We present evidence for a novel TLR2 function in transmodulating the adhesive activities of human monocytes in response to the fimbriae of Porphyromonas gingivalis, a pathogen implicated in chronic periodontitis and atherosclerosis. Monocyte recruitment into the subendothelium is a crucial step in atherosclerosis, and we investigated the role of P. gingivalis fimbriae in stimulating monocyte adhesion to endothelial cells and transendothelial migration. Fimbriae induced CD11b/CD18-dependent adhesion of human monocytes or mouse macrophages to endothelial receptor ICAM-1; these activities were inhibited by TLR2 blockade or deficiency or by pharmacological inhibitors of PI3K. Moreover, this inducible adhesive activity was sensitive to the action of Clostridium difficile toxin B, but was not affected by Clostridium botulinum C3 exoenzyme, pertussis toxin, or cholera toxin. Accordingly, we subsequently showed through the use of dominant negative signaling mutants of small GTPases, that Rac1 mediates the ability of fimbria-stimulated monocytes to bind ICAM-1. A dominant negative mutant of Rac1 also inhibited the lipid kinase activity of PI3K suggesting that Rac1 acts upstream of PI3K in this proadhesive pathway. Furthermore, fimbriae stimulated monocyte adhesion to HUVEC and transmigration across HUVEC monolayers; both activities required TLR2 and Rac1 signaling and were dependent upon ICAM-1 and the high-affinity state of CD11b/CD18. P. gingivalis-stimulated monocytes displayed enhanced transendothelial migration compared with monocytes stimulated with nonfimbriated isogenic mutants. Thus, P. gingivalis fimbriae activate a novel proadhesive pathway in human monocytes, involving TLR2, Rac1, PI3K, and CD11b/CD18, which may constitute a mechanistic basis linking P. gingivalis to inflammatory atherosclerotic processes.

The β2 integrin heterodimer CD11b/CD18 (Mac-1, CR3) is a multifunctional receptor with significant and diverse roles in immunity and inflammation (1, 2). The functional versatility of this integrin is attributed, at least partly, to its ability to recognize multiple and structurally unrelated molecules, including its endothelial counter-receptor ICAM-1, fibrinogen, iC3b, Factor X, and platelet glycoprotein Ibα (3, 4, 5, 6). Abundantly expressed by neutrophils and monocytes, CD11b/CD18 plays a role in their migration to sites of extravascular inflammation (3, 7, 8, 9). The interactions of CD11b/CD18 with fibrinogen and ICAM-1 mediate adhesion of neutrophils or monocytes to sites of fibrinogen deposition and the endothelium, respectively (3, 9). These adhesive interactions are tightly regulated. Although the default conformation of CD11b/CD18 in resting cells is of low affinity, a rapid and transient shift to a high-affinity binding state (referred to as CD11b/CD18 activation) can be triggered by inside-out signaling (1, 10). Inside-out signaling pathways for CD11b/CD18 activation can be induced upon stimulation of other surface receptors, such as chemotactic receptors (1, 10) or TLRs (11, 12).

The potential of CD11b/CD18 for vascular cell interactions by binding to ICAM-1 or to endothelial-associated matrix proteins, such as fibrinogen, may contribute to cardiovascular inflammation (5, 13). In this context, the adhesion of bloodborne leukocytes to the arterial endothelium, followed by their migration into the subendothelial area is a hallmark of early atherogenesis (14). The transmigratory process is mediated by interacting sets of cell adhesion molecules, including the CD11b/CD18-ICAM-1 pair, which has been experimentally implicated in atherosclerosis and other inflammatory conditions (13, 15, 16, 17). It is thought that infectious agents contribute to vascular inflammation and certain bacterial pathogens such as Chlamydia pneumoniae, Helicobacter pylori, and Porphyromonas gingivalis have been implicated as accessory factors in the development or acceleration of atherosclerosis (14, 18, 19). In this regard, infection-driven chronic inflammatory diseases, including periodontitis, are associated with increased risk for cardiovascular disease (20, 21, 22).

P. gingivalis is a Gram-negative oral bacterium that is strongly associated with chronic periodontitis (23). This pathogen may disseminate from periodontal lesions into the systemic circulation and P. gingivalis-specific DNA has been detected in human atherosclerotic plaques (24). Studies in animal models of periodontitis or atherosclerosis have established the P. gingivalis fimbriae (filamentous appendages on the cell surface) as a major virulence factor of this pathogen (25, 26). Although both wild-type P. gingivalis and an isogenic non-fimbriated mutant are detected in the blood and aortic arch tissue of orally infected hyperlipidemic mice, only the presence of wild-type P. gingivalis is associated with periodontal disease and increased atherosclerotic plaque formation (26).

In this study, we have identified a plausible inflammatory mechanism whereby P. gingivalis fimbriae may contribute to the atherosclerotic process. Based on earlier findings that P. gingivalis fimbriae bind CD14 and stimulate TLR2/PI3K-mediated inside-out signaling for CD11b/CD18 activation (11, 27), we now show that activation of this pathway leads to increased monocyte adhesion to fibrinogen, ICAM-1, and endothelial cells. This inducible proadhesive pathway is distinct from other CD11b/CD18 activation pathways stimulated by FMLP or PMA, as shown by differential toxin sensitivity. On the basis of toxin sensitivity data and additional experiments using dominant negative (DN)3 signaling mutants of small GTPases, we found that Rac1 acts upstream of PI3K and is essential for the ability of fimbria-stimulated monocytes to bind CD11b/CD18 ligands, endothelial cells, and transmigrate across endothelial monolayers. The property of P. gingivalis fimbriae to induce CD11b/CD18-dependent adhesive interactions may contribute to the role of monocytes in the process of atherosclerosis or other inflammatory conditions.

mAbs to human TLR2 (clone TL2.1), TLR4 (HTA125), CD11b (CBRM1/5, FITC-labeled), MHC class I (W6/32), and Ig isotype controls (IgG1, IgG2a) were purchased from eBioscience. FITC-labeled mAb to human CD11b (Bear-1) and mAbs to human CD14 (MEM-18), mouse CD11b (M1/70), and its IgG2b isotype control were from Caltag Laboratories. mAb to human CD11b (2LPM19c) was from DakoCytomation. mAb to ICAM-1 (BBIG-I1) and human rICAM-1 plus rIL-1β were from R&D Systems. FMLP, PMA, wortmannin, LY294002, LY30351, GF109203X, and cell culture-grade BSA were purchased from Sigma-Aldrich. Human fibrinogen (depleted of plasminogen, von Willebrand factor, and fibronectin) was obtained from Enzyme Research Laboratories. XVA143, a β2 integrin allosteric antagonist (28), was generously provided by Dr. N. Fotouhi (Roche, Nutley, NJ). P. gingivalis was grown anaerobically at 37°C in brain-heart infusion broth supplemented with hemin (5 μg/ml) and menadione (l μg/ml). P. gingivalis strains used included wild-type strains 381 and A7436 (donated by Dr. H. Kuramitsu, University of Buffalo, Buffalo, NY) and their respective isogenic fimbria-deficient mutants (25, 29). Fimbriae were purified from P. gingivalis strain 381 as previously described (11). Recombinant fimbrillin (rFimA) was purified from Escherichia coli BL21 (DE3) transformed with the fimA gene of P. gingivalis 381 as previously described (30) with an additional step involving chromatography through agarose-immobilized polymyxin B to remove residual endotoxin. The final fimbrial preparations were free of any contaminating substances on silver-stained SDS-PAGE and tested negative for endotoxin (<6 EU/mg of protein) according to a quantitative Limulus amebocyte lysate assay (BioWhittaker).

Monocytes were purified from the peripheral blood of healthy human volunteers as previously described (11). Briefly, monocytes were separated from lymphocytes upon centrifugation of peripheral blood over NycoPrep 1.068 (Axis-Shield). Incidental nonmonocytes were removed by magnetic depletion using a mixture of biotin-conjugated mAbs and magnetic microbeads coupled to anti-biotin mAb (Monocyte isolation kit II; Miltenyi Biotec). Purified monocytes were cultured at 37°C and 5% CO2 atmosphere, in RPMI 1640 (Invitrogen Life Technologies) supplemented with 10% heat-inactivated FBS, 2 mM l-glutamine, 10 mM HEPES, 100 U/ml penicillin G, 100 μg/ml streptomycin, and 0.05 mM 2-ME (complete RPMI). Human blood collections were conducted in compliance with established guidelines approved by the institutional review board. THP-1 cell lines stably transfected with human CD14 (THP-1/CD14) or with empty vector (THP-1/RSV) (31) were cultured in complete RPMI. Both cell lines were provided by Dr. P. S. Tobias (The Scripps Research Institute, La Jolla, CA). HUVEC (PromoCell) were cultured at 37°C and 5% CO2 atmosphere in PromoCell Endothelial Cell Growth medium (2% FCS, 0.1 ng/ml epidermal growth factor, 1.0 ng/ml basic fibroblast growth factor, 1.0 μg/ml hydrocortisone, 0.4% endothelial cell growth supplement/heparin, 50 μg/ml gentamicin, 50 ng/ml amphotericin B), according to the supplier’s recommendations. Thioglycolate-elicited macrophages were isolated from the peritoneal cavity of mice deficient in CD14, TLR2, TLR4, both TLR2 and TLR4, or from wild-type control mice, as previously described (27, 32). The mice deficient in CD14, TLR2, or TLR4 were of C57BL/6 genetic background, whereas mice harboring homozygous TLR2 and TLR4 mutations were 9-fold backcrossed toward the C3H genetic background (kindly donated by Dr. C. Kirschning, Technical University of Munich, Munich, Germany). Mouse macrophages were cultured in complete RPMI as described. The use of animals was reviewed and approved by the Institutional Animal Care and Use Committee. Human or mouse cell viability was monitored using the CellTiter-Blue Cell Viability assay kit (Promega). The use of fimbriae and other agonists as well as treatments with blocking mAbs or other antagonists did not affect cell viability as compared with medium-only control treatments.

