The cell adhesion molecule CD44, which is the major hyaluronan receptor, has been implicated in the binding, endocytosis, and metabolism of hyaluronan. Previous studies have revealed that CD44 plays crucial roles in a variety of inflammatory diseases. In recent years, TLRs, which are ancient microbial pattern recognition receptors, have been shown to initiate an innate immune response and have been linked to a variety of inflammatory diseases. The present study shows that CD44 negatively regulates in vivo inflammation mediated by TLRs via NF-κB activation, which leads to proinflammatory cytokine production. Furthermore, our results show that CD44 directly associates with TLR2 when stimulated by the TLR2 ligand zymosan and that the cytoplasmic domain of CD44 is crucial for its regulatory effect on TLR signaling. This study indicates that CD44 plays a protective role in TLR-mediated inflammation and is the first to demonstrate a direct association between CD44 and a TLR.

The adhesion molecule CD44 is a broadly distributed type I transmembrane glycoprotein receptor for the glycosaminoglycan hyaluronan (HA).4 This receptor is known to be involved in the binding, endocytosis, and metabolism of HA and also has additional functions in innate and adaptive immunity (1). CD44 is constitutively expressed on both hemopoietic and parenchymal cells. A considerable number of publications have reported that CD44 plays crucial roles in a variety of inflammatory diseases (2, 3, 4, 5, 6, 7, 8), in which CD44 expression is up-regulated on inflammatory cells. Most evidence for the involvement of CD44 in inflammatory diseases has come from animal studies. Animal studies showed that administration of anti-CD44 Abs inhibited inflammation in murine models of collagen- and proteoglycan-induced arthritis (3), cutaneous inflammation (4), experimental autoimmune encephalomyelitis (5), and IL-2-induced vascular leak syndrome (6). Furthermore, CD44-deficient mice survived better than their wild-type counterparts, presumably because the lack of CD44 tips physiological responses toward a favorable outcome. Wang et al. (7) reported that CD44 in ischemic brain tissue seems to be associated with a selective reduction in inflammatory cytokines. These findings suggest an inflammatory role for CD44. In contrast, in a study of bleomycin-induced acute lung injury, CD44-deficient mice showed an enhanced and persistent inflammatory response due to impaired clearance of apoptotic neutrophils and HA fragments from the injury site (8). This study implies an anti-inflammatory role for CD44. Although CD44 plays a role in inflammation, it is becoming increasingly evident that the function of CD44 in inflammation is complex and involves multiple cell types, ligands, and signaling pathways (2). Therefore, it is important to clearly understand the function of CD44 and its mechanisms in inflammatory processes.

The current study aimed to investigate the role of CD44 in inflammation using a zymosan-induced arthritis (ZIA) model of CD44-deficient mice developed by our group (9). ZIA is a popular model of acute arthritis and thought to be mediated by activation of the alternative pathway of complement and the release of lysosomal hydrolases from activated macrophages (10). The recent discovery of pattern recognition receptors and their roles in innate immunity has led to a re-evaluation of our concepts of zymosan-induced inflammation (11). Recently, it became clear in a study on TLR2-deficient mice that TLR signaling pathways have pivotal roles in ZIA (12).

TLRs are a family of type 1 transmembrane proteins that are expressed on diverse cell types. Regulation of TLR signaling is a key step in acute inflammation, septic shock, and innate/adaptive immunity. Both common-to-all and receptor-specific signals have been shown to be elicited from individual receptors belonging to the TLR/IL-1 receptor (TIR) superfamily (13). The ligands for TLR2 include lipopeptides, peptidoglycans (14), and zymosan (11).

The present study also aimed to examine whether there were any differences in the severity of ZIA between CD44-deficient (CD44−/−) and wild-type (CD44+/+) mice, and to determine whether there are molecular interactions between CD44 and TLRs in inflammation.

Zymosan, a wall product of Saccharomyces cerevisiae, LPS from Salmonella enteriditis, and TNF-α were all purchased from Sigma-Aldrich. Flagellin from Salmonella typhimurium and CpG-DNA (ODN2006) were from Invivogen. R848, an imidazoquinoline-derived antiviral agent, and dsRNA poly(I:C) (polyIC) were from Amersham Biosciences/GE Healthcare.

Two types of purified low molecular mass HA (LMW-HA) fragments and one type of high molecular mass HA (HMW-HA) from chicken comb (Seikagaku) were free of protein and other glycosaminoglycans and had peak molecular mass of 3, 22, and 940 kDa. These HA fragments contained an endotoxin content of <0.002 ng/mg as determined by Limulus amebocyte lysate assays.

The CD44−/− mice were described previously (15). These mice and their wild-type counterparts (CD44+/+ mice) (hybrids of 129 and C57BL/6 strains) were kept and bred in the Animal Unit of Chiba University Graduate School of Medicine in environmentally controlled and specific pathogen-free conditions. All data were generated from littermates. CD44−/− mice that had been backcrossed onto the C57BL/6J (B6) background for more than six generations were obtained from The Jackson Laboratory.

All experimental procedures were approved by the Institutional Animal Care and Use Committee. Typing of CD44 genes was performed by PCR of tail DNA using the following primers: CD44 wild-type: 5′-GGCGACTAGATCCCTCCGTT-3′ and 5′-ACCCAGAGGCATACCAGCTG-3′; CD44 knockout: 5′-GTTTCATCCAGCACGCCAT-3′ and 5′-ATTCAGGCTGCGCAACTGT-3′. All experiments were performed using 10–14-wk-old mice.

CD44−/− or CD44+/+ mice (10-wk old) were used as irradiated recipients of BM cells. The mice were placed in a plastic box and X-irradiated with 10 Gy using an MBR-1520R machine (Hitachi) at a dose rate of 2 Gy/min. At 24 h after the irradiation, the mice were given an i.v. injection of other genotype BM cells (2 × 107) through the tail vein. Donor mice (also 10-wk old) were sacrificed by cervical dislocation. BM cells were obtained from the tibia and femur using cold RPMI 1640 (Sigma-Aldrich) supplemented with heat-inactivated FBS (Vitromex), 100 IU/ml penicillin (Invitrogen Life Technologies), and 100 μg/ml streptomycin (Invitrogen Life Technologies). After centrifugation, the BM cells were resuspended in RPMI 1640 and adjusted to 1 × 108 cells/ml. Next, 200 μl of each BM single-cell suspension was injected i.v. into lethally irradiated recipient mice. At 6 wk after the transplantation, the injected mice were used for experiments. Chimeras prepared by injecting CD44+/+ BM cells into irradiated CD44−/− mice are herein referred to as CD44+/+ BM chimeras (44+/+→44−/−). Other chimeras prepared by injecting CD44−/− BM cells into irradiated CD44+/+ mice are referred to as CD44−/− BM chimeras (44−/−→44+/+).

For chimera control groups, CD44+/+ BM cells were injected into irradiated CD44+/+ mice (B6 background) and CD44−/− BM cells were injected into irradiated CD44−/− mice (B6 background), and the resulting chimeras are referred to as CD44+/+ mice (44+/+→44+/+) and CD44−/− mice (44−/−→44−/−), respectively.

