Published online before print April 27, 2005
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* Department of Surgery (Immunology), Brigham and Women’s Hospital and Harvard Medical School, Boston, Massachusetts; and
Department of Surgery, Massachusetts General Hospital, Boston, Massachusetts
1 Correspondence: Brigham and Women’s Hospital, Harvard Medical School, 75 Francis Street, Boston, MA 02115. E-mail: jlederer{at}rics.bwh.harvard.edu
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production by LPS-stimulated burn macrophages requires p38 activation. Although we demonstrated that injury increases macrophage TLR4 mRNA expression and intracellular expression of TLR4-myeloid differentiation protein-2 (MD-2) protein, macrophage cell-surface expression of TLR4-MD-2 was not changed by burn injury. Our results suggest that the injury-induced increase in TLR4 reactivity is mediated, at least in part, by enhanced activation of the p38 signaling pathway.
Key Words: monocytes • macrophages • inflammation • signal transduction
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(TNF-), IL-6] in response to Toll-like receptor (TLR) stimuli [1
2
3
4
]. The innate-immune system and specifically macrophages have been shown to be the major source of these cytokines [3
, 4
]. However, the molecular mechanism(s) that are involved in the enhanced TLR responsiveness by macrophages following injury are not well-defined. TLRs are a family of IL-1 receptor (IL-1R)-related molecules that recognize conserved pathogen-associated molecules such as bacterial lipopolysaccharide (LPS), bacterial lipoprotein (BLP), CpG DNA, or viral double-stranded RNA to name a few. Each receptor recognizes a unique set of ligands such as LPS for TLR4 or BLP for TLR2. These receptors share a common [myeloid differentiation primary-response protein 88 (MyD88)-dependent] signaling pathway [5 , 6 ]; however, TLR3 and TLR4 have also been shown to activate cells via a MyD88-independent signaling mechanism [6 , 7 ].
Upon binding of the ligand to the TLR, MyD88 is recruited to the receptor’s cytoplasmic tail and mediates the phosphorylation of IL-1R-associated kinase (IRAK)-1 by IRAK-4. Phosphorylated IRAK-1 then associates with TNF receptor-associated factor 6 and through several intermediate steps, leads to the activation of the mitogen-activated protein kinase (MAPK) signaling molecules, which are signaling enzymes from three protein families: p38 MAPK, extracellular signal-regulated kinase (ERK), and c-jun N-terminal kinase/stress-activated protein kinase (JNK/SAPK) [8
]. Following activation, MAPKs can initiate the synthesis and assembly of a number of transcription factors including nuclear factor-
B (NF-B), activating transcription factor (ATF)-2, and activator protein-1 (AP-1). The existence of several recently described TLR signaling pathway inhibitors, IRAK-M and Toll-IL-1R-8/single immunoglobulin (Ig) IL-1R-related (Tir8/SIGIRR) molecule, indicates that this important innate-immune signaling pathway is under tight control [9
, 10
].
The aim of this study was to determine if changes in the well-defined MyD88-dependent TLR4 signaling pathway might be responsible for the enhanced TLR4 responsiveness by murine macrophages at 7 days post-injury described previously. Using flow cytometry and specific antibodies, we examined the TLR signaling cascade at several levels including changes in cell-surface and cytoplasmic TLR4 expression, expression of MyD88, and IRAK-M as well as expression and activation of p38, ERK, and SAPK/JNK in F4/80+ splenic macrophages. We show that burn injury primes splenic macrophages for augmented LPS-induced p38 activation, which persists for at least 7 days as opposed to ERK1/ERK2 or SAPK/JNK activation, suggesting that increased p38 signaling may be responsible for the augmented TLR4 reactivity of macrophages following burn injury.