Transfections of THP-1/CD14 cells were performed using the FuGene 6 transfection reagent (Roche Applied Science) at a reagent to DNA ratio of 3:1, according to the manufacturer’s instructions. Plasmids expressing DN versions of the human Rac1, Cdc42, and RhoA genes (Rac1/T17N, Cdc42/T17N, and RhoA/T19N, respectively) as well as the empty control vector pUSEamp+ were obtained from Upstate Biotechnology. A plasmid expressing a DN mutant of human TLR2 (pZERO-hTLR2tirless) and empty vector control (pZERO-mcs) were obtained from InVivogen. The cells were used in functional assays 48 h posttransfection. Transient transfection efficiency was 40–45% as determined by GFP reporter plasmid transfection and fluorescence microscopy to detect cells presenting green fluorescence.

The 96-well microtiter plates were coated with 10 μg/ml human fibrinogen or rICAM-1. Following overnight incubation at 4°C, remaining uncoated sites on the plates were blocked with 5 mg/ml BSA for 1 h at room temperature. Monocytes were labeled with the fluorescent dye calcein AM (2.5 μM; Molecular Probes) for 30 min, washed, and resuspended in assay buffer (HBSS, 10 mM HEPES (pH 7.4), 1 mM Mg2+, and 0.42 mM Ca2+). Labeled cells were added to the ligand-coated wells (5 × 104 cells per well) at 37°C in the absence or presence of 30-min stimulation with native fimbriae or rFimA (1 μg/ml) or positive control agonists (FMLP, 10−7 M; PMA, 0.1 μg/ml). In certain experiments, the cells were pretreated for 30 min with blocking mAbs or pharmacological inhibitors before stimulation. At the end of the 30-min binding time, nonadherent cells were removed by careful washing repeated four times. Cell adhesion was quantified using a fluorescence microplate reader (FL600, Bio-Tek Instruments) with excitation/emission wavelength settings of 485/530 nm, and was expressed as a percentage of total cells added using formula (bound fluorescence/total fluorescence added × 100). HUVEC (between passages 2 and 5) were seeded on 96-well plates at 5 × 104 cells/well and cultured for 2 days to form a confluent monolayer. Calcein AM-labeled monocytes were added at 2 × 105 cells per well of confluent HUVEC monolayer and incubated for 30 min at 37°C and 5% CO2 atmosphere. Nonadherent monocytes were gently washed off and monocyte adhesion was assessed as described using a Bio-Tek fluorescence microplate reader. Wells containing only HUVEC were used to determine background fluorescence, which was minimal and subtracted from each experimental value.

To assess monocyte transmigration across a HUVEC monolayer, we used the Transwell plate system (6.5-mm Transwell inserts with 8.0-μm pores; Corning Costar) and a modification of a previously described protocol (33). Briefly, HUVEC (5 × 104) were seeded in the upper chamber of each transwell and grown for 2 days to form a confluent monolayer. Confluency was confirmed by microscopic inspection. Calcein AM-labeled primary monocytes or THP-1 cells were added to the upper chamber (2 × 105 cells) and were allowed to migrate through the HUVEC monolayer into the lower chamber at 37°C for 3 or 4 h, respectively. Subsequently, the upper chamber was removed to stop transmigration. The fluorescence intensity in the lower compartment was measured using a Bio-Tek fluorescence microplate reader and was expressed as a percentage of total cell-associated fluorescence added in the upper compartment (percentage of transmigrated cells).

PI3K activity was measured as enzymatic production of phosphatidylinositol-3,4,5-trisphosphate (PIP3) from phosphatidylinositol 4,5-bisphosphate (PIP2) substrate by means of a PI3K ELISA kit (Echelon Biosciences) as previously described (27). Briefly, PI3K was immunoprecipitated from cell lysates using anti-PI3K Ab and protein A-agarose beads, and the bead-bound enzyme was subsequently incubated with PIP2 substrate in kinase reaction buffer for 2 h at room temperature. The generation of PIP3 product was determined by competitive ELISA.

The CBRM1/5 epitope induction assay was used to monitor the activation state of CD11b/CD18, as we have previously described (11). The assay is based on the property of the CBRM1/5 mAb to detect a conformational change on CD11b that signifies the high-affinity binding state of CD11b/CD18 (34).

Data were evaluated by ANOVA and the Dunnett multiple-comparison test using the InStat program (GraphPad). Where appropriate (comparison of two groups only), two-tailed t tests were also performed. Statistical differences were considered significant at the level of p < 0.05. Experiments were performed using triplicate samples and were performed twice or more to verify the results.

We have previously shown that P. gingivalis fimbriae induce an activation-specific neoepitope (CBRM1/5) on CD11b, via a novel inside-out signaling pathway involving CD14, TLR2, and PI3K (11). We now investigated the functional significance of this signaling pathway. First, we examined the ability of fimbria-stimulated monocytes to bind well-characterized ligands, such as ICAM-1 and fibrinogen, which are bound by CD11b/CD18 only when this integrin is activated (6). Upon 30-min stimulation at 37°C with 1 μg/ml native fimbriae or rFimA, human monocytes bound efficiently to ICAM-1- or fibrinogen-coated microtiter wells, in contrast to medium-only-treated monocytes that bound poorly (6–16% of the binding activity of stimulated cells; Fig. 1, A and B). When monocytes were pretreated with anti-CD11b mAb (2LPM19c, 10 μg/ml) or with an allosteric antagonist of CD11b/CD18 (XVA143, 1 μM) before stimulation with fimbriae, their ability to bind immobilized ICAM-1 (Fig. 1,A) or fibrinogen (Fig. 1,B) was significantly (p < 0.05) diminished. Pretreatment with IgG1 isotype control or a mAb to an unrelated surface Ag (MHC class I) had no effect in this regard (Fig. 1, A and B). These data indicate that P. gingivalis fimbriae (in native or recombinant form) stimulate monocyte adhesion to immobilized ICAM-1 or fibrinogen in a CD11b/CD18-dependent way.

FIGURE 1.

P. gingivalis fimbriae stimulate CD11b/CD18-dependent monocyte adhesion via TLR2 inside-out signaling. Fluorescently labeled human monocytes were added to 96-well plates coated with ICAM-1 (A, C, and E) or fibrinogen (B, D, and F). The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis native fimbriae or rFimA, or additionally with 0.1 μg/ml PMA (E and F). Before the assays, monocytes were pretreated for 30 min with the indicated inhibitors or control molecules. All mAbs and isotype controls were used at 10 μg/ml; XVA143 (allosteric antagonist of CD11b/CD18) at 1 μM; wortmannin (WTM) at 50 nM; LY294002 and LY30351 (inactive analog) at 20 μM; and GF109203X at 10 μM. After washing to remove nonadherent monocytes, cell adhesion was measured on a microplate fluorescence reader and was expressed as a percentage of total cells added. Results are presented as the mean ± SD of triplicate determinations, from one of two independent sets of experiments that yielded similar findings. Statistically significant (∗, p < 0.05) inhibition of cell adhesion due to various treatments is indicated.

FIGURE 1.

P. gingivalis fimbriae stimulate CD11b/CD18-dependent monocyte adhesion via TLR2 inside-out signaling. Fluorescently labeled human monocytes were added to 96-well plates coated with ICAM-1 (A, C, and E) or fibrinogen (B, D, and F). The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis native fimbriae or rFimA, or additionally with 0.1 μg/ml PMA (E and F). Before the assays, monocytes were pretreated for 30 min with the indicated inhibitors or control molecules. All mAbs and isotype controls were used at 10 μg/ml; XVA143 (allosteric antagonist of CD11b/CD18) at 1 μM; wortmannin (WTM) at 50 nM; LY294002 and LY30351 (inactive analog) at 20 μM; and GF109203X at 10 μM. After washing to remove nonadherent monocytes, cell adhesion was measured on a microplate fluorescence reader and was expressed as a percentage of total cells added. Results are presented as the mean ± SD of triplicate determinations, from one of two independent sets of experiments that yielded similar findings. Statistically significant (∗, p < 0.05) inhibition of cell adhesion due to various treatments is indicated.