Zymosan was prepared and sterilized as a 15 mg/ml suspension in saline. The material was thoroughly agitated before use to ensure that the suspension was homogeneous. Knees were prepared for injection by depilation with a razor to expose the patellar ligament, and the area was cleaned with 70% ethanol. Under diethyl ether anesthesia, the left knee was injected with 10 μl of the zymosan suspension through a 28-gauge needle fitted to a syringe, and the right knee was injected with saline as a control (9). There were 14 tibiofemoral joints in seven mice of each genotype and 16 tibiofemoral joints in eight mice of each chimera type. To eliminate undefined genetic influences and inflammatory effects, there were 46 additional tibiofemoral joints in 46 BM-transplanted mice (44+/+→44+/+: n = 22; 44−/−→44+/+: n = 16; and 44−/−→44−/−: n = 8) on the B6 background. At 1 wk after the injection, the mice were sacrificed with an overdose of diethyl ether. Half of the joints were arthritic and the other half were saline controls to eliminate injection effects. The knees were removed, fixed in 10% neutral-buffered formalin, decalcified in 5% EDTA-2Na solution, and embedded in paraffin. Next, the specimens were cut into 4-μm sections and stained with H&E. Sections of ZIA tissues were microscopically evaluated for the intensities of three parameters of arthritis, namely cellular infiltration, synovial cell hyperplasia, and pannus formation as described by Ohshima et al. (16) with minor modifications. The intensity of each parameter was scored from 0 to 3 as follows: 0, within the normal range; 1, mild changes; 2, moderate changes; and 3, severe changes.

BM-derived macrophages were prepared by 4 days of culture in 10% FBS-containing RPMI 1640 supplemented with 25 ng/ml M-CSF (R&D Systems). BM was collected from the femurs of adult mice and resuspended in complete RPMI 1640 containing 25 ng/ml M-CSF. After 2 days in culture, media and nonadherent cells were aspirated and replaced with fresh media, which was replaced again after 2 days. Assays were performed on day 5 (17).

Primary mouse embryonic fibroblasts (MEFs) from CD44−/− mice or CD44+/+ littermates were isolated from embryos at 12–14 days of gestation. The embryos were minced and then disaggregated with 0.25% trypsin and 20 U/ml DNase I in PBS for 2 h at 37°C. The obtained MEFs were maintained in DMEM containing 10% FBS.

Raw 264.7 cells were maintained in DMEM supplemented with 10% FBS.

The expression vectors for the standard and epithelial forms of human CD44 (pCIneo-CD44s and pCIneo-CD44E, respectively), pCMV-MyD88 (MyD88), pCMV-Mal (Mal), pCMV-IRAK1 (IRAK1), pCMV-TRAF6 (TRAF6), and pFlag-CMV1-TLR2 were described previously (17, 18). Deletion mutants of CD44E and TLR2 were constructed by digestion with appropriate restriction endonucleases, filling in by T4 DNA polymerases, ligation to synthetic oligonucleotide linkers containing in-frame termination codons, and cloning into plasmid vectors. The deletion mutant of CD44E (CD44Ed: deletion of aa 292–391) was created by the introduction of a stop codon at alanine 291 (TGC)→Stop (TGA). The deletion mutant of TLR2 (TLR2cd: deletion of aa 612–784) was created by the introduction of a stop codon at phenyalanine 612 (TTC)→Stop (TGA).

The details of the EMSA used were described previously (19).

BM-derived macrophages were maintained in 10% FBS-containing RPMI 1640. Cells plated in 6-well dishes (1 × 106 cells/well) were stimulated with 10 μg/ml zymosan or 10 ng/ml LPS as samples, 1 ng/ml TNF-α as positive controls, or left untreated as negative controls. At 24 h after the stimulation, the cells were harvested, washed with PBS, and suspended in ice-cold 20 mM HEPES-NaOH (pH 7.9) buffer containing 0.5% Nonidet P-40, 15% glycerol, 300 mM NaCl, 1 mM EDTA, 10 mM NaF, 1 mM DTT, 1 mM sodium orthovanadate, 0.5 mM PMSF, 50 mM calpain inhibitor-1, 1 mg/ml leupeptin, 1 mg/ml pepstatin, and 1 mg/ml aprotinin. After 30 min of gentle agitation at 4°C, the supernatants were collected by centrifugation as whole-cell extracts. DNA-protein binding reactions (15 μl) were performed by incubating the whole-cell extracts (20 μg equivalent of protein) in a 13 mM HEPES-NaOH (pH 7.9) buffer containing 8% glycerol, 50 mM NaCl, 0.4 mM MgCl2, 0.5 mM DTT, 66.6 μg/ml poly(dI-dC), and 33.3 μg/ml salmon sperm DNA for 15 min on ice, followed by an additional 30-min incubation with a 32P-end-labeled synthetic double stranded oligonucleotide probe (0.1 ng, 5 nCi) at room temperature. The final concentration of the probe was 0.25 fmol/μl. Half of the mixture was loaded onto a polyacrylamide gel (5%) in 0.5 × Tris-borate-EDTA buffer to separate the DNA-protein complexes, and the separated complexes were detected by exposing the dried gels to x-ray films. The oligonucleotide sequences for the labeled probe were from Ig κ L chain enhancer B as follows: 5′-GAT CCA GAG GGG ACT TTC CGA GAG-3′ and 5′-GAT CCT CTC GGA AAG TCC CCT CTG-3′.

Wild-type or CD44−/− MEFs were seeded in 12-well dishes (1 × 105 cells/well). Transient transfection was conducted using Lipofectamine 2000 (Invitrogen Life Technologies) and the cells were cotransfected with 0.5 μg of p-55Igκ-luc reporter plasmid for NF-κB and 10 ng of Renilla luciferase internal control plasmid (pRL-CMV; Promega) (20). For CD44 re-expression experiments, CD44+/+ MEFs were cotransfected with 0.5 μg of p-55Igκ-luc reporter plasmid for NF-κB and 10 ng of Renilla luciferase internal control plasmid (pRL-CMV), while CD44−/− MEFs were cotransfected with 0.5 μg of p-55Igκ-luc reporter plasmid for NF-κB, 10 ng of Renilla luciferase internal control plasmid (pRL-CMV), and 0.5 μg of pCIneo (Mock), 0.5 μg of pCIneoCD44s (for full-length CD44s re-expression), or 0.5 μg of pCIneoCD44E (for full-length CD44E re-expression). At 24 h after the transfection, cells were either left unstimulated (as negative controls) or stimulated with 10 μg/ml zymosan, 10 ng/ml LPS as samples, or 1 ng/ml TNF-α as positive controls for 24 h. Next, the firefly and Renilla luciferase activities were determined using a Dual Luciferase assay kit (Promega) and a TD20/20 dual luminometer (Turner Designs). The firefly luciferase activity was normalized by the Renilla luciferase control activity. Values were expressed as the mean relative stimulation and SD for representative experiments (19). Data were analyzed as the fold induction of luciferase activity over unstimulated cells and are representative of the average of three identical experiments.