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Reagents
Culture medium for in vitro studies consisted of RMPI 1640 supplemented with 5% heat-inactivated fetal calf serum, 1 mM glutamine, penicillin/streptomycin/fungizone, 10 mM HEPES buffer, 100 µM nonessential amino acids, and 2.5 x 10–5 M 2-mercaptoethanol, all purchased from Gibco-Invitrogen Corp. (Grand Island, NY). LPS Escherichia coli 0111:B4 was obtained from Sigma-Aldrich Chemical Corp. (St. Louis, MO) and BLP, from Bachem (King of Prussia, PA), and unmethylated CpG DNA sequence 1826 was custom-prepared by Invitrogen (Carlsbad, CA). The LPS was repurified following the procedure described by Hirschfeld et al. [11
] to remove contaminating TLR2 reactivity. The LPS used in these studies was kindly confirmed to be a TLR4-restricted agonist by Dr. Evelyn Kurt-Jones of the University of Massachusetts (Amherst) using human embryonic kidney 2 cells transfected with TLR2 or TLR4 [11
, 12
].
Mouse injury model
The thermal injury protocol, approved by NIH and Harvard Medical School Standing Committee on Animal Research, was performed as described previously [13
]. Mice were anesthetized by intraperitoneal (i.p.) injection of ketamine (125 mg/kg) with xylazine (6.8 mg/kg). The dorsal fur was shaved, and the animal was placed in an insulated plastic mold to expose 25% total body surface area. This part of the dorsum was then immersed in 90°C water to induce a well-demarcated, full thickness, anesthetic burn injury. The exposed dorsum of sham animals was immersed in isothermic (24°C) water. All animals were resuscitated with an i.p. injection of 1 ml 0.9% pyrogen-free saline. The mortality from burn injury was less than 5%.
Preparation of spleen cell cultures
Mice were killed by CO2 asphyxiation. Spleens were harvested into serum-free RPMI-1640 medium and digested with Liberase CI enzyme (400 µg/ml final concentration; Roche Applied Science, Indianapolis, IN) for 10 min. Pilot experiments performed in our laboratory have demonstrated that Liberase CI increases the cell yield but does not significantly alter the expression of a number of cell-surface markers (CD4, CD8, CD19, F4/80, CD11c, TLR2, or TLR4). Cell suspensions were then prepared by mincing the tissues on wire mesh. Cells were treated with Tris-ammonium chloride solution for 3 min to lyse red blood cells and washed twice before suspension in culture medium in Costar round-bottom, 96-well plates at a concentration of 5 x 105 cells/well.
Measurement of total intracellular signaling molecules by fluorescein-activated cell sorter (FACS) analysis
At 1 and 7 days after injury, total spleen-cell suspensions were prepared from sham and burn-injured mice. The cells were fixed with 1.5% paraformaldehyde (PAF) for 10 min at 37°C, permeabilized with ice-cold methanol (100%) for an additional 10 min, and washed with phosphate-buffered saline, supplemented with 1% bovine serum albumin and 0.1% sodium azide (PBA). Prior to antibody staining, samples were incubated with FcBlock (PharMingen, San Diego, CA) to prevent nonspecific binding. Fluorescein isothiocyanate (FITC)-labeled anti-F4/80 antibody (PharMingen) was added along with unlabeled, primary rabbit polyclonal antibodies specific for MyD88 (StressGen, San Diego, CA), IRAK-M (Chemicon International, Temecula, CA), p38, ERK, or SAPK/JNK (Cell Signaling, Beverly, MA). An isotype-matched rabbit IgG was used as a nonspecific staining control. Cells were then washed and stained with a goat anti-rabbit phycoerythrin (PE)-labeled secondary antibody. Cells were subsequently washed twice and resuspended in 0.3% PAF. Flow cytometry was performed using the FACSCalibur instrument (BD Biosciences, Mountain View, CA), and the results were analyzed using the accompanying CellQuest Pro software.
Detection of phosphorylated (activated) MAPK signaling molecules by FACS
To distinguish between the activated and nonactivated forms of MAPK signaling molecules, we used antibodies specific for the phosphorylated (active) forms of p38, ERK1/2, and SAPK/JNK (Cell Signaling). Spleen cells were harvested at 1 and 7 days after injury, as described above, and then rested for 2 h at 37°C in culture medium to allow cells to reach steady-state conditions. This was done, as our preliminary studies showed that cell harvesting could cause a transient activation of the MAPK signaling cascade, but following 2 h of culture, the levels of phosphorylated MAPKs would return to baseline.