Close modal

However, the data described do not necessarily show that the observed fimbria-induced monocyte adhesion is mediated via the CD14-TLR2-PI3K inside-out signaling pathway (11). We thus pretreated monocytes with mAbs to CD14 or TLR2, and determined their ability to bind immobilized ICAM-1 or fibrinogen upon activation with native fimbriae or rFimA. In contrast to isotype controls or mAbs with irrelevant specificities (TLR4 or MHC class I), anti-CD14 or anti-TLR2 significantly inhibited adhesion (p < 0.05; Fig. 1, C and D). The effect of combined anti-TLR2 and anti-TLR4 treatment was not significantly different from the use of anti-TLR2 alone (Fig. 1, C and D). We next determined the effect of PI3K inhibitors on fimbria-stimulated monocyte adhesion. Specifically, pretreatment with wortmannin or LY294002 (but not with its inactive analog, LY303511) resulted in significantly (p < 0.05) reduced cell adhesion to immobilized ICAM-1 or fibrinogen (Fig. 1, E and F, respectively). The inhibitory action of wortmannin or LY294002 could not be attributed to nonspecific toxic effects, because both compounds had no influence on PMA-stimulated cell adhesion, which was however inhibitable by GF109203X, a PKC inhibitor (Fig. 1, E and F).

The involvement of CD14 and TLR2, but not of TLR4, in P. gingivalis fimbria-induced cell adhesion was conclusively shown using pattern-recognition receptor (PRR) knockout mouse macrophages. We found that wild-type or TLR4-deficient macrophages could readily bind to immobilized ICAM-1 or fibrinogen upon stimulation with native fimbriae or rFimA (Fig. 2, A and B). In stark contrast, similarly stimulated macrophages deficient in CD14 or TLR2 or macrophages with combined TLR2 and TLR4 deficiencies failed to display enhanced binding to the same molecules (Fig. 2, A and B). FMLP and PMA were used as controls and their stimulatory effect on macrophage adhesion was not influenced by any PRR deficiency tested (Fig. 2, A and B). The observed cell adhesion to ICAM-1 or fibrinogen was dependent upon CD11b/CD18 as shown by the inhibitory effect of treatments with anti-mouse CD11b or with XVA143 (Fig. 2, C and D). The data from Figs. 1 and 2 collectively indicate that fimbriae interact with CD14 and TLR2 leading to activation of PI3K-mediated intracellular signaling for enhanced CD11b/CD18-dependent cell adhesion.

FIGURE 2.

Adhesion of P. gingivalis fimbria-stimulated mouse macrophages to ICAM-1 or fibrinogen is inhibited by CD14 or TLR2 deficiency. Fluorescently labeled macrophages from wild-type mice or mice deficient in CD14, TLR2, TLR4, or both TLR2 and TLR4 (TLR2/4) were added to 96-well plates coated with ICAM-1 (A and C) or fibrinogen (B and D). The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with P. gingivalis native fimbriae or rFimA (both at 1 μg/ml), or with control agonists (FMLP, 10−7 M; PMA, 0.1 μg/ml). Before the assays (C and D), wild-type macrophages were pretreated for 30 min with 10 μg/ml IgG2b isotype control or anti-CD11b, or with 1 μM XVA143. Cell adhesion was assessed fluorometrically as outlined in the legend to Fig. 1. Data are presented as the mean ± SD (n = 3) from a typical set of experiments that were repeated yielding similar results. Significantly reduced macrophage adhesion (∗, p < 0.05) due to receptor deficiency (A and B) or due to CD11b/CD18 blockade (C and D) is shown.

FIGURE 2.

Adhesion of P. gingivalis fimbria-stimulated mouse macrophages to ICAM-1 or fibrinogen is inhibited by CD14 or TLR2 deficiency. Fluorescently labeled macrophages from wild-type mice or mice deficient in CD14, TLR2, TLR4, or both TLR2 and TLR4 (TLR2/4) were added to 96-well plates coated with ICAM-1 (A and C) or fibrinogen (B and D). The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with P. gingivalis native fimbriae or rFimA (both at 1 μg/ml), or with control agonists (FMLP, 10−7 M; PMA, 0.1 μg/ml). Before the assays (C and D), wild-type macrophages were pretreated for 30 min with 10 μg/ml IgG2b isotype control or anti-CD11b, or with 1 μM XVA143. Cell adhesion was assessed fluorometrically as outlined in the legend to Fig. 1. Data are presented as the mean ± SD (n = 3) from a typical set of experiments that were repeated yielding similar results. Significantly reduced macrophage adhesion (∗, p < 0.05) due to receptor deficiency (A and B) or due to CD11b/CD18 blockade (C and D) is shown.

Close modal

The ability of P. gingivalis fimbriae to induce the CD11b activation-specific CBRM1/5 neoepitope is not affected by the G protein inhibitor, pertussis toxin (PTx), in contrast with PTx-sensitive induction of CBRM1/5 by FMLP (11). Similarly, PTx had no effect on the ability of fimbriae to stimulate monocyte adhesion to immobilized ICAM-1 or fibrinogen, although the same toxin significantly inhibited (p < 0.05) FMLP-stimulated cell adhesion (Figs. 3,A and 4,A). To further characterize the fimbria-induced proadhesive pathway, we have investigated three additional toxins that inhibit integrin activation by interfering with activation signals, namely cholera toxin (CTx), Clostridium difficile toxin B (CdTxB), and Clostridium botulinum C3 transferase exoenzyme (C3 exoenzyme) (35, 36, 37). Following pretreatment with various doses of the toxins, human monocytes were allowed to bind immobilized ICAM-1 (Fig. 3) or fibrinogen (Fig. 4) for 30 min at 37°C in the absence or presence of stimulation with P. gingivalis native fimbriae, FMLP, or PMA. CTx did not significantly influence monocyte adhesion to ICAM-1 (Fig. 3,B) or fibrinogen (Fig. 4,B) regardless of the agonists used for cell stimulation. However, there was a statistically significant linear trend (p < 0.05) regarding the effect of CTx on cell adhesion induced by fimbriae or FMLP, but not with PMA. Specifically, CTx appeared to have a modest enhancing effect on fimbria-induced cell adhesion, whereas the opposite trend was observed for FMLP-induced cell adhesion (Figs. 3,B and 4 B). Interestingly, within the same dose range we tested, CTx acts as a potent inhibitor of P. gingivalis fimbria-induced cytokine release (38).

FIGURE 3.

Toxin sensitivity of stimulated monocyte adhesion to ICAM-1. Fluorescently labeled human monocytes were added to 96-well plates coated with ICAM-1. The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis fimbriae, 10−7 M FMLP, or 0.1 μg/ml PMA. Before the assay, monocytes were pretreated with PTx for 2 h (A); CTx for 2 h (B); CdTxB for 2 h (C); and C3 exoenzyme for 24 h (D) at the indicated doses. After washing to remove nonadherent monocytes, cell adhesion was assessed fluorometrically and was expressed as a percentage of total cells added. Results are presented as the mean ± SD of triplicate determinations, from one of two independent sets of experiments that yielded similar findings. Statistically significant (∗, p < 0.05) inhibition of cell adhesion due to toxin treatment is indicated.

FIGURE 3.

Toxin sensitivity of stimulated monocyte adhesion to ICAM-1. Fluorescently labeled human monocytes were added to 96-well plates coated with ICAM-1. The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis fimbriae, 10−7 M FMLP, or 0.1 μg/ml PMA. Before the assay, monocytes were pretreated with PTx for 2 h (A); CTx for 2 h (B); CdTxB for 2 h (C); and C3 exoenzyme for 24 h (D) at the indicated doses. After washing to remove nonadherent monocytes, cell adhesion was assessed fluorometrically and was expressed as a percentage of total cells added. Results are presented as the mean ± SD of triplicate determinations, from one of two independent sets of experiments that yielded similar findings. Statistically significant (∗, p < 0.05) inhibition of cell adhesion due to toxin treatment is indicated.

Close modal
FIGURE 4.

Toxin sensitivity of stimulated monocyte adhesion to fibrinogen. Fluorescently labeled human monocytes were added to 96-well plates coated with fibrinogen. The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis fimbriae, 10−7 M FMLP, or 0.1 μg/ml PMA. Before the assay, monocytes were pretreated with PTx for 2 h (A); CTx for 2 h (B); CdTxB for 2 h (C); and C3 exoenzyme for 24 h (D) at the indicated doses. Cell adhesion was assessed as shown in Fig. 3. Data are presented as the mean ± SD (n = 3), from one of two independent sets of experiments that yielded similar results. Statistically significant (∗, p < 0.05) inhibition of cell adhesion due to toxin treatment is indicated.

FIGURE 4.

Toxin sensitivity of stimulated monocyte adhesion to fibrinogen. Fluorescently labeled human monocytes were added to 96-well plates coated with fibrinogen. The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis fimbriae, 10−7 M FMLP, or 0.1 μg/ml PMA. Before the assay, monocytes were pretreated with PTx for 2 h (A); CTx for 2 h (B); CdTxB for 2 h (C); and C3 exoenzyme for 24 h (D) at the indicated doses. Cell adhesion was assessed as shown in Fig. 3. Data are presented as the mean ± SD (n = 3), from one of two independent sets of experiments that yielded similar results. Statistically significant (∗, p < 0.05) inhibition of cell adhesion due to toxin treatment is indicated.