To investigate the effects of normal or mutant CD44 molecules on TLR signaling, CD44−/− MEFs were seeded in 12-well dishes (1 × 105 cells/well). Transient transfection was conducted using Lipofectamine 2000, and the cells were transfected with p-55Igκ-luc reporter plasmid (0.5 μg), internal control plasmid (0.01 μg), and pCIneoCD44E (0.5 μg), pCIneoCD44E (0.1 μg) plus pCIneo (0.4 μg), pCIneoCD44Ed (deletion of aa 291–361; 0.5 μg), pCLneoCD44Ed (deletion of aa 291–361; 0.4 μg) plus pCIneo (0.1 μg), or pCIneo (Mock; 0.5 μg), the volume of which was 1.001 μg/1 ml. At 24 h after the transfection, the cells were stimulated with 10 μg/ml zymosan or 100 ng/ml LPS for 24 h. The enzyme activity was measured as described above.

Primary cultured BM macrophages from CD44−/− mice or CD44+/+ littermates were plated in 24-well plates at a density of 2 × 105 cells/well in 0.5 ml of medium. Next, the cells were stimulated with zymosan (10 μg/ml), LPS (10 ng/ml), polyIC (10 ng/ml), CpG-DNA (ODN2006; 2 μM), flagellin (10 ng/ml), or R848 (1 μg/ml) for 24 h or left untreated (as negative controls). ELISA kits for mouse IL-6 (R&D Systems or Biosource International) and mouse TNF-α (Biosource International) were used to determine cytokine production.

To assess the effect of HA on the LPS signaling pathway, CD44+/+ and CD44−/− macrophages were stimulated with 20 μg/ml HA of 3, 22, or 940 kDa in molecular mass (18). At 2 h after the HA treatment, the cells were further stimulated with LPS (10 ng/ml) or medium. After another 24 h, the cytokine production was determined as described above. Experiments were repeated at least twice in triplicate.

To investigate the topological region of the CD44 molecule involved in the signaling cascade mediated by TIR-containing receptors, CD44−/− and CD44+/+ MEFs were transfected with p-55Igκ-luc reporter plasmid (0.5 μg), internal control pRL-CMV (0.01 μγ), and pCMV-MyD88 (MyD88), pCMV-Mal (Mal), pCMV-IRAK1 (IRAK1), pCMV-TRAF6 (TRAF6), or their empty counterpart pCMV vector (Mock) (0.5 μg) (17, 21, 22).

The MEFs were maintained in DMEM, containing 20% FBS, and subjected to luciferase assays at 48 h after the transfection.

HEK293 cells were transfected with Lipofectamine 2000. To perform immunoprecipitation experiments, HEK293 cells transfected with various plasmids were washed with serum-free DMEM and stimulated with zymosan in serum-free DMEM. Whole cell lysates were prepared with RIPA buffer (0.1 M Tris-HCl (pH 7.4), 100 mM NaCl, 0.5% Triton X-100, 1% deoxycholate, and 0.1% SDS) containing 0.5% deoxycholic acid, 2 mmol/L sodium vanadate, 50 mmol/L sodium fluoride, 50 mg/ml leupeptin, 25 mg/ml aprotinin, and 10 mg/ml pepstatin (23). Lysates were incubated with an anti-Flag Ab (M2; Sigma-Aldrich) or an anti-HA Ab (12CA5) as a control. Immune complexes were precipitated with protein G-Sepharose (Amersham Biosicences) and analyzed with anti-Flag and anti-CD44 (Hermes3) Abs as described previously (19).

RAW 264.7 cells grown on glass coverslips were washed with serum-free DMEM and incubated with zymosan in serum-free DMEM for 10 min at 37°C. After fixation and permeabilization, non-specific sites and Fc receptors were blocked by incubation in blocking buffer [PBS containing 10% calf serum and 10 μg/ml anti-mouse Fc receptors (2.4G2)]. Double-labeling studies were performed using a primary rabbit polyclonal Ab against CD44 (1: 20; Santa Cruz Biotechnology) and a mouse mAb against TLR2 (1:20; Abcam). For the double-labeling experiments, the cells were incubated with the primary Abs together overnight. After a 40-min incubation with secondary Abs (FITC-conjugated anti-mouse (1:40) and tetramethylrhodamine isomer R [TRITC]-conjugated anti-rabbit (1:40) Abs; both from Dako A/S) in PBS containing 1% BSA, the cells were washed three times in PBS and mounted in 0.1 M Tris-HCl-glycerol containing 5% n-propyl gallate (WAKO). Control experiments were performed in the same manner except that the primary Abs were excluded. Confocal microscopy was performed using an LSM510 META system (Carl Zeiss), equipped with appropriate dual laser excitation and emission filters for maximum separation of the tetramethylrhodamine isomer R and FITC signals. Stained cells were analyzed with the LSM imaging software. Histogram analyses of the fluorescence intensities were performed using the same software.

Differences between groups were analyzed using the StatView J 4.5 software statistical package (23). Scores for histological grading were analyzed by the Mann-Whitney U test. Transcriptional transactivation assays and cytokine production by macrophages were analyzed using Student’s t test.

To obtain insights into the role of CD44 and the relationships between CD44 and TLRs in inflammatory diseases, we initially attempted to develop experimental arthritis in CD44−/− and CD44+/+ mice using zymosan, a classic inducer of acute joint disease.

At 7 days after a zymosan injection, CD44−/− mice developed markedly more severe lesions in their arthritic joints than CD44+/+ mice (Fig. 1, A and B). Histologically, ZIA is characterized by inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in the joint space (24). Mice from each genotype were examined histologically for these three features of arthritis and graded for their severities to examine whether there were any significant differences. The arthritis assessments of the individual mice of each group are summarized in Fig. 1 C. The histological scores of both inflammatory cell infiltration and synovial cell hyperplasia were significantly elevated in CD44−/− mice compared with CD44+/+ mice. Histopathologically, increased infiltration of polymorphonuclear cells, mononuclear cells, and macrophages, and remarkable hyperplasia of the synovial tissue, were seen in the arthritis in CD44−/− mice. Although the pannus formation tended to be more severe in CD44−/− mice than in CD44+/+ mice, statistical analysis did not show any significant differences between the two groups. These results suggest that CD44 may have a protective role against the development of ZIA.

FIGURE 1.

Zymosan-induced arthritis in CD44−/− and CD44+/+ mice. A and B, Sagittal sections of whole knee joints from CD44+/+ (A) and CD44−/− (B) mice at 7 days after induction of arthritis with zymosan. Scale bars = 200 μm. C, Scores for histological grading of inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in CD44+/+ and CD44−/− mice. The mean values are indicated by solid black lines. The statistical significance of differences was evaluated using the Mann-Whitney U test. Each open or closed circle corresponds to the score of an individual mouse.

FIGURE 1.

Zymosan-induced arthritis in CD44−/− and CD44+/+ mice. A and B, Sagittal sections of whole knee joints from CD44+/+ (A) and CD44−/− (B) mice at 7 days after induction of arthritis with zymosan. Scale bars = 200 μm. C, Scores for histological grading of inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in CD44+/+ and CD44−/− mice. The mean values are indicated by solid black lines. The statistical significance of differences was evaluated using the Mann-Whitney U test. Each open or closed circle corresponds to the score of an individual mouse.