The cells were then incubated with 200 ng/ml phorbol 12-myristate 13-acetate (PMA) or 0.1, 1, or 10 µg/ml LPS for dose-response experiments and 1 µg/ml LPS or culture medium for stimulation experiments. After 5-, 15-, 30-, or 60-min incubation periods, the cells were rapidly fixed with 1.5% PAF, permeabilized with methanol, and stained and analyzed as described above using antibodies specific for the phosphorylated MAPKs.
To confirm the specificity of the antiphospho-p38 antibody, we performed an additional staining study in the presence of blocking peptides. Fifteen minutes after stimulation with 200 ng/ml PMA, the cells were stained with antibody specific for phospho-p38, which was preincubated with an excess amount (10 µg) of phosphorylated p38 peptide or nonphosphorylated p38 peptide.
FACS analysis of cell-surface and intracellular TLR4-MD-2 expression
Cell-surface and intracytoplasmic TLR4-MD-2 levels were measured using flow cytometry. Spleen cells from sham and burn mice were isolated as described above. After 2 h preincubation in culture medium, spleen cells were incubated with FcBlock for 15 min and stained with FITC-labeled anti-F4/80 antibody to identify the macrophage population. The cells were then divided into two groups: surface and intracellular. The surface group was stained with PE-labeled anti TLR4-MD-2 antibodies (eBiosciences, San Diego, CA) and then fixed with 1.5% PAF. The intracellular group was fixed, first with 1.5% PAF to preserve the F4/80 surface stain, permeabilized by adding 0.5% saponin (weight/volume in PBA), and then stained for intracellular TLR4-MD-2. Both groups of cells were washed twice with PBA, stored in 0.3% PAF, and subsequently analyzed by flow cytometer.
Measurement of cytokine production following MAPK inhibition
Spleen cells prepared from mice 7 days after sham or burn injury were cultured in the presence of a dimethyl sulfoxide control, p38 inhibitor SB203580 (Sigma-Aldrich Chemical Corp.), or ERK (MAPK kinase 1/2) inhibitor UO126 (Cell Signaling) at concentrations of 0.1, 0.5, 1, 5, 10, and 20 µM/ml for 1.5 h. LPS (1 µg/ml) was then added as a stimulant as well as Brefeldin A (10 µg/ml). Six hours later, the cells were fixed, surface-stained for F4/80, permeabilized with saponin, and stained for intracellular TNF- expression using PE-labeled, anti-TNF- antibody (eBiosciences).
Quantitative reverse transcriptase-polymerase chain reaction (RT-PCR)
FITC-labeled F4/80+ macrophages were purified (>90% purity) from spleen cell preparations at 7 days after injury using anti-FITC magnetic beads (Miltenyi Biotec, Auburn, CA). RNA was isolated using the TRIzol Reagent (Gibco-BRL, Grand Island, NY), per the manufacturer’s instructions. cDNA was synthesized using the SuperScript III RT system (Invitrogen). RT-PCR was then performed using glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as a housekeeping gene control, TLR4 gene-specific primers sets, and the 2x ABsolute QPCR SYBR Green reagent (ABgene Inc., Rochester, NY; TLR4 primer forward CCTGGCTGGTTTACACGTC; reverse GACATTGCAGAAACATTCGC). The RT-PCR reactions were run on the GeneAmp 5700 sequence detection system, and the results were calculated using the accompanying software program (PE Biosystems, Foster City, CA).
Statistical analysis
Results in figures are presented as mean ± SEM. Statistical analysis was performed using the GraphPad Prism 4 software (GraphPad Software, San Diego, CA). Unpaired Student’s t-tests were used to generate two-tailed P values.