Close modal

In contrast to CTx, CdTxB had a profound inhibitory effect (p < 0.05) on both fimbria- and FMLP-stimulated monocyte adhesion to immobilized ICAM-1 or fibrinogen (Figs. 3,C or 4,C, respectively). However, suppression of PMA-induced cell adhesion by CdTxB did not reach statistical significance, despite a significant linear trend with increasing toxin dose (Figs. 3,C and 4,C). The C3 exoenzyme had no effect, whatsoever, on fimbria-stimulated cell adhesion, although the same toxin significantly inhibited (p < 0.05) the activity of FMLP and PMA (Figs. 3,D and 4 D). The toxins did not influence basal cell adhesion, which was ≤8% of that seen in the presence of agonists (data not shown). These data suggest that the signaling pathway involved in P. gingivalis fimbria-stimulated monocyte adhesion to a ligand (fibrinogen) or a counterreceptor (ICAM-1) of CD11b/CD18 is distinct from those stimulated by FMLP or PMA. The fimbria-stimulated pathway is sensitive to the action of CdTxB but is not affected by PTx, CTx, or C3 exoenzyme.

CdTxB is known to inhibit the small molecular weight GTPases, Rho, Rac, and Cdc42, whereas C3 exoenzyme specifically inhibits Rho (RhoA, B, and C), but not Rac or Cdc42 (39). Therefore, on the basis of the toxin sensitivity data (Figs. 3 and 4), we hypothesized that the proadhesive pathway activated by P. gingivalis fimbriae involves participation of Rac or Cdc42, but not of Rho GTPase. To test this hypothesis, we determined the ability of human monocytic THP-1/CD14 cells to bind immobilized ICAM-1 or fibrinogen, upon cell transfection with empty vector control or with DN inhibitors of Rac1, Cdc42, or RhoA. We found that THP-1/CD14 cells transfected with Rac1-DN, but not with Cdc42-DN or RhoA-DN, displayed significantly reduced adhesion to ICAM-1 in response to fimbriae (p < 0.05, compared with empty vector-transfected cells) (Fig. 5,A). Similar results were obtained when adhesion was tested on fibrinogen-coated plates (data not shown). In a parallel experiment using similarly transfected but FMLP-stimulated THP-1/CD14 cells, cell adhesion to ICAM-1 was significantly inhibited (p < 0.05) by RhoA-DN but not by Rac1-DN or Cdc42-DN (Fig. 5 A). Therefore, Rac1 appears to be a second signaling intermediate, in addition to PI3K, involved in P. gingivalis fimbria-stimulated cell adhesion to ICAM-1. These data further support that P. gingivalis fimbriae and FMLP activate distinct intracellular signaling pathways, involving Rac1 and RhoA, respectively, for CD11b/CD18 activation.

FIGURE 5.

Rac1 is involved in P. gingivalis fimbria-induced cell adhesion to ICAM-1 (A) and stimulation of the lipid kinase activity of PI3K (B). A, THP-1/CD14 cells transfected with empty vector control or with DN mutants of Rac1, Cdc42, or RhoA (at the indicated microgram amounts of plasmid DNA per 2 × 105 cells) were fluorescently labeled and added to 96-well plates coated with ICAM-1. The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis fimbriae or 10−7 M FMLP. After washing to remove nonadherent monocytes, cell adhesion was assessed fluorometrically and was expressed as a percentage of total cells added. B, THP-1/CD14 cells transfected with DN mutants of TLR2, Rac1, or RhoA were stimulated for 30 min with 1 μg/ml fimbriae. Subsequently, PI3K was immunoprecipitated from cell lysates and its enzymatic activity was assessed as described in Materials and Methods. Data are presented as the mean ± SD (n = 3), from one of two (A) or three (B) independent sets of experiments that yielded similar results. Statistically significant (∗, p < 0.05) inhibition of cell adhesion (A) or of PIP3 production (B) due to transfection with DN mutants is indicated.

FIGURE 5.

Rac1 is involved in P. gingivalis fimbria-induced cell adhesion to ICAM-1 (A) and stimulation of the lipid kinase activity of PI3K (B). A, THP-1/CD14 cells transfected with empty vector control or with DN mutants of Rac1, Cdc42, or RhoA (at the indicated microgram amounts of plasmid DNA per 2 × 105 cells) were fluorescently labeled and added to 96-well plates coated with ICAM-1. The cells were allowed to bind for 30 min at 37°C in the absence or presence of stimulation with 1 μg/ml P. gingivalis fimbriae or 10−7 M FMLP. After washing to remove nonadherent monocytes, cell adhesion was assessed fluorometrically and was expressed as a percentage of total cells added. B, THP-1/CD14 cells transfected with DN mutants of TLR2, Rac1, or RhoA were stimulated for 30 min with 1 μg/ml fimbriae. Subsequently, PI3K was immunoprecipitated from cell lysates and its enzymatic activity was assessed as described in Materials and Methods. Data are presented as the mean ± SD (n = 3), from one of two (A) or three (B) independent sets of experiments that yielded similar results. Statistically significant (∗, p < 0.05) inhibition of cell adhesion (A) or of PIP3 production (B) due to transfection with DN mutants is indicated.

Close modal

Rac1 and PI3K regulate cellular function through various, often overlapping, signaling pathways and either of the two intracellular enzymes can activate the other, depending on the specific pathway involved (40). We examined whether Rac1-DN could additionally inhibit fimbria-induced PI3K activation to determine whether Rac1 acts upstream of PI3K. Specifically, we examined whether Rac1-DN inhibits the lipid kinase activity of PI3K, monitored through the generation of PIP3 from PIP2 substrate. TLR2-DN and RhoA-DN were used as positive and negative controls, respectively. We found that Rac1-DN and TLR2-DN (but not RhoA-DN) could significantly inhibit (p < 0.05) the ability of fimbriae to activate PI3K (Fig. 5 B). Therefore, Rac1 appears to be a signaling intermediate, acting between TLR2 and PI3K, in the P. gingivalis fimbria-stimulated pathway for CD11b/CD18-dependent cell adhesion.

Our findings that P. gingivalis fimbriae stimulate monocyte adhesion to ICAM-1, a major endothelial receptor, suggested that fimbriae may similarly up-regulate monocyte adhesion to HUVEC. To investigate this possibility, monocytes were added with or without fimbriae (1 μg/ml) to a HUVEC monolayer that was previously either activated (by 1 ng/ml IL-1β for 16 h) or maintained unstimulated (medium only). Unstimulated or IL-1β-stimulated HUVEC were washed before addition of monocytes to remove IL-1β from the coculture system. We found that the ability of monocytes to adhere to the HUVEC was significantly (p < 0.05) higher in the presence of fimbriae than in the presence of medium only, regardless of whether the HUVECs were prestimulated with IL-1β (Fig. 6,A). However, monocyte adhesion to IL-1β-stimulated HUVECs was significantly enhanced (p < 0.05) compared with unstimulated HUVECs (Fig. 6,A). When monocytes were preincubated with anti-CD11b mAb or the XVA143 allosteric antagonist of CD11b/CD18 before being exposed to fimbriae and added to the HUVEC monolayer, their adhesive activity was significantly reduced (p < 0.05; Fig. 6,B). In contrast, an IgG1 isotype control or a mAb to an unrelated surface Ag (MHC class I) were without effect in this regard (Fig. 6,B). Furthermore, when unstimulated or IL-1β-stimulated HUVECs were preincubated with anti-ICAM-1 mAb before addition of monocytes and fimbriae, their ability to support monocyte adhesion was significantly decreased (p < 0.05; Fig. 6,C). In contrast, control treatments (isotype control or irrelevant mAb) had no effect (Fig. 6 C). These data collectively suggest that P. gingivalis fimbriae promote monocyte adhesion to HUVEC in a CD11b/CD18- and ICAM-1-dependent way.

FIGURE 6.

P. gingivalis fimbriae promote monocyte adhesion to HUVEC in a CD11b/CD18- and ICAM-1-dependent way. Confluent HUVEC monolayers were stimulated with IL-1β (1 ng/ml; 16 h) or not and washed before addition of fluorescently labeled monocytes (2 × 105 per well) with or without P. gingivalis fimbriae (1 μg/ml). The monocytes were allowed to bind for 30 min at 37°C. B, Monocytes were preincubated for 30 min with IgG1 isotype control, anti-CD11b mAb, or anti-MHC class I (all at 10 μg/ml) or an allosteric antagonist of CD11b/CD18 (XVA413, 1 μM) before exposure to fimbriae and addition to the HUVEC monolayer. C, HUVEC were preincubated for 30 min with IgG1 isotype control, anti-ICAM-1 mAb, or anti-MHC class I mAb (all at 10 μg/ml) before addition of monocytes and fimbriae. After a 30-min incubation of monocytes with HUVEC, nonadherent monocytes were gently washed off and monocyte adhesion was measured fluorometrically and expressed as a percentage of total cells added. Results are presented as the mean ± SD (n = 3), from one of three (A) or two (B and C) independent experiments that yielded similar findings. Statistically significant (∗, p < 0.05) differences in cell adhesion due to differential stimulation treatments (A) or due to the use of inhibitory treatments (B and C) are indicated.

FIGURE 6.