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CD44 is present on both hemopoietic cells and parenchymal cells, such as epithelial cells and fibroblasts (2). To examine the cell populations responsible for the severity of the inflammation, we transferred BM cells from CD44−/− mice into irradiated CD44+/+ mice (CD44−/− BM chimeras, 44−/−→44+/+) and CD44+/+ BM cells into irradiated CD44−/− mice (CD44+/+ BM chimeras, 44+/+→44−/−). When arthritis was induced by zymosan in these mice, CD44−/− BM chimeras exhibited more severe changes in inflammatory cell infiltration and synovial cell hyperplasia than CD44+/+ BM chimeras (Fig. 2, A and B). The histological scores of these two parameters were significantly elevated in CD44−/− BM chimeras compared with CD44+/+ BM chimeras (Fig. 2 C). Although the pannus formation tended to be more severe in CD44−/− BM chimeras than in CD44+/+ BM chimeras, statistical analysis did not show any significant differences between the two groups. Furthermore, the grade of arthritis in CD44−/− BM chimeras appeared to be similar to that in CD44−/− mice, and the grade in CD44+/+ BM chimeras appeared to be similar to that in CD44+/+ mice. These results indicate that CD44 on hemopoietic cells could be responsible for the severities of inflammatory cell infiltration and synovial cell hyperplasia in ZIA.

FIGURE 2.

Zymosan-induced arthritis in CD44−/− and CD44+/+ chimeric mice. A and B, Sagittal sections of whole knee joints from a CD44+/+ (A) and CD44−/− (B) BM chimeras at 7 days after induction of arthritis with zymosan. Scale bars = 200 μm. C, Scores for histological grading of inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in CD44+/+ and CD44−/− BM chimeras. The mean values are indicated by solid black lines. The statistical significance of differences was evaluated using the Mann-Whitney U test. Each open or closed circle corresponds to the score of an individual mouse.

FIGURE 2.

Zymosan-induced arthritis in CD44−/− and CD44+/+ chimeric mice. A and B, Sagittal sections of whole knee joints from a CD44+/+ (A) and CD44−/− (B) BM chimeras at 7 days after induction of arthritis with zymosan. Scale bars = 200 μm. C, Scores for histological grading of inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in CD44+/+ and CD44−/− BM chimeras. The mean values are indicated by solid black lines. The statistical significance of differences was evaluated using the Mann-Whitney U test. Each open or closed circle corresponds to the score of an individual mouse.

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To eliminate other undefined genetic influences and inflammatory effects while generating chimeras, we generated CD44+/+ mice with BM from CD44+/+ mice (44+/+→ 44+/+), CD44−/− mice with BM from CD44−/− mice (44−/−→44−/−), and CD44+/+ mice with BM from CD44−/− mice (CD44−/− BM chimeras, 44−/−→44+/+), all of which had been backcrossed onto a B6 background for more than six generations. Arthritis was then induced by zymosan in these mice. CD44−/− mice (44−/−→44−/−) and CD44−/− BM chimeras (44−/−→44+/+) exhibited more severe changes in inflammatory cell infiltration and synovial cell hyperplasia than CD44+/+ mice (44+/+→44+/+) (Fig. 3). Although the pannus formation tended to be more severe in CD44−/− mice (44−/−→44−/−) and CD44−/− BM chimeras (44−/−→44+/+) than in CD44+/+ mice (44+/+→44+/+), statistical analysis did not show any significant differences among these three groups. The results for CD44−/− mice (44−/−→ 44−/−), CD44−/− BM chimeras (44−/−→44+/+), and CD44+/+ mice (44+/+→44+/+) on the B6 background were comparable to those from CD44−/− mice, CD44−/− BM chimeras, and CD44+/+ mice on the hybrid background, respectively.

FIGURE 3.

Zymosan-induced arthritis in CD44+/+ mice (44+/+→ 44+/+), CD44−/− mice (44−/−→44−/−), and CD44−/− BM chimeras (44−/−→44+/+). Scores for histological grading of inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in CD44+/+ mice (44+/+→44+/+), CD44−/− mice (44−/−→44−/−), and CD44−/− BM chimeras (44−/−→44+/+). The mean values are indicated by solid black lines. The statistical significance of differences was evaluated using the Mann-Whitney U test. Each open or closed circle, or open triangle corresponds to the score of an individual mouse.

FIGURE 3.

Zymosan-induced arthritis in CD44+/+ mice (44+/+→ 44+/+), CD44−/− mice (44−/−→44−/−), and CD44−/− BM chimeras (44−/−→44+/+). Scores for histological grading of inflammatory cell infiltration, synovial cell hyperplasia, and pannus formation in CD44+/+ mice (44+/+→44+/+), CD44−/− mice (44−/−→44−/−), and CD44−/− BM chimeras (44−/−→44+/+). The mean values are indicated by solid black lines. The statistical significance of differences was evaluated using the Mann-Whitney U test. Each open or closed circle, or open triangle corresponds to the score of an individual mouse.

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CD44 on both hemopoietic and non-hemopoietic cells has been shown to contribute to the extent of inflammation (2). However, in the case of ZIA, CD44 on hemopoietic cells may have a more important role than CD44 on non-hemopoietic cells, since similar results were observed for CD44−/− mice (44−/−→44−/−) and CD44−/− BM chimeras (44−/−→44+/+) in ZIA. These results also suggest that CD44 on hemopoietic cells may have a role in TLR signaling, since TLR signaling plays a pivotal role in ZIA.

To assess the effects of CD44-deficency on TLR signaling pathways, we examined the activation of NF-κB, one of the principal signal transducers of TLR signaling, in BM macrophages. The activation of NF-κB was monitored using an EMSA at 24 h after TLR ligand stimulation (13). CD44−/− macrophages exhibited elevated NF-κB activity compared with their CD44+/+ counterparts in response to zymosan. Gram-negative bacteria-derived LPS, a well-characterized ligand for TLR4 (25), also induced elevated activation of NF-κB in CD44−/− cells compared with CD44+/+ cells. In contrast to TLR ligands, TNF-α stimulation provoked a similar robust response in each cell type (Fig. 4,A). Similar results were also obtained when primary MEFs from CD44−/− mice were stimulated with either zymosan or LPS (data not shown). These results indicate that the activation of NF-κB was enhanced and prolonged in CD44−/− cells compared with CD44+/+ cells. In time-course studies, the two genotypes of macrophages displayed similar levels of NF-κB activation at 10 min after LPS stimulation, whereas significantly increased levels of NF-κB activation were observed in CD44−/− macrophages compared with CD44+/+ controls at 20 and 120 min after the stimulation (Fig. 4 B). These data indicate that CD44 diminished the NF-κB activation induced by the TLR signaling pathway.

FIGURE 4.