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Figure 1. Expression levels of signaling molecules in F4/80+ macrophages at 1 and 7 days after sham or burn injury measured by flow cytometry. Total spleen cell suspensions were prepared as described in Materials and Methods, permeabilized with methanol, and stained with antibodies for surface F4/80+ and intracellular signaling molecules. At day 1 (A) and day 7 (B), the levels of MyD88 and IRAK-M were comparable in sham and burn mice. However, although similar on day 1, the total levels (phosphorylated and nonphosphorylated) of p38, ERK, SAPK/JNK were decreased by day 7 in burn mice compared with sham. Results are presented as mean value ± SEM of two experiments (n=8 mice; *, P<0.05).
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We first validated this method and determined the optimal LPS concentration needed to induce MAPK signaling by examining levels of phospho-p38 in LPS-stimulated, splenic F4/80+ macrophages from control C57BL/6J mice. As shown in Figure 2A
, LPS caused a rapid (within 15 min) and easily detectable increase in the levels of phospho-p38. These preliminary studies also demonstrated that the magnitude of the response appeared to plateau at 
1 µg/ml. To assure efficient staining of the MAPK molecule examined, we used the strong stimulant PMA as a positive control in each experiment. We include a representative FACS profile of LPS-stimulated macrophages in Figure 2B
to illustrate the levels of staining attained with this approach. To validate the specificity of the antiphospho-p38 antibody, we stained cells with antibody preincubated with phosphorylated or nonphosphorylated p38 peptide (Fig. 2C)
. Phosphorylated p38 peptide completely blocked the antiphospho-p38 antibody-mediated staining of F4/80+ cells, bringing the staining levels down to isotype control. Conversely, the addition of nonphosphorylated p38 peptide to the staining reaction had no effect on the staining. Taken together, these control studies confirm the specificity of this antibody for the detection of phosphorylated p38 MAPK.
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Figure 2. Flow cytometric approach to detect p38 activation in LPS-stimulated, F4/80+ splenic macrophages. (A) After 15 min, LPS caused a rapid and easily detectable increase in the levels of phosphorylated p38. The magnitude of this response appeared to plateau at approximately the 1-µg/ml LPS concentration. PMA was used as a positive control. Results are presented as mean value ± SEM of two experiments (n=8 mice). (B) Representative FACS plots that illustrate the level of staining attainable using this approach. (C) Representative FACS histogram plots demonstrating specificity of the antiphospho-p38 antibody. Cells were stained with standard phospho-p38 antibody or antibody preincubated with phosphorylated or nonphosphorylated p38 peptide. Staining was blocked with the phosphorylated p38 peptide but not with the nonphosphorylated peptide, thus confirming the specificity of the antibody for the phosphorylated form of p38. MFI, Mean fluorescence intensity.
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Figure 3. MAPK activation kinetics. Spleen-cell cultures were stimulated with 1 µg/ml LPS or culture medium for 5, 15, 30, and 60 min. Using flow cytometry, we measured the activation of p38, ERK, and SAPK/JNK in F4/80+ cells with phospho-specific antibodies. LPS caused a rapid increase in the number of cells with detectable phospho-MAPKs. The maximum difference between stimulated and unstimulated cells occurred at the 15- and 30-min time-points. Results are presented as mean value ± SEM of four experiments (n=16 mice; *, P<0.05).
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Figure 4. The influence of injury on LPS activation of MAPK. Total spleen cell suspensions were prepared and rested in a 2-h preculture. The cells were then stimulated with 1 µg/ml LPS or culture medium for 15 and 30 min. Levels of phosphorylated p38, ERK, and SAPK/JNK were measured in F4/80+ cells after permeabilization with methanol. (A) The p38 response. Although similar in unstimulated cells, upon stimulation with LPS, cells from burn mice showed a twofold increase in p38 activation at days 1 and 7 compared with sham animals. (B and C) The ERK and SAPK/JNK responses, respectively. ERK and SAPK/JNK show increased activation in burn mice in response to LPS on day 1 but not by day 7. (D) Activation of MAPK after 15 min of LPS stimulation presented as MFI. Cells from burn mice show significantly increased activation of p38 compared with sham animals at days 1 and 7. Results are presented as mean value ± SEM of three experiments (n=12 mice; *, P<0.05; **, P<0.001).