P. gingivalis fimbriae promote monocyte adhesion to HUVEC in a CD11b/CD18- and ICAM-1-dependent way. Confluent HUVEC monolayers were stimulated with IL-1β (1 ng/ml; 16 h) or not and washed before addition of fluorescently labeled monocytes (2 × 105 per well) with or without P. gingivalis fimbriae (1 μg/ml). The monocytes were allowed to bind for 30 min at 37°C. B, Monocytes were preincubated for 30 min with IgG1 isotype control, anti-CD11b mAb, or anti-MHC class I (all at 10 μg/ml) or an allosteric antagonist of CD11b/CD18 (XVA413, 1 μM) before exposure to fimbriae and addition to the HUVEC monolayer. C, HUVEC were preincubated for 30 min with IgG1 isotype control, anti-ICAM-1 mAb, or anti-MHC class I mAb (all at 10 μg/ml) before addition of monocytes and fimbriae. After a 30-min incubation of monocytes with HUVEC, nonadherent monocytes were gently washed off and monocyte adhesion was measured fluorometrically and expressed as a percentage of total cells added. Results are presented as the mean ± SD (n = 3), from one of three (A) or two (B and C) independent experiments that yielded similar findings. Statistically significant (∗, p < 0.05) differences in cell adhesion due to differential stimulation treatments (A) or due to the use of inhibitory treatments (B and C) are indicated.

Close modal

We next examined the ability of fimbria-stimulated monocytes for transmigration across HUVEC monolayers. For this purpose, monocytes were added with or without fimbriae (1 μg/ml) to the upper chamber of Transwells containing HUVEC monolayers, and the incubation was conducted for 3 h. We found that the ability of monocytes for transmigration was significantly enhanced (p < 0.05) in the presence of fimbriae than in the presence of medium only (Fig. 7,A). However, when monocytes were preincubated with anti-CD11b mAb or the XVA143 antagonist before being exposed to fimbriae and added to HUVEC, their ability for transendothelial migration was significantly inhibited (p < 0.05; Fig. 7,B). On the contrary, an IgG1 isotype control or mAb with irrelevant specificity (MHC class I) did not affect monocyte transmigration (Fig. 7,B). Furthermore, pretreatment of HUVEC with anti-ICAM-1 mAb before addition of monocytes and fimbriae resulted in significantly reduced (p < 0.05) monocyte transmigration, whereas control treatments had no effect in this regard (Fig. 7 C). These data jointly suggest that P. gingivalis fimbriae promote transendothelial migration of monocytes in a CD11b/CD18- and ICAM-1-dependent mode.

FIGURE 7.

P. gingivalis fimbriae promote monocyte transmigration through HUVEC monolayers in a CD11b/CD18- and ICAM-1-dependent way. A, Fluorescently labeled monocytes (2 × 105) were added with or without P. gingivalis fimbriae (1 μg/ml) to confluent HUVEC monolayers in the upper chamber of Transwells. B, Monocytes were preincubated for 30 min with IgG1 isotype control, anti-CD11b mAb, or anti-MHC class I (all at 10 μg/ml) or an allosteric antagonist of CD11b/CD18 (XVA413, 1 μM) before exposure to fimbriae and addition to the HUVEC monolayer. C, HUVECs were preincubated for 30 min with IgG1 isotype control, anti-ICAM-1 mAb, or anti-MHC class I (all at 10 μg/ml) before addition of monocytes and fimbriae. The monocytes were allowed to migrate into the lower chamber at 37°C for 3 h. Subsequently, the fluorescence intensity in the lower chamber was measured and expressed as a percentage of total cell-associated fluorescence added in the upper chamber (percentage of transmigrated cells). Results are presented as the mean ± SD of triplicate determinations from one of three (A) or two (B and C) independent experiments that yielded similar findings. Statistically significant (∗, p < 0.05) differences in monocyte transmigration due to differential stimulation treatments (A) or due to the use of inhibitory treatments (B and C) are indicated.

FIGURE 7.

P. gingivalis fimbriae promote monocyte transmigration through HUVEC monolayers in a CD11b/CD18- and ICAM-1-dependent way. A, Fluorescently labeled monocytes (2 × 105) were added with or without P. gingivalis fimbriae (1 μg/ml) to confluent HUVEC monolayers in the upper chamber of Transwells. B, Monocytes were preincubated for 30 min with IgG1 isotype control, anti-CD11b mAb, or anti-MHC class I (all at 10 μg/ml) or an allosteric antagonist of CD11b/CD18 (XVA413, 1 μM) before exposure to fimbriae and addition to the HUVEC monolayer. C, HUVECs were preincubated for 30 min with IgG1 isotype control, anti-ICAM-1 mAb, or anti-MHC class I (all at 10 μg/ml) before addition of monocytes and fimbriae. The monocytes were allowed to migrate into the lower chamber at 37°C for 3 h. Subsequently, the fluorescence intensity in the lower chamber was measured and expressed as a percentage of total cell-associated fluorescence added in the upper chamber (percentage of transmigrated cells). Results are presented as the mean ± SD of triplicate determinations from one of three (A) or two (B and C) independent experiments that yielded similar findings. Statistically significant (∗, p < 0.05) differences in monocyte transmigration due to differential stimulation treatments (A) or due to the use of inhibitory treatments (B and C) are indicated.

Close modal

Under the experimental conditions used, it was possible that fimbriae also activated the HUVEC. Indeed, the incubation time (3 h) appeared adequate for fimbria-induced up-regulation of ICAM-1 expression in HUVEC (41). We therefore set out to confirm that fimbria-induced transendothelial migration of monocytes was mediated, at least in part, by direct effects of fimbriae on monocytes (i.e., through induction of CD11b/CD18 activation). To this end, we compared the transmigration activity of monocytic THP-1/CD14 cells to that of CD14-nonexpressing THP-1/RSV cells. The rationale was that fimbriae would not effectively induce CD11b/CD18 activation in the latter cell line (due to diminished CD14 expression, required for fimbria-induced inside-out signaling (11)), resulting in reduced transmigration activity of THP-1/RSV cells compared with THP-1/CD14 cells. Indeed, fimbriae readily induced CD11b/CD18 activation in THP-1/CD14 cells (but not in THP-1/RSV cells), as evidenced by induction of the CD11b activation-specific CBRM1/5 neoepitope (Fig. 8,A). Moreover, the fimbria-induced transendothelial migration activity of THP-1/CD14 cells was significantly higher (p < 0.05) compared with that of THP-1/RSV cells (Fig. 8,B), confirming that CD11b/CD18 activation plays an important role in the transmigration process. The relative inability of THP-1/RSV cells to respond to fimbriae was not due to any inherent defects in this cell line. Indeed, when PMA was used instead as an agonist, both cell lines could equally well induce the CBRM1/5 neoepitope (Fig. 8,A) and stimulate transmigration (Fig. 8 B).

FIGURE 8.

CD11b/CD18 activation is important for transendothelial migration of THP-1 cells in response to P. gingivalis fimbriae. A, THP-1/CD14 or CD14-nonexpressing THP-1/RSV cells were incubated with medium only, fimbriae (1 μg/ml), or PMA (0.1 μg/ml). After 30 min, the cells were assessed for induction of a CD11b/CD18 activation-specific neoepitope (CBRM1/5) by staining with FITC-labeled CBRM1/5 mAb. Cell-associated fluorescence was measured and expressed in relative fluorescence units (RFU). B, Fluorescently labeled THP-1/CD14 and THP-1/RSV cells were added with or without fimbriae (1 μg/ml) or PMA (0.1 μg/ml) to the upper chamber of Transwells containing HUVEC monolayers, and the incubation was conducted for 4 h. The percentage of transmigrated cells was determined as outlined in the legend to Fig. 7. Results are presented as the mean ± SD (n = 3) from one of two (A) or three (B) independent set of experiments that yielded similar findings. Significantly enhanced transmigration (∗, p < 0.05) compared with indicated control is shown.

FIGURE 8.

CD11b/CD18 activation is important for transendothelial migration of THP-1 cells in response to P. gingivalis fimbriae. A, THP-1/CD14 or CD14-nonexpressing THP-1/RSV cells were incubated with medium only, fimbriae (1 μg/ml), or PMA (0.1 μg/ml). After 30 min, the cells were assessed for induction of a CD11b/CD18 activation-specific neoepitope (CBRM1/5) by staining with FITC-labeled CBRM1/5 mAb. Cell-associated fluorescence was measured and expressed in relative fluorescence units (RFU). B, Fluorescently labeled THP-1/CD14 and THP-1/RSV cells were added with or without fimbriae (1 μg/ml) or PMA (0.1 μg/ml) to the upper chamber of Transwells containing HUVEC monolayers, and the incubation was conducted for 4 h. The percentage of transmigrated cells was determined as outlined in the legend to Fig. 7. Results are presented as the mean ± SD (n = 3) from one of two (A) or three (B) independent set of experiments that yielded similar findings. Significantly enhanced transmigration (∗, p < 0.05) compared with indicated control is shown.