NF-κB activities analyzed by EMSA. A, Increased activities of NF-κB in response to zymosan or LPS in CD44−/− macrophages. Cells were stimulated with 10 ng/ml zymosan, 10 ng/ml LPS, or 1 ng/ml TNF-α for 24 h or left untreated (Control). B, Kinetics of NF-κB activation in response to LPS in macrophages. Cells were stimulated with 10 ng/ml LPS for the indicated periods. Open and closed triangles indicate activated NF-κB dimers and free probes, respectively.

FIGURE 4.

NF-κB activities analyzed by EMSA. A, Increased activities of NF-κB in response to zymosan or LPS in CD44−/− macrophages. Cells were stimulated with 10 ng/ml zymosan, 10 ng/ml LPS, or 1 ng/ml TNF-α for 24 h or left untreated (Control). B, Kinetics of NF-κB activation in response to LPS in macrophages. Cells were stimulated with 10 ng/ml LPS for the indicated periods. Open and closed triangles indicate activated NF-κB dimers and free probes, respectively.

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To evaluate the role of CD44 in the activation of NF-κB stimulation by either zymosan or LPS, we examined the transcriptional transactivation activity of NF-κB by luciferase assays and used MEFs from CD44−/− and CD44+/+ mice. CD44+/+ MEFs did not exhibit NF-κB-dependent transcriptional activation by zymosan, whereas significant NF-κB-dependent activation was detected in CD44−/− MEFs. This CD44-dependent suppression of NF-κB activation was also found in the experiments with LPS (Fig. 5 A). These results are consistent with our notion obtained by the EMSA that CD44 suppresses NF-κB activation.

FIGURE 5.

NF-κB activities analyzed by luciferase assays. A, Increased transcriptional activities of NF-κB in response to zymosan or LPS in CD44−/− MEFs. CD44+/+ (□) and CD44−/− (▪) MEFs were transfected with a reporter plasmid (0.5 μg) and an internal control plasmid (0.01 μg). Cells were left unstimulated (Control) or stimulated with 10 μg/ml zymosan or 10 ng/ml LPS. Vertical axes represent the normalized luciferase activities relative to the mean activity of unstimulated MEFs. Error bars indicate the SD (n = 3). B, Expression of CD44 reverses the hyperactivation of NF-κB in CD44−/− MEFs. CD44+/+ and CD44−/− MEFs were transfected with a reporter plasmid (0.5 μg), internal control plasmid (0.01 μg), and pCIneo (Mock; 0.5 μg) pCIneoCD44s (0.5 μg), or pCIneoCD44E (0.5 μg). Cells were left unstimulated (Control) or stimulated with 10 μg/ml zymosan or 1 ng/ml TNF-α. Vertical axes represent the normalized luciferase activities relative to the mean activity of unstimulated CD44+/+ MEFs. Error bars indicate the SD (n = 3). Data are given as means ± SD. ∗∗, p < 0.01.

FIGURE 5.

NF-κB activities analyzed by luciferase assays. A, Increased transcriptional activities of NF-κB in response to zymosan or LPS in CD44−/− MEFs. CD44+/+ (□) and CD44−/− (▪) MEFs were transfected with a reporter plasmid (0.5 μg) and an internal control plasmid (0.01 μg). Cells were left unstimulated (Control) or stimulated with 10 μg/ml zymosan or 10 ng/ml LPS. Vertical axes represent the normalized luciferase activities relative to the mean activity of unstimulated MEFs. Error bars indicate the SD (n = 3). B, Expression of CD44 reverses the hyperactivation of NF-κB in CD44−/− MEFs. CD44+/+ and CD44−/− MEFs were transfected with a reporter plasmid (0.5 μg), internal control plasmid (0.01 μg), and pCIneo (Mock; 0.5 μg) pCIneoCD44s (0.5 μg), or pCIneoCD44E (0.5 μg). Cells were left unstimulated (Control) or stimulated with 10 μg/ml zymosan or 1 ng/ml TNF-α. Vertical axes represent the normalized luciferase activities relative to the mean activity of unstimulated CD44+/+ MEFs. Error bars indicate the SD (n = 3). Data are given as means ± SD. ∗∗, p < 0.01.

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To further examine the effect of CD44 on NF-κB, CD44 re-expression experiments were performed. Activation of NF-κB by zymosan in CD44−/− MEFs was significantly decreased to the level in wild-type MEFs after transfection of a CD44s expression vector. Similar effects were found after transfection of a CD44E expression vector. These results indicate that both variants of CD44 could suppress NF-κB reporter activation. In contrast, TNF-α-dependent NF-κB activation was not altered in MEFs regardless of the presence or absence of CD44 (Fig. 5 B) to rule out non-specific effects of CD44 transfection. These results indicate that CD44-deficiency is a genetically determined cause of the NF-κB hyperactivation induced by zymosan, one of the TLR ligands.

To investigate whether CD44 affects signaling from TLRs, we prepared primary BM macrophages from CD44−/− mice or CD44+/+ littermates, treated the cells with zymosan, and examined the production of the proinflammatory cytokines IL-6 and TNF-α. CD44−/− BM macrophages stimulated by zymosan produced significantly increased amounts of both IL-6 and TNF-α compared with their CD44+/+ counterparts (Fig. 6, A and B). Furthermore, we assessed the production of both cytokines in response to several other TLR ligands, namely LPS (TLR4 ligand), polyIC (TLR3 ligand), unmethylated CpG-DNA (ODN2006; TLR9 ligand), flagellin (TLR5 ligand),and the imidazoquinoline-derived antiviral agent R848 (TLR7 ligand). The production of both cytokines was significantly increased in CD44−/− BM macrophages compared with CD44+/+ BM macrophages in response to all the TLR ligands tested.

FIGURE 6.

Proinflammatory cytokine production by murine BM macrophages. A and B, Increased production of IL-6 (A) and TNF-α (B) in CD44−/− BM macrophages stimulated by zymosan (10 μg/ml) or left untreated (Control). C and D, Increased production of IL-6 (C) and TNF-α (D) in CD44−/− macrophages stimulated by various TLR ligands or left untreated (Control). The cells were stimulated with LPS (10 ng/ml), polyIC (10 ng/ml), CpG-DNA (ODN; 2 μM), flagellin (10 ng/ml), or R848 (1 μg/ml). The cytokine production was determined by ELISA. Error bars indicate the SD (n = 3). Experiments were repeated at least twice in triplicate, and similar results were obtained. ∗, p < 0.05; ∗∗, p < 0.01.

FIGURE 6.

Proinflammatory cytokine production by murine BM macrophages. A and B, Increased production of IL-6 (A) and TNF-α (B) in CD44−/− BM macrophages stimulated by zymosan (10 μg/ml) or left untreated (Control). C and D, Increased production of IL-6 (C) and TNF-α (D) in CD44−/− macrophages stimulated by various TLR ligands or left untreated (Control). The cells were stimulated with LPS (10 ng/ml), polyIC (10 ng/ml), CpG-DNA (ODN; 2 μM), flagellin (10 ng/ml), or R848 (1 μg/ml). The cytokine production was determined by ELISA. Error bars indicate the SD (n = 3). Experiments were repeated at least twice in triplicate, and similar results were obtained. ∗, p < 0.05; ∗∗, p < 0.01.