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by LPS-stimulated, splenic macrophages as a prototypical, proinflammatory cytokine, as previous studies have shown that burn injury significantly increases the LPS-stimulated, intracellular TNF- expression [3
, 17
]. We were therefore interested in the effect of adding specific p38 or ERK inhibitors on LPS-stimulated TNF- production by splenic macrophages from sham and burn mice.
After preculturing spleen cells with graded doses of a specific p38 inhibitor (SB203580) or an ERK inhibitor (UO126), LPS was added for a 6-h activation period in the presence of Brefeldin A, a protein transport inhibitor. Intracellular TNF- was then measured in F4/80+ cells by two-color flow cytometry. As shown in Figure 5
, as expected, LPS induced higher levels of intracellular TNF- expression in cells from burn as compared with sham-injured mice. Adding the p38 inhibitor to these cultures caused a dose-dependent reduction in this enhanced TNF- expression seen in cells from burn mice but at least at the lower doses (0.1–0.5 µM), had no effect on the TNF- expression in sham cells. At the 0.5 µM concentration and higher, there were no longer any significant differences in the TNF- expression between sham and burn-injured mice. In comparison, we found that the pERK inhibitor had virtually no effect on TNF- expression, suggesting that p38 is indeed the dominant MAPK pathway used in these cells for TLR4-induced TNF- production. Taken together, these results support the finding that the p38 signaling pathway plays a prominent role in the augmented TLR4 responsiveness displayed by splenic macrophages at 7 days after injury.
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Figure 5. Effect of inhibition of p38 or ERK signaling on LPS-induced TNF- expression by splenic macrophages. At 7 days after burn or sham injury, total spleen cells were stimulated for 6 h with LPS in the presence of Brefeldin A after a 1.5-h preculture with the indicated doses (µM) of p38 inhibitor (SB203580) or ERK inhibitor (UO126). Intracellular TNF- was measured in F4/80+ cells by flow cytometry. Cells from burn mice showed higher levels of TNF- expression compared with cells from sham mice. The addition of the p38 inhibitor progressively decreased this enhanced production of TNF- in cells from burn mice. At the 0.5-µM and higher concentrations of SB203580, there were no significant differences between the sham and burn groups. In contrast, ERK inhibition had no effect on TNF- levels. Results are presented as mean value ± SEM of two experiments (n=8; *, P<0.05, comparison between sham and burn groups).
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Figure 6. Effects of injury on cell-surface and intracellular expression of TLR4-MD-2. Using flow cytometry and antibodies specific for TLR4-MD-2, we measured the surface and intracellular levels of this receptor in F4/80+ cells at 1 and 7 days after injury. (A) The results presented as percent of cells positive for the TLR receptor; (B) the data as MFI. On day 1, injured animals showed increased levels of intracellular TLR4-MD-2, which persisted at day 7. The surface expression of TLR4-MD-2 was not changed on day 1 or by day 7 in injured versus sham animals. Results are presented as mean value ± SEM of two experiments (n=12 mice; *, P<0.05).
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To determine if injury influenced TLR4 gene expression, we used quantitative RT-PCR to assess changes in expression of TLR4 mRNA in purified F4/80+ macrophages from sham or burn mice at 7 days after injury. Using GAPDH as housekeeping gene control, we detected a 2.5-fold increase in TLR4 mRNA levels in burn mice compared with sham (Fig. 7 ). Taken together, these findings indicate that injury induces a significant increase in TLR4 gene expression levels and intracellular TLR4-MD-2 expression in splenic macrophages.
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Figure 7. Changes in TLR4 gene expression levels following injury. Using quantitative RT-PCR, we measured the TLR4 mRNA in highly purified F4/80+ cells at 7 days after sham or burn injury. Results are presented as 1/ Ct, the difference in the number of PCR cycles needed to reach a threshold between the measured gene (TLR4) and the housekeeping gene GAPDH. This corresponds to approximately a 2.5-fold increase in TLR4 mRNA in burn mice compared with sham (*, P<0.05).
