. Original magnification ×1000. (D and E) Cell lysates were subjected to immunoblotting with anti-CD11b, anti-cPKC, and anti–p-PKC Abs (D), and the intensity of CD11b, cPKC, and p-PKC signals in each band was normalized by β-actin (E). (F and G) Murine macrophages pretreated with Gö6983 (5 μM) for 60 min were incubated with PBS or stimulated with 200 ng/ml LPS for 6 h. Cell lysates were subjected to immunoprecipitation with anti-HMGB1 Ab and immunoblotting with anti-cPKC Ab (F), and the intensity of immunoprecipitated cPKC in each band was corrected by total cPKC in the lysate (B). Data shown are mean ± SD of three separate experiments. *p < 0.05, **p < 0.01 versus cells incubated with PBS. p < 0.05 versus LPS-stimulated, scrRNA-transfected cells or LPS-stimulated cells.

Close modal

To further examine the impact of LPS-stimulated interaction of cPKC with HMGB1 on HMGB1 nucleocytoplasmic translocation, we used a PKC inhibitor Gö6983 to block PKC activation. As shown in Fig. 6F and 6G, LPS stimulation resulted in significantly increased cPKC–HMGB1 immunocomplex formation (p < 0.05 versus cells incubated with PBS), whereas the interaction of cPKC with HMGB1 stimulated by LPS was substantially suppressed by the PKC inhibitor Gö6983 (p < 0.05 versus LPS-stimulated cells). Moreover, inhibition of PKC with Gö6983 strongly inhibited LPS stimulation-induced nucleocytoplasmic translocation of HMGB1 (Fig. 6H), indicating that PKC activation and its interaction with HMGB1 is critical for HMGB1 nucleocytoplasmic translocation and subsequent extracellular release.

CD11b is a member of the β2 integrin family. Early studies mainly focused on expression and the function of PMN CD11b and found a correlation of the increased expression of CD11b on PMNs with the progression of sepsis, indicating that CD11b may serve as a biomarker with a diagnostic value in sepsis (31). Our recent studies have shown that a lactosyl derivative Gu-4 functions as an inhibitor of CD11b (25), and administration of Gu-4 attenuates LPS-induced acute lung injury (26) and protects animals against CLP-induced polymicrobial sepsis (27). In the present study, we demonstrated that antagonism of CD11b with either the CD11b blocking Ab or the CD11b inhibitor Gu-4 protected mice against LPS-induced endotoxin shock and CLP-induced polymicrobial sepsis. In supporting this finding, we further revealed that mice with deficiency in CD11b were more resistant to microbial sepsis with significantly higher survival compared with their wild-type littermates. These results suggest that CD11b contributes to the development of microbial sepsis.

In response to bacterial infection, innate inflammatory cells, including monocytes/macrophages and PMNs, produce a large amount of proinflammatory cytokines TNF-α, IL-1β, IL-6, and IL-12. Because of their fast production upon stimulation, these proinflammatory cytokines and mediators are referred to as acute-phase proteins and play important roles in the initiation and progression of microbial sepsis (32, 33). In recent years, a ubiquitous nuclear protein HMGB1 was identified as a novel and late-phase proinflammatory mediator, which can be released by activated innate inflammatory cells in response to stimulation with pathogen-associated molecular patterns and inflammatory cytokines (15, 34). The extensive intracellular and extracellular activity of HMGB1 renders itself as a key pathogenic contributor in microbial sepsis, and, furthermore, blockage of HMGB1 release has been show to protect against endotoxin- and microbial sepsis-related lethality (36). Consistent with previous findings, we found that the protection afforded by antagonism of CD11b was closely associated with significantly reduced serum levels of HMGB1, but not TNF-α and other inflammatory cytokines, in mice treated with the CD11b blocking Ab or the CD11b inhibitor Gu-4. Moreover, CD11b-deficient mice impervious to microbial sepsis displayed substantially blunted circulating HMGB1 upon septic challenge. These results indicate that antagonism of CD11b affords protection against endotoxin shock and polymicrobial sepsis, predominantly via attenuating HMGB1 release.