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These results indicate that CD44−/− cells are hyperresponsive to both zymosan and the other TLR ligands examined, suggesting that CD44 has an inhibitory role in a common signaling pathway of all the TLRs tested.

HA is widely distributed under homeostatic conditions as an extracellular and cell surface nonsulfated polysaccharide of high molecular mass, usually several million Da.In contrast, accumulation of low molecular mass forms of HA occurs at sites of inflammation (8). HA interacts with cell surface receptors including CD44, thus influencing cell behavior through direct receptor-mediated effects (18, 26). Recently Jiang et al. (1) reported that both TLR2 and TLR4 seem to drive lung inflammation in response to the fragmented form of HA. Currently, both TLR2 and TLR4 are candidates for HA-binding receptors. Therefore, we assessed whether HA stimulation of macrophages affected TLR signaling in a CD44-dependent or -independent manner. In these experiments, we used three different molecular mass ranges of HA, all of which induced activation of focal adhesion kinase through CD44 in our earlier study (18). In our experiments, the sole addition of each different molecular mass HA (3, 22, and 940 kDa) did not significantly affect the cytokine production by BM macrophages. In LPS-treated cells, IL-6 and TNF-α production levels by both CD44−/− and CD44+/+ macrophages were significantly reduced by the addition of LMW-HA (3 and 22 kDa), indicating that the inhibitory effect of LMW-HA for LPS-induced NF-κB activation is not related to CD44 expression. In contrast, TNF-α production by both CD44 −/− and CD44+/+ macrophages and IL-6 production by CD44−/− macrophages were not significantly influenced by the addition of HMW-HA (940 kDa) (Fig. 7, A and B), indicating that HMW-HA may not be involved in the inhibitory effect of CD44 on LPS-induced NF-κΒ activation. As a result, it seems that the inhibitory effect of CD44 on TLR signaling differs in its mechanism from that of LMW-HA.

FIGURE 7.

Proinflammatory cytokine production in response to LPS and HA. A and B, Inhibitory effect of CD44 on TLR signaling is not affected by HA. CD44+/+ and CD44−/− macrophages were stimulated by HA with the indicated molecular mass. After HA treatment, the cells were further stimulated with LPS or medium alone. Production of IL-6 (A) and TNF-α (B) was determined by ELISA. Experiments were repeated at least twice in triplicate with comparable results. ND: Not detected. Error bars indicate the SD (n = 3). ∗, p < 0.05 for LPS-stimulated CD44+/+ or CD44−/− macrophages with HA vs those without HA, respectively.

FIGURE 7.

Proinflammatory cytokine production in response to LPS and HA. A and B, Inhibitory effect of CD44 on TLR signaling is not affected by HA. CD44+/+ and CD44−/− macrophages were stimulated by HA with the indicated molecular mass. After HA treatment, the cells were further stimulated with LPS or medium alone. Production of IL-6 (A) and TNF-α (B) was determined by ELISA. Experiments were repeated at least twice in triplicate with comparable results. ND: Not detected. Error bars indicate the SD (n = 3). ∗, p < 0.05 for LPS-stimulated CD44+/+ or CD44−/− macrophages with HA vs those without HA, respectively.

Close modal

Overexpression of several intracellular components of TLR signaling has been shown to activate NF-κB, probably by mimicking the signaling process (17, 22, 27). These components include TRAF6 (a RING finger protein), IRAK1 (a serine/threonine kinase), and MyD88 and Mal/TIRAP (adaptor molecules) (13, 28, 29). We cotransfected CD44+/+ and CD44−/− MEFs with TRAF6, IRAK1, MyD88 or Mal/TIRAP, and the NF-κB-dependent luciferase reporter. Overexpression of TRAF6, IRAK1, MyD88, or Mal/TIRAP resulted in activation of NF-κB in both CD44+/+ and CD44−/− MEFs to similar degrees (Fig. 8). These results suggest that CD44 can restrain TLR signaling upstream of TRAF6, IRAK1, MyD88, and Mal/TIRAP. These results imply that CD44 may inhibit signaling from TLRs proximally to receptors on the cell membrane.

FIGURE 8.

Effects of CD44 on NF-κB activation after overexpression of intracellular mediators in TLR signaling. CD44 has no significant effects on the NF-κB activation induced by intracellular mediators (MyD88, Mal, IRAK1, and TRAF6) of TLR signaling. CD44−/− and CD44+/+ MEFs were transfected with the p-55Igκ-luc reporter plasmid (0.5 μg), internal control plasmid (0.01 μg) and pCMV-MyD88 (MyD88), pCMV-Mal (Mal), pCMV-IRAK1 (IRAK1), pCMV-TRAF6 (TRAF6), or their empty counterpart pCMV vector (Mock; 0.5 μg). Error bars indicate the SD (n = 3).

FIGURE 8.

Effects of CD44 on NF-κB activation after overexpression of intracellular mediators in TLR signaling. CD44 has no significant effects on the NF-κB activation induced by intracellular mediators (MyD88, Mal, IRAK1, and TRAF6) of TLR signaling. CD44−/− and CD44+/+ MEFs were transfected with the p-55Igκ-luc reporter plasmid (0.5 μg), internal control plasmid (0.01 μg) and pCMV-MyD88 (MyD88), pCMV-Mal (Mal), pCMV-IRAK1 (IRAK1), pCMV-TRAF6 (TRAF6), or their empty counterpart pCMV vector (Mock; 0.5 μg). Error bars indicate the SD (n = 3).

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To obtain insights into the molecular mechanisms for how CD44 inhibits TLR signaling, we first examined the subcellular localizations of CD44 and TLR2 using double-labeling studies involving immunofluorescence and confocal laser microscopy. In agreement with previous examinations (11), TLR2 was enriched on phagosomal membranes containing zymosan particles (Fig. 9,A, particles 1 and 2) in a murine macrophage cell line (RAW 264.7). CD44 was also found in the areas around zymosan particles (Fig. 9,B, particles 1 and 2), which are comparable to phagosomal membranes. Simultaneous detection of TLR2 and CD44 revealed that the two molecules resided in the areas around zymosan particles (Fig. 9,C, particles 1 and 2), indicating that TLR2 and CD44 were colocalized around zymosan particles. Histograms of the intensities of TLR2 and CD44 on a dashed white line (Fig. 9,C) revealed that the intensity pattern of TLR2 showed a similar distribution to that of CD44 (Fig. 9 D), confirming that the two molecules were preferentially colocalized around zymosan particles in endogenous cells.

FIGURE 9.