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Other research groups have reported the critical role played by the p38 signaling pathway in the post-injury LPS response. In particular, Schwacha and colleagues [24 , 25 ] have recently documented that burn injury causes increased p38 activation by crude LPS in plastic-adherent, splenic macrophages. Consistent with our findings, they found that the ERK and SAPK/JNK activation pathways played a lesser role in LPS signaling in murine macrophages. Others have shown that burn injury enhanced p38 activation in renal cells and that inhibiting p38 resulted in a reduced neutrophil influx into the glomerulus and prevented the development of burn-induced renal failure [26 ]. Increased p38 activation has also been shown to occur after cardiac ischemia reperfusion injury [27 , 28 ] and in cardiac myocytes exposed to LPS [29 ]. In the latter study, inhibiting p38 activation not only caused decreased LPS-induced cardiac inflammatory cytokine gene expression but also attenuated the LPS-induced decline in left ventricular-developed pressure and coronary blood flow.
Although we have consistently observed similar levels of cell-surface TLR4 expression in burn and sham macrophages [3 , 17 ], recent evidence supports the idea that higher levels of TLR4 reside in the Golgi apparatus within the cytoplasm of monocytes and other cells and that these intracellular receptors have the capacity to rapidly cycle to the cell surface to mediate LPS signaling. Although Wright and colleagues [30 , 31 ] have reported that LPS traffics directly to the Golgi apparatus once internalized, and additional reports have suggested that TLR4 signals intracellularly in coronary artery endothelial [32 ] and intestinal epithelial cells [33 , 34 ], work by Golenbock and colleagues [18 , 19 , 35 ] suggests that LPS signaling in monocytes/macrophages is initiated on the cell surface. Using fluorescent microscopy, they have illustrated that TLR4/CD14/MD-2 complexes recycle rapidly between the Golgi apparatus and the plasma membrane. In these experiments, they demonstrated that the plasma membrane-impermeable, TLR4-specific antibody was sufficient to induce signaling and that the pharmacological disruption of LPS trafficking to the Golgi apparatus did not disrupt the cell membrane-initiated TLR4 response. In light of their findings, we are uncertain what role the injury-associated increase in the intracellular TLR4 pool that we observed plays in the increased LPS responsiveness exhibited by splenic macrophages at 7 days after injury. Although there appears to be significantly more intracellular TLR4-MD-2 receptors available for recycling to the surface of burn versus sham macrophages, we have so far been unable to document any increase in the surface expression of these receptors by stimulated cells from burn animals.
Recently available flow cytometry techniques were used to study macrophage TLR4-MD-2 signaling in the present report, as the number of purified cells obtainable from the spleens of individual animals was insufficient to assay multiple time-points and multiple signaling molecules by Western immunoblotting and immunoprecipitation assays. Moreover, flow cytometry has the added advantage of permitting activation of TLR4 in macrophages within a more physiologic mixed spleen-cell population rather than an isolated fraction of purified macrophages, which would be required for Western analysis. The results of our experiments showed similar levels of MyD88 and IRAK-M molecules in F4/80+ cells from sham and burn-injured mice, suggesting that changes in the expression of these two molecules do not play a significant role in post-injury signaling. However, suitable reagents are not yet available for flow cytometric detection of a number of other molecules in the TLR4 signaling complex and for determination of their activation/phosphorylation. Thus, possible injury-induced alterations in TLR4 signaling proximal to MAPK activation cannot be ruled in or out by the present investigation.
The results presented here suggest the possibility of controlling the marked and potentially harmful up-regulation of TLR4 reactivity by severe injury in a clinical setting by modulating or inhibiting the p38 signaling pathway. However, the potential, beneficial effects of p38 inhibition would have to be weighted against the negative influence of such inhibition on host immunity.
Received December 2, 2004; revised April 1, 2005; accepted April 7, 2005.
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