Close modal

Rac1, but not RhoA, is required for enhanced THP-1/CD14 adhesion to ICAM-1 in response to P. gingivalis fimbriae (Fig. 5). We now determined whether Rac-1 is similarly involved in the ability of fimbria-stimulated THP-1/CD14 cells to adhere to HUVEC and transmigrate across the HUVEC monolayer, and moreover, examined whether these activities correlate with CD11b/CD18 activation. For this purpose, we used THP-1/CD14 cells transiently transfected with Rac1-DN. Cells transfected with TLR2-DN or RhoA-DN were used as positive or negative controls, respectively. We found that cells transfected with Rac1-DN or TLR2-DN displayed significantly reduced ability (p < 0.05) for CD11b/CD18 activation (CBRM1/5 epitope induction), adhesion to HUVEC, and transendothelial migration in response to P. gingivalis fimbriae, as compared with untransfected cells or cells transfected with empty vector control (Fig. 9). Transfection with Rac1-DN or TLR2-DN had no effect on surface expression of CD11b/CD18, as shown by staining with Bear-1, a mAb that detects CD11b regardless of its activation state (data not shown). In contrast to THP-1/CD14 cells transfected with Rac1-DN or TLR2-DN, transfection with RhoA-DN did not influence their ability for CD11b/CD18 activation and adhesion to HUVEC (Fig. 9). However, the transmigrating activity of RhoA-DN-transfected cells was similarly affected and was significantly diminished (p < 0.05) relative to untransfected or empty vector-transfected cells (Fig. 9). Therefore, RhoA may be involved in fimbria-induced monocyte transmigration by acting on a process that is independent of CD11b/CD18 activation and adhesion to HUVEC. These data demonstrate that Rac1 is an essential component of the P. gingivalis fimbria-induced inside-out signaling pathway that leads to enhanced monocyte adhesion to HUVEC and transendothelial migration.

FIGURE 9.

Rac1 is involved in induction of CD11b/CD18 activation, adhesion to HUVEC, and transendothelial migration of fimbria-stimulated monocytic cells. THP-1/CD14 cells, transfected with empty vector control (EVC) or with DN mutants of TLR2, Rac1, or RhoA (at the indicated microgram amounts of plasmid DNA per 2 × 105 cells), were stimulated with 1 μg/ml P. gingivalis fimbriae and tested for induction of CD11b/CD18 activation, adhesion to HUVECs, and transendothelial migration. Results were normalized to the activity of untransfected THP-1/CD14 cells, and are presented as the mean ± SD (n = 3) from one of two independent sets of experiments that yielded similar findings. Statistically significant inhibition (∗, p < 0.05) of cell activity due to transfection with DN mutants is indicated.

FIGURE 9.

Rac1 is involved in induction of CD11b/CD18 activation, adhesion to HUVEC, and transendothelial migration of fimbria-stimulated monocytic cells. THP-1/CD14 cells, transfected with empty vector control (EVC) or with DN mutants of TLR2, Rac1, or RhoA (at the indicated microgram amounts of plasmid DNA per 2 × 105 cells), were stimulated with 1 μg/ml P. gingivalis fimbriae and tested for induction of CD11b/CD18 activation, adhesion to HUVECs, and transendothelial migration. Results were normalized to the activity of untransfected THP-1/CD14 cells, and are presented as the mean ± SD (n = 3) from one of two independent sets of experiments that yielded similar findings. Statistically significant inhibition (∗, p < 0.05) of cell activity due to transfection with DN mutants is indicated.

Close modal

We then determined whether fimbriae could activate monocyte transendothelial migration in cell-associated form. This examination used whole cells of P. gingivalis from wild-type strains 381 and A7436 or their respective isogenic non-fimbriated mutants. We found that the wild-type strains were significantly more potent (p < 0.05) than their non-fimbriated mutants in stimulating monocyte adhesion to HUVEC and transmigration across HUVEC monolayers (Fig. 10). These data suggest that P. gingivalis has the potential to contribute to the inflammatory processes in atherosclerosis by stimulating monocyte recruitment into subendothelial areas.

FIGURE 10.

The ability of P. gingivalis to promote monocyte adhesion (A) and transmigration through HUVEC monolayers (B) correlates with its fimbriation state. Fluorescently labeled monocytes (2 × 105) were added to confluent HUVEC monolayers and assayed for adhesion or transmigration. The monocytes were added in the presence or absence of wild-type (Wt) P. gingivalis strains 381 or A7436 or their non-fimbriated isogenic mutants (Mt) at the indicated multiplicity of infection. Results are presented as the mean ± SD (n = 3) from one of two independent experiments that yielded similar findings. Statistically significant enhancement (∗, p < 0.05) of monocyte adhesion or transmigration by wild-type P. gingivalis compared with corresponding mutant is shown.

FIGURE 10.

The ability of P. gingivalis to promote monocyte adhesion (A) and transmigration through HUVEC monolayers (B) correlates with its fimbriation state. Fluorescently labeled monocytes (2 × 105) were added to confluent HUVEC monolayers and assayed for adhesion or transmigration. The monocytes were added in the presence or absence of wild-type (Wt) P. gingivalis strains 381 or A7436 or their non-fimbriated isogenic mutants (Mt) at the indicated multiplicity of infection. Results are presented as the mean ± SD (n = 3) from one of two independent experiments that yielded similar findings. Statistically significant enhancement (∗, p < 0.05) of monocyte adhesion or transmigration by wild-type P. gingivalis compared with corresponding mutant is shown.

Close modal

It has been speculated that inside-out signaling mechanisms evolved to resolve two competing objectives (42). On the one hand, there is need for highly mobile leukocytes to roam and detect potential threats; in contrast, these wandering cells should hold fast to sites of infection they encounter and thus need to be rapidly reprogrammable. In this context, we consider that TLRs are appropriate transmodulators of the adhesive activities of leukocytes because these PRRs can both detect infection and transduce activating intracellular signals (43). Our identification of a novel, TLR2-mediated inside-out signaling pathway (11) and, most importantly, the current demonstration that this pathway regulates the adhesive and transmigrating activities of human monocytes clearly support this notion. CD11b/CD18 and β2 integrins in general are appropriate effectors of such proadhesive pathways due to their ability to engage, once activated, diverse ligands or counterreceptors implicated in the inflammatory response (1, 2, 4, 6). The significance of inside-out signaling in the regulation of leukocyte adhesion was emphasized by the discovery of an alternative form of leukocyte adhesion deficiency (44). In this type of deficiency, β1, β2, and β3 integrins are normally expressed on the cell surface but fail to be activated by intracellular signaling pathways to bind ligands (44).

This study showed that P. gingivalis fimbriae stimulate monocyte adhesion to β2 integrin ligands (Fig. 1) or to endothelial cells (Fig. 6) and activate monocyte transendothelial migration (Figs. 7 and 8,B) through induction of inside-out signaling. On the basis of previous (11) and current data, this proadhesive signaling pathway involves the sequential participation of CD14, TLR2, Rac1, PI3K, and the CD11b/CD18 (Fig. 11). Initial evidence that this mechanism of CD11b/CD18 activation involves a distinct intracellular signaling pathway from those induced by FMLP or PMA came from data of differential toxin sensitivity (Figs. 3 and 4). The novel pathway activated by P. gingivalis fimbriae was sensitive to the action of CdTxB, but was not influenced by C3 exoenzyme, PTx, or CTx. These findings pointed to a possible participation of Rac1 in this pathway because Rac1 is inhibitable by CdTxB but not by C3 exoenzyme (39). At the same time, these data suggested that RhoA was not a likely signaling molecule candidate in the fimbria-induced pathway because RhoA is inhibitable by both CdTxB and C3 exoenzyme (39). Experiments examining the effects of DN signaling mutants on the ability of fimbriae to induce CD11b/CD18-dependent monocyte adhesive activities confirmed the importance of Rac1 and the irrelevance of RhoA, although the latter was important for the FMLP-induced proadhesive pathway. An additional difference regarding the mechanisms whereby fimbriae and FMLP stimulate monocyte adhesion involves the lack of PI3K requirement in the case of FMLP. Specifically, although FMLP activates PI3K in our experimental system, the use of specific PI3K inhibitors (wortmannin or LY294002) does not inhibit the ability of FMLP to stimulate monocyte adhesion (our unpublished observation). Therefore, PI3K is not a point of convergence in the proadhesive pathways activated by fimbriae or FMLP. FMLP-activated PI3K may mediate other effector functions in monocytes, such as induction of NF-κB activation (45). The ability of fimbriae to stimulate monocyte transendothelial migration was inhibited not only by the Rac1-DN but also by the RhoA-DN mutant (Fig. 9 C). Because RhoA is not involved in fimbria-induced activation of CD11b/CD18 and monocyte adhesion to endothelial cells, this finding appeared somewhat unexpected. However, RhoA was previously found to be essential for the retraction of the tail of the migrating monocyte to complete diapedesis (33) and this may account for the inhibitory effect of the RhoA-DN mutant in our transmigration model.

FIGURE 11.

P. gingivalis fimbria-activated TLR2 transmodulates the adhesive activity of monocytes. P. gingivalis fimbriae interact with CD14 and TLR2 and induce Rac1- and PI3K-mediated inside-out signaling leading to activation of the ligand-binding capacity of CD11b/CD18. Activated CD11b/CD18 can thereby bind endothelial ICAM-1, thus promoting monocyte-endothelial cell interactions.