Double-staining and immunoprecipitation of TLR2 and CD44. A–D, Double-staining confocal microscopy study with anti-TLR2 (A) and anti-CD44 (B) Abs in RAW 264.7 cells. A, TLR2 staining (green) is detected around internal zymosan (particles 1 and 2) in a RAW 264.7 cell. B, CD44 staining (red) is also detected around internal zymosan particles (1 and 2) in a RAW 264.7 cell. C, A merged confocal image of TLR2 (green) and CD44 (red) indicates that TLR2 and CD44 colocalize around the zymosan particles (1 and 2). N: Nucleus of the RAW 264.7 cell. D, Histogram of the intensities of TLR2 (green) and CD44 (red) staining on the white dashed line in C. The x-axis represents the point-to-point distance on the white dashed line in C in mm, and the y-axis shows the fluorescence intensities of TLR2 (green) and CD44 (red) on an arbitrary scale. Similar patterns of intensity distribution are observed for TLR2 and CD44. E, Schematic diagrams of CD44E and CD44Ed (CD44E deletion mutant: deletion of aa 291–361). The deletion mutant lacks most of the cytoplasmic section (71 amino acids) of CD44E, which includes two functional domains (ezrin-radixin-moesin-binding domain and ankyrin-binding domain). F, Schematic diagrams of TLR2 and TLR2cd (cytoplasmic deletion mutant of TLR2). The deletion mutant lacks most of the cytoplasmic section (173 amino acids; deletion of aa 612–784) of TLR2, which includes the TIR domain. G–I, CD44 is coimmunoprecipitated with TLR2. HEK293 cells were transfected in the presence (+) or absence (−) of expression vectors for CD44 (150 kDa), cytoplasmic domain-deleted CD44 (CD44cd; 150 kDa), TLR2 (90 kDa) (G and H), and cytoplasmic domain-deleted TLR2 (TLR2cd; 75 kDa) (I) as indicated. After 10 min of incubation with or without zymosan, cells were lysed and immunoprecipitated with an anti-Flag Ab for TLR2 or TLR2cd, or an anti-HA Ab as a control. TLR2 and CD44 in the precipitate were detected by Western blotting.

FIGURE 9.

Double-staining and immunoprecipitation of TLR2 and CD44. A–D, Double-staining confocal microscopy study with anti-TLR2 (A) and anti-CD44 (B) Abs in RAW 264.7 cells. A, TLR2 staining (green) is detected around internal zymosan (particles 1 and 2) in a RAW 264.7 cell. B, CD44 staining (red) is also detected around internal zymosan particles (1 and 2) in a RAW 264.7 cell. C, A merged confocal image of TLR2 (green) and CD44 (red) indicates that TLR2 and CD44 colocalize around the zymosan particles (1 and 2). N: Nucleus of the RAW 264.7 cell. D, Histogram of the intensities of TLR2 (green) and CD44 (red) staining on the white dashed line in C. The x-axis represents the point-to-point distance on the white dashed line in C in mm, and the y-axis shows the fluorescence intensities of TLR2 (green) and CD44 (red) on an arbitrary scale. Similar patterns of intensity distribution are observed for TLR2 and CD44. E, Schematic diagrams of CD44E and CD44Ed (CD44E deletion mutant: deletion of aa 291–361). The deletion mutant lacks most of the cytoplasmic section (71 amino acids) of CD44E, which includes two functional domains (ezrin-radixin-moesin-binding domain and ankyrin-binding domain). F, Schematic diagrams of TLR2 and TLR2cd (cytoplasmic deletion mutant of TLR2). The deletion mutant lacks most of the cytoplasmic section (173 amino acids; deletion of aa 612–784) of TLR2, which includes the TIR domain. G–I, CD44 is coimmunoprecipitated with TLR2. HEK293 cells were transfected in the presence (+) or absence (−) of expression vectors for CD44 (150 kDa), cytoplasmic domain-deleted CD44 (CD44cd; 150 kDa), TLR2 (90 kDa) (G and H), and cytoplasmic domain-deleted TLR2 (TLR2cd; 75 kDa) (I) as indicated. After 10 min of incubation with or without zymosan, cells were lysed and immunoprecipitated with an anti-Flag Ab for TLR2 or TLR2cd, or an anti-HA Ab as a control. TLR2 and CD44 in the precipitate were detected by Western blotting.

Close modal

Next, we addressed the direct association between CD44 and TLR2 using immunoprecipitation assays. As shown in Fig. 9,G, CD44 and TLR2 were coimmunoprecipitated from an extract of HEK293 cells stimulated by zymosan, but were not coimmunoprecipitated from an extract without stimulation. To assess the roles of the cytoplasmic domains of TLR2 and CD44 in the direct association, we constructed a truncated CD44 mutant (CD44Ed) pCIneoCD44Ed291–361 that lacked the carboxyl-terminal 71 amino acids and almost all the cytoplasmic domain (Fig. 9,E). We also constructed a truncated TLR2 mutant pFlag-CMV1-TLR2cd that lacked the cytoplasmic carboxyl-terminal 173 amino acids including the TIR domain (Fig. 9,F). CD44Ed and full-length TLR2 (Fig. 9, G and H) or full-length CD44 and TLR2cd (Fig. 9 I) were not coimmunoprecipitated in the presence or absence of zymosan stimulation.

These results suggest that zymosan stimulation leads to an association between CD44 and TLR2. Furthermore, it seems that the TLR2 cytoplasmic domain including the TIR region and the CD44 cytoplasmic domain are necessary for the association between these two molecules.

Previous studies have shown that the cytoplasmic domain of CD44 has intercellular signaling motifs and protein domains necessary for interactions with cytoskeletal components (30). To assess the role of the cytoplasmic domain of CD44 in TLR signaling, we examined the transcriptional activation of NF-κB in MEFs transfected with a truncated CD44 mutant pCIneoCD44Ed (deletion of aa 291–361), which behaves as a dominant-negative molecule in the functioning of CD44 (31, 32). Transfection of the CD44Ed (deletion of aa 291–361) mutant into CD44+/+ MEFs enhanced the activation of NF-κB while transfection of full-length CD44E into CD44−/− MEFs reduced both the zymosan and LPS-induced activations of NF-κB (Fig. 10, A and B). These results indicate that the cytoplasmic domain of CD44 has a regulatory role in TIR signaling and leads to NF-κB activation by zymosan or LPS.

FIGURE 10.

Roles of the cytoplasmic domain of CD44. A and B, NF-κB activities analyzed by luciferase assays. Expression of full-length CD44E reduces zymosan-induced (A) and LPS-induced (B) NF-κB activation in CD44−/− MEFs, whereas expression of CD44Ed (deletion of aa 291–361) appears to enhance LPS-induced NF-κB activation in CD44−/− MEFs (B).

FIGURE 10.

Roles of the cytoplasmic domain of CD44. A and B, NF-κB activities analyzed by luciferase assays. Expression of full-length CD44E reduces zymosan-induced (A) and LPS-induced (B) NF-κB activation in CD44−/− MEFs, whereas expression of CD44Ed (deletion of aa 291–361) appears to enhance LPS-induced NF-κB activation in CD44−/− MEFs (B).

Close modal

Eleven TLR paralogues encoded by mammalian genomes sense macromolecules specific to microbes and initiate the production of cytokines that elicit an inflammatory response (13, 28, 29). In their intracellular domains, TLRs share structural homology with the receptor for a proinflammatory cytokine, IL-1. Both common-to-all and receptor-specific signals have been shown to be elicited from individual receptors belonging to the TIR superfamily (13). One of the major pathways activated by TLRs culminates in the activation of the transcription factor NF-κB, which acts as a master switch for inflammation. Throughout this study of ZIA in mice, the absence of CD44 molecules in BM cells aggravated inflammation and enhanced TLR2-mediated NF-κB activation. These results indicated that CD44 has a regulatory role in the inflammatory responses induced by TLR ligands. Recruitment of inflammatory cells is one of the earliest events in the acute inflammatory response and CD44 plays a significant role in the localization of cells in an inflammatory lesion (2). This role of CD44 may affect the kinetics of the response in ZIA models. We histologically confirmed the maximal severity of ZIA at day 7, when the most prominent features of ZIA should be observed (24, 33). The increased severity of ZIA observed for CD44−/− BM cells at the peak period supports the notion that CD44 regulates the magnitude of TLR-mediated inflammation.