Consistent with our in vivo finding that antagonism of CD11b markedly attenuates circulating HMGB1, pharmacological blockage, or genetic knockdown/knockout of CD11b in vitro by using either the CD11b blocking Ab, CD11b inhibitor Gu-4, CD11b-specific shRNA, or CD11b-deficient macrophages markedly suppressed HMGB1 release from LPS-stimulated murine macrophages, suggesting the involvement of CD11b in the regulation of HMGB1 release. In naive innate inflammatory cells, HMGB1 shuttles between the nucleus and the cytoplasm. Upon activation by pathogen-associated molecular patterns such as LPS or proinflammatory cytokines, HMGB1 relocalizes from the nucleus into the cytoplasm (9), which is a crucial prerequisite for subsequent HMGB1 extracellular release from activated monocytes/macrophages (8, 9). In the present study, we observed that stimulation of murine macrophages with LPS led to a substantial HMGB1 translocation from the nucleus into the cytoplasm, whereas knockdown of CD11b by its specific shRNA almost totally abrogated LPS-stimulated HMGB1 nucleocytoplasmic translocation. Moreover, LPS stimulation failed to induce cytoplasmic translocation of HMGB1 in CD11b-deficient macrophages. These results suggest that CD11b facilitates the translocation of nuclear HMGB1 into the cytoplasm in activated macrophages, and, thus, antagonism of CD11b-attenuated HMGB1 release is, at least in part, via the interruption of nucleocytoplasmic translocation of HMGB1.

Upon LPS stimulation, HMGB1 is posttranslationally modified by phosphorylation or acetylation at specific lysines within the nuclear localization sequence, which leads to the occurrence of HMGB1 translocation from the nucleus into the secretory lysosome for its active release (10, 11). A number of signal and molecular events have been implicated in the modulation of HMGB1 nucleocytoplasmic translocation and subsequent extracellular secretion upon stimulation. For example, CKD712, a derivative of higenamine, inhibits LPS-stimulated HMGB1 secretion by suppressing PI3K–PKC signaling-induced HMGB1 phosphorylation (35). Tanshinone IIA sodium sulfonate attenuates LPS-stimulated HMGB1 release by facilitating HMGB1 endocytic uptake (36). EGCG, a major component of green tea, blocks LPS-induced extracellular release of HMGB1 by promoting LC3-II production and autophagosome formation (37). More importantly, a nuclear export factor CRM1 appears to be critical in assisting HMGB1 active export from the nucleus into the cytoplasm (13, 14, 38). cPKC, in response to LPS stimulation, is capable of translocating from the cytoplasm into the nucleus to phosphorylate HMGB1 and promote HMGB1 extracellular secretion (12). In this study, we demonstrated that silencing CD11b by its specific shRNA disrupted LPS-stimulated immunocomplex formation of HMGB1 with CRM1 and cPKC, and suppressed cPKC-induced HMGB1 phosphorylation. Moreover, inhibition of cPKC strongly impaired LPS-induced interaction of cPKC with HMGB1 and blocked HMGB1 cytoplasmic translocation. These results indicate that CRM1 and cPKC are two potential molecular targets by which CD11b facilitates nucleocytoplasmic translocation and extracellular release of HMGB1.

Taken together, our in vivo data suggest that CD11b contributes to the development of sepsis, as antagonism of CD11b protects mice against endotoxin shock and microbial sepsis, and this protection is associated with diminished circulating HMGB1. Our in vitro data further show that CD11b facilitates LPS-stimulated HMGB1 nucleocytoplasmic translocation and extracellular release by promoting the interaction of HMGB1 with CRM1 and cPKC. Thus, targeting CD11b may have a therapeutic potential for microbial sepsis and other inflammation-related diseases.

This work was supported by National Natural Science Foundation of China Grants 81501703, 81273232, 81501364, 81671967, and 81420108022, Natural Science Foundation of Jiangsu Province Grants BK20150294 and BK20151206, and by Natural Science Foundation for Colleges and Universities in Jiangsu Province Grant 15KJB310018.

The online version of this article contains supplemental material.

Abbreviations used in this article:

BMM

bone marrow–derived macrophage

CLP

cecal ligation and puncture

cPKC

classical protein kinase C

CRM1

chromosome region maintenance 1

Gu-4

N-[2-(1,3-dilactosyl)-propanyl]-2-amino-pentandiamide

HMGB1

high mobility group box 1

PMN

polymorphonuclear neutrophil

scrRNA

scrambled shRNA

shCD11b

CD11b-specific shRNA

shRNA

short hairpin RNA.

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The authors have no financial conflicts of interest.

Supplementary data