FIGURE 11.

P. gingivalis fimbria-activated TLR2 transmodulates the adhesive activity of monocytes. P. gingivalis fimbriae interact with CD14 and TLR2 and induce Rac1- and PI3K-mediated inside-out signaling leading to activation of the ligand-binding capacity of CD11b/CD18. Activated CD11b/CD18 can thereby bind endothelial ICAM-1, thus promoting monocyte-endothelial cell interactions.

Close modal

Rac1 and other small GTPases of the Rho family are regulated by GTP/GDP exchange and function as molecular switches that control signaling pathways involved in kinase regulation, gene transcription, cytoskeleton organization, cell motility, and other cellular processes (46, 47). The property of Rac1 to function as a molecular on-off switch is consistent with its involvement in P. gingivalis fimbria-induced inside-out signaling pathway, which needs to be rapidly and transiently activated. PI3K is also a component of this signaling pathway and appears to act downstream of Rac1 (Fig. 11). In this regard, it was shown that Rac1, but not RhoA, can bind PI3K and augment its activity (48). These findings are in line with our observations that a DN inhibitor of Rac1, but not of RhoA, inhibits PI3K activity (Fig. 5 B). Two PI3K binding motifs are present on the TLR2 cytoplasmic tail and PI3K is recruited to TLR2 upon activation with heat-killed Staphylococcus aureus (49). This pathway proceeds downstream of PI3K through the Ser/Thr kinase Akt and results in NF-κB activation (49). Rac1 is also essential for NF-κB activation in the S. aureus-stimulated TLR2 pathway and it similarly acts upstream of PI3K (49). Therefore, it appears that P. gingivalis fimbriae and heat-killed S. aureus may both activate a TLR2-Rac1-PI3K pathway, which may bifurcate at the level of PI3K; downstream activation of Akt induces NF-κB-dependent transcription (49), whereas stimulation of alternative PI3K effectors may result in activation of CD11b/CD18-dependent adhesion as seen in the present study.

The ability of CD14/TLR2 to detect P. gingivalis fimbriae and initiate inside-out signaling for CD11b/CD18 activation is a potentially protective mechanism, which can contribute to monocyte recruitment to sites of P. gingivalis infection. Fimbriae could stimulate this proadhesive pathway in bacterial cell-associated form or more effectively as free molecules shed from the bacterial cell surface or as components of released outer membrane vesicles that can readily infiltrate tissues (50). However, the ability of P. gingivalis fimbria-stimulated TLR2 to transmodulate the adhesive activity of CD11b/CD18 may also constitute a potentially harmful mechanism contributing to destructive inflammation in conditions associated with P. gingivalis. Evidence from biopsy studies on human carotid endarterectomy specimens or from experimental atherosclerosis in a mouse model suggests that P. gingivalis can localize to sites of atheroma development (24, 26). Moreover, epidemiological and experimental evidence suggests that periodontal disease and P. gingivalis may be risk factors contributing to the pathogenesis of atherosclerosis (20, 21, 22, 26, 51, 52). Because monocyte recruitment into the subendothelium is a crucial step in atherosclerosis (14), our findings that P. gingivalis stimulates monocyte transendothelial migration suggest a plausible mechanistic link between this pathogen and atherogenesis. Furthermore, our observation that wild-type P. gingivalis is significantly more potent than non-fimbriated mutants in this proinflammatory activity is consistent with earlier findings that fimbriation of P. gingivalis is an essential virulence attribute of this pathogen for stimulating atherosclerotic plaque formation in orally infected hyperlipidemic mice (26). The prominent role of fimbriae in mediating P. gingivalis-induced host responses may be attributable to their binding versatility as well as hydrophobicity and polymeric nature, which can thereby promote the host-P. gingivalis molecular cross-talk in both specific ways as well as through increased avidity of interactions (26, 27, 53).

Besides periodontitis, other infection-driven chronic inflammatory diseases have also been implicated as contributory factors in the pathogenesis of atherosclerosis (14, 18, 54, 55). C. pneumoniae, a respiratory pathogen associated with atherosclerosis (54), induces adhesion of human monocytes to aortic endothelial cells in vitro (56) and activates recruitment of mouse macrophages to the carotid artery in vivo (57). C. pneumoniae promotes monocyte-endothelial interactions through integrin activation in monocytes (56, 57) or through up-regulation of ICAM-1 expression in endothelial cells (58). This pathogen also induces foam cell formation, characteristic of atherosclerotic lesions (55). In addition to our findings on monocyte activation by P. gingivalis, others have shown that this oral pathogen invades endothelial cells and up-regulates ICAM-1 expression (41). This function is similarly dependent on the presence of fimbriae (41). An additional mechanism whereby P. gingivalis may promote atherogenesis is via induction of foam cell formation (59). Indeed, infection of human monocyte-derived macrophages with P. gingivalis in the presence of low-density lipoprotein results in foam cell formation (59). This process is augmented when macrophages are exposed to wild-type P. gingivalis rather than a non-fimbriated mutant (59). The parallels between P. gingivalis and C. pneumoniae suggest that similar pathogenic mechanisms induced by different infectious agents may link chronic inflammatory conditions with the development of cardiovascular disease. However, this investigation is the first time that pathogen-induced TLR-dependent inside-out signaling has been demonstrated and suggested as a possible mechanism for infection-driven monocyte accumulation into the subendothelium.

In conclusion, innate recognition of P. gingivalis fimbriae results in activation of TLR2, which transmodulates the adhesive activity of CD11b/CD18 via Rac1 and PI3K (Fig. 11). Activated CD11b/CD18 renders the monocytes capable of binding endothelial ICAM-1 and transmigrating across endothelial cells. Because monocyte recruitment into subendothelial areas plays an important role in the early steps of atherogenesis, the inducible proadhesive pathway described in this study may form a mechanistic basis linking P. gingivalis to inflammatory atherosclerotic processes.

We thank Dr. Sarah Gaffen (University at Buffalo, Buffalo, NY) for critical review 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 U.S. Public Health Service Grant DE015254 from the National Institutes of Health.

3

Abbreviations used in this paper: DN, dominant negative; PRR, pattern-recognition receptor; PTx, pertussis toxin; CTx, cholera toxin; CdTxB, C. difficile toxin B; PIP3, phosphatidylinositol-3,4,5-trisphosphate; PIP2, phosphatidylinositol 4,5-bisphosphate.