CD44 plays a role in inflammation, and CD44 on both hemopoietic and non-hemopoietic cells has been known to contribute to the extent of inflammation (2). However, our findings for ZIA in chimeric mice imply that CD44 expression on hemopoietic cells is more important for the extent of ZIA than CD44 expression on non-hemopoietic cells. ZIA is thought to be mediated by the activation of the alternative pathway of complement from activated macrophages (9, 10). Recently, zymosan has become accepted as one of the ligands for TLRs, and a study of the ZIA model of TLR2-deficient mice revealed that TLR signaling contributes to the extent of ZIA (12). Thus, our findings indicate that CD44 may contribute to the extent of ZIA in TLR signaling.

When we examined the cytokine production in response to TLR ligands for TLR2, TLR3, TLR4, TLR6, TLR8, and TLR9, enhanced cytokine production was always found in CD44−/− BM macrophages compared with CD44+/+ cells. These results suggest that CD44 negatively regulates TLR-mediated cytokine production, possibly through its common-to-all signaling.

TLR signaling is negatively regulated by several molecules, such as SOCS-1 (21, 22), IRAK-M (34), single immunoglobulin IL-1R-related molecule (35), and ST2 (27). Among these, SOCS-1, IRAK-M, and ST2 were barely expressed at steady-state and strongly induced after LPS stimulation in macrophages. These three factors are required to establish LPS tolerance (21, 22, 27, 34, 35), a protection mechanism against LPS-induced tissue damage. Thus, we examined the ability of CD44−/− macrophages to develop LPS tolerance. Contrary to our expectations, CD44−/− cells did not differ from CD44+/+ cells in their LPS tolerance (data not shown). It seems likely that CD44 is a factor that defines the overall magnitude of the LPS response but not LPS tolerance, in contrast to other negative regulators. Although CD44-deficiency enhanced the LPS-mediated response as effectively as zymosan in vitro, the in vivo role of LPS remains unclear. Thus, we generated LPS-sublethal shock models of CD44−/− and CD44+/+ mice. Surprisingly, CD44−/− mice displayed dramatic hypersensitivity to LPS compared with CD44+/+ mice (data not shown), consistent with data in a recent report (36). From this finding, it is clear that CD44 plays an indispensable role in the regulation of TLR-mediated responses in vivo. Further analyses could lead to the development of new treatments for human LPS-induced disease.

CD44 is a well-known HA-binding receptor protein, and now both TLR2 and TLR4 are also candidates for HA-binding receptor proteins. Thus, we assessed the effects of HA on TLR-mediated cytokine production by BM macrophages. We showed that LMW-HA (3 and 22 kDa) had inhibitory effects on TLR signaling, but these effects were independent of CD44 expression. As a result, it seems that the inhibitory effect of CD44 on TLR signaling differs in its mechanism from that of LMW-HA. Furthermore, it is known that synthesis of HA increases under inflammatory circumstances, and that TNF-α is an important stimulator of HA production (37). In our experiments, TNF-α stimulation resulted in similar levels of NF-κB activation in both CD44+/+ and CD44−/− genotypes of BM macrophages and MEFs. These results support the conclusion that the inhibitory effect of CD44 on TLR signaling is independent of HA.

An emerging concept in signal transduction is that cell adhesion molecules can function as coreceptors. CD44 is known to serve as a coreceptor for growth factors and associate with their receptors, such as tyrosine kinase receptor ErbB or c-Met (38, 39). These findings prompted us to examine the subcellular localizations of CD44 and TLRs. As expected, CD44 was found in the area around phagosomes containing zymosan where TLR2 can also exist. We also found that CD44 directly associated with TLR2 by using HEK293 cells after endocytosis of zymosan. In addition, no significant differences were found between CD44−/− and CD44+/+ BM macrophages regarding alterations in their expressions of TLR2 and TLR4, LPS-binding abilities, and endocytic activities for zymosan (data not shown). The study involving overexpression of intracellular signaling molecules for TLRs suggested that CD44 may inhibit signaling from TLR2 and TLR4 proximally to receptors on the cell membrane. Transfection of a CD44 cytoplasmic deletion mutant did not reproduce the inhibitory effect on TLR signaling, and the mutant did not associate with TLR2 after stimulation with zymosan. Therefore, the cytoplasmic domain of CD44 may be critical for the regulation of TLR signaling, possibly by interacting with the TIR domain. Indeed, a truncated TLR2 mutant that lacked the TIR domain could not associate with CD44 under zymosan stimulation. From these results, it is tempting to speculate that CD44 functions as an inhibitory molecule that rapidly moves to phagosomes and physically associates with TLR2. However, it remains possible that the inhibitory effect of CD44 is modulated by endogenous ligands for CD44 other than HA.

This report is the first to demonstrate that CD44 directly associates with TLR2 and negatively regulates TLR-mediated responses. Accordingly, the ultimate function of CD44 may be to protect tissues or organs against damage due to a severe inflammatory response in vivo.

It is clear that CD44 is a potentially important molecule that regulates inflammatory responses. Therefore, we believe that elucidating the molecular mechanisms of CD44 in TLR signaling pathways will help toward achieving an understanding of the mechanisms of inflammatory diseases and thereby provide novel insights into new therapies against inflammatory disorders.

We thank B. Seed (Harvard Medical School, Boston, MA), T. Fujita (The Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan), N. Watanabe, H. Shimada, A. Asari (Seikagaku, Tokyo, Japan), S. Akira, S. Sato (Osaka University, Osaka, Japan), and N. Suzuki and S. Suzuki (RIKEN Research Center for Allergy and Immunology, Kanagawa, Japan) for the reagents used in this study; K. Yoshida (National Institute of Radiological Science, Chiba, Japan) for the BM transplantation; N. Suzuki and S. Suzuki for useful comments and helpful discussions; and T. Saito (RIKEN Research Center for Allergy and Immunology, Kanagawa, Japan) for critical reading of the manuscript.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by a Grant-in-Aid for Scientific Research Priority Area 12215018 from the Ministry of Education, Culture, Sports, Science and Technology, Japan (to K.H.) and a Grant-in-Aid for Scientific Research 13670163 from the Japan Society for the Promotion of Science (to K.H.).

4

Abbreviations used in this paper: HA, hyaluronan; ZIA, zymosan-induced arthritis; TIR, TLR/IL-1 receptor; polyIC, poly(I:C); LMW-HA, low molecular mass HA; HMW-HA, high molecular mass HA; B6, C57BL/6J; BM, bone marrow; MEF, mouse embryonic fibroblasts.

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