1
Shimaoka, M., J. Takagi, T. A. Springer.
2002
. Conformational regulation of integrin structure and function.
Annu. Rev. Biophys. Biomol. Struct.
31
:
485
-516.
2
Ehlers, M. R. W..
2000
. CR3: a general purpose adhesion-recognition receptor essential for innate immunity.
Microbes Infect.
2
:
289
-294.
3
Carlos, T. M., J. M. Harlan.
1994
. Leukocyte-endothelial adhesion molecules.
Blood
84
:
2068
-2101.
4
Yakubenko, V. P., V. K. Lishko, S. C. Lam, T. P. Ugarova.
2002
. A molecular basis for integrin αMβ2 ligand binding promiscuity.
J. Biol. Chem.
277
:
48635
-48642.
5
Wang, Y., M. Sakuma, Z. Chen, V. Ustinov, C. Shi, K. Croce, A. C. Zago, J. Lopez, P. Andre, E. Plow, D. I. Simon.
2005
. Leukocyte engagement of platelet glycoprotein Ibα via the integrin Mac-1 is critical for the biological response to vascular injury.
Circulation
112
:
2993
-3000.
6
Diamond, M. S., J. Garcia-Aguilar, J. K. Bickford, A. L. Corbi, T. A. Springer.
1993
. The I domain is a major recognition site on the leukocyte integrin Mac-1 (CD11b/CD18) for four distinct adhesion ligands.
J. Cell Biol.
120
:
1031
-1043.
7
Meerschaert, J., M. B. Furie.
1995
. The adhesion molecules used by monocytes for migration across endothelium include CD11a/CD18, CD11b/CD18, and VLA-4 on monocytes and ICAM-1, VCAM-1, and other ligands on endothelium.
J. Immunol.
154
:
4099
-4112.
8
Luster, A. D., R. Alon, U. H. von Andrian.
2005
. Immune cell migration in inflammation: present and future therapeutic targets.
Nat. Immunol.
6
:
1182
-1190.
9
Issekutz, A. C., D. Rowter, T. A. Springer.
1999
. Role of ICAM-1 and ICAM-2 and alternate CD11/CD18 ligands in neutrophil transendothelial migration.
J. Leukocyte Biol.
65
:
117
-126.
10
Laudanna, C., J. Y. Kim, G. Constantin, E. C. Butcher.
2002
. Rapid leukocyte integrin activation by chemokines.
Immunol. Rev.
186
:
37
-46.
11
Harokopakis, E., G. Hajishengallis.
2005
. Integrin activation by bacterial fimbriae through a pathway involving CD14, Toll-like receptor 2, and phosphatidylinositol-3-kinase.
Eur. J. Immunol.
35
:
1201
-1210.
12
Sendide, K., N. E. Reiner, J. S. Lee, S. Bourgoin, A. Talal, Z. Hmama.
2005
. Cross-talk between CD14 and complement receptor 3 promotes phagocytosis of mycobacteria: regulation by phosphatidylinositol 3-kinase and cytohesin-1.
J. Immunol.
174
:
4210
-4219.
13
Simon, D. I., H. Xu, S. Ortlepp, C. Rogers, N. K. Rao.
1997
. 7E3 Monoclonal antibody directed against the platelet glycoprotein IIb/IIIa cross-reacts with the leukocyte integrin Mac-1 and blocks adhesion to fibrinogen and ICAM-1.
Arterioscler. Thromb. Vasc. Biol.
17
:
528
-535.
14
Libby, P..
2002
. Inflammation in atherosclerosis.
Nature
420
:
868
-874.
15
Nageh, M. F., E. T. Sandberg, K. R. Marotti, A. H. Lin, E. P. Melchior, D. C. Bullard, A. L. Beaudet.
1997
. Deficiency of inflammatory cell adhesion molecules protects against atherosclerosis in mice.
Arterioscler. Thromb. Vasc. Biol.
17
:
1517
-1520.
16
Collins, R. G., R. Velji, N. V. Guevara, M. J. Hicks, L. Chan, A. L. Beaudet.
2000
. P-selectin or intercellular adhesion molecule (ICAM)-1 deficiency substantially protects against atherosclerosis in apolipoprotein E-deficient mice.
J. Exp. Med.
191
:
189
-194.
17
Rosen, H., S. Gordon.
1990
. The role of the type 3 complement receptor in the induced recruitment of myelomonocytic cells to inflammatory sites in the mouse.
Am. J. Respir. Cell Mol. Biol.
3
:
3
-10.
18
Meurman, J. H., M. Sanz, S. J. Janket.
2004
. Oral health, atherosclerosis, and cardiovascular disease.
Crit. Rev. Oral Biol. Med.
15
:
403
-413.
19
Hajishengallis, G., A. Sharma, M. W. Russell, R. J. Genco.
2002
. Interactions of oral pathogens with Toll-like receptors: possible role in atherosclerosis.
Ann. Periodontol.
7
:
72
-78.
20
Beck, J. D., J. R. Elter, G. Heiss, D. Couper, S. M. Mauriello, S. Offenbacher.
2001
. Relationship of periodontal disease to carotid artery intima-media wall thickness: the Atherosclerosis Risk in Communities (ARIC) Study.
Arterioscler. Thromb. Vasc. Biol.
21
:
1816
-1822.
21
Wu, T., M. Trevisan, R. J. Genco, J. P. Dorn, K. L. Falkner, C. T. Sempos.
2000
. Periodontal disease and risk of cerebrovascular disease: the first national health and nutrition examination survey and its follow-up study.
Arch. Intern. Med.
160
:
2749
-2755.
22
Desvarieux, M., R. T. Demmer, T. Rundek, B. Boden-Albala, D. R. Jacobs, Jr, R. L. Sacco, P. N. Papapanou.
2005
. Periodontal microbiota and carotid intima-media thickness: the Oral Infections and Vascular Disease Epidemiology Study (INVEST).
Circulation
111
:
576
-582.
23
Zambon, J. J., S. Grossi, R. Dunford, V. I. Harazsthy, H. Preus, R. J. Genco.
1994
. Epidemiology of subgingival bacterial pathogens in periodontal diseases. R. J. Genco, Jr, and S. Hamada, Jr, and J. R. Lehrer, Jr, and J. R. McGhee, Jr, and S. Mergenhangen, Jr, eds.
Molecular Pathogenesis of Periodontal Disease
3
-12. American Society for Microbiology, Washington, D.C.
24
Haraszthy, V. I., J. J. Zambon, M. Trevisan, M. Zeid, R. J. Genco.
2000
. Identification of periodontal pathogens in atheromatous plaques.
J. Periodontol.
71
:
1554
-1560.
25
Malek, R., J. G. Fisher, A. Caleca, M. Stinson, C. J. van Oss, J. Y. Lee, M. I. Cho, R. J. Genco, R. T. Evans, D. W. Dyer.
1994
. Inactivation of Porphyromonas gingivalis fimA gene blocks periodontal damage in gnotobiotic rats.
J. Bacteriol.
176
:
1052
-1059.
26
Gibson, F. C., III, H. Yumoto, Y. Takahashi, H. H. Chou, C. A. Genco.
2006
. Innate immune signaling and Porphyromonas gingivalis-accelerated atherosclerosis.
J. Dent. Res.
85
:
106
-121.
27
Hajishengallis, G., P. Ratti, E. Harokopakis.
2005
. Peptide mapping of bacterial fimbrial epitopes interacting with pattern recognition receptors.
J. Biol. Chem.
280
:
38902
-38913.
28
Shimaoka, M., T. A. Springer.
2004
. Therapeutic antagonists and the conformational regulation of the β2 integrins.
Curr. Top. Med. Chem.
4
:
1485
-1495.
29
Walter, C., J. Zahlten, B. Schmeck, C. Schaudinn, S. Hippenstiel, E. Frisch, A. C. Hocke, N. Pischon, H. K. Kuramitsu, J.-P. Bernimoulin, N. Suttorp, M. Krüll.
2004
. Porphyromonas gingivalis strain-dependent activation of human endothelial cells.
Infect. Immun.
72
:
5910
-5918.
30
Amano, A., A. Sharma, J. Y. Lee, H. T. Sojar, P. A. Raj, R. J. Genco.
1996
. Structural domains of Porphyromonas gingivalis recombinant fimbrillin that mediate binding to salivary proline-rich protein and statherin.
Infect. Immun.
64
:
1631
-1637.
31
Pugin, J., V. V. Kravchenko, J. D. Lee, L. Kline, R. J. Ulevitch, P. S. Tobias.
1998
. Cell activation mediated by glycosylphosphatidylinositol-anchored or transmembrane forms of CD14.
Infect. Immun.
66
:
1174
-1180.
32
Hajishengallis, G., R. I. Tapping, M. H. Martin, H. Nawar, E. A. Lyle, M. W. Russell, T. D. Connell.
2005
. Toll-like receptor 2 mediates cellular activation by the B subunits of type II heat-labile enterotoxins.
Infect. Immun.
73
:
1343
-1349.
33
Worthylake, R. A., S. Lemoine, J. M. Watson, K. Burridge.
2001
. RhoA is required for monocyte tail retraction during transendothelial migration.
J. Cell Biol.
154
:
147
-160.
34
Diamond, M. S., T. A. Springer.
1993
. A subpopulation of Mac-1 (CD11b/CD18) molecules mediates neutrophil adhesion to ICAM-1 and fibrinogen.
J. Cell Biol.
120
:
545
-556.
35
Poggi, A., F. Spada, P. Costa, E. Tomasello, V. Revello, N. Pella, M. R. Zocchi, L. Moretta.
1996
. Dissection of lymphocyte function-associated antigen 1-dependent adhesion and signal transduction in human natural killer cells shown by the use of cholera or pertussis toxin.
Eur. J. Immunol.
26
:
967
-975.
36
Hmama, Z., K. L. Knutson, P. Herrera-Velit, D. Nandan, N. E. Reiner.
1999
. Monocyte adherence induced by lipopolysaccharide involves CD14, LFA-1, and cytohesin-1: regulation by Rho and phosphatidylinositol 3-kinase.
J. Biol. Chem.
274
:
1050
-1057.
37
Laudanna, C., J. J. Campbell, E. C. Butcher.
1996
. Role of Rho in chemoattractant-activated leukocyte adhesion through integrins.
Science
271
:
981
-983.
38
Hajishengallis, G., H. Nawar, R. I. Tapping, M. W. Russell, T. D. Connell.
2004
. The type II heat-labile enterotoxins LT-IIa and LT-IIb and their respective B pentamers differentially induce and regulate cytokine production in human monocytic cells.
Infect. Immun.
72
:
6351
-6358.
39
Aktories, K..
1997
. Bacterial toxins that target Rho proteins.
J. Clin. Invest.
99
:
827
-829.
40
Welch, H. C., W. J. Coadwell, L. R. Stephens, P. T. Hawkins.
2003
. Phosphoinositide 3-kinase-dependent activation of Rac.
FEBS Lett.
546
:
93
-97.
41
Khlgatian, M., H. Nassar, H. H. Chou, F. C. Gibson, III, C. A. Genco.
2002
. Fimbria-dependent activation of cell adhesion molecule expression in Porphyromonas gingivalis-infected endothelial cells.
Infect. Immun.
70
:
257
-267.
42
Abraham, R. T..
2003
. Rap1 redux.
Nat. Immunol.
4
:
725
-727.
43
Akira, S., K. Takeda.
2004
. Toll-like receptor signalling.
Nat. Rev. Immunol.
4
:
499
-511.
44
McDowall, A., D. Inwald, B. Leitinger, A. Jones, R. Liesner, N. Klein, N. Hogg.
2003
. A novel form of integrin dysfunction involving β1, β2, and β3 integrins.
J. Clin. Invest.
111
:
51
-60.
45
Pan, Z. K., L. Y. Chen, C. G. Cochrane, B. L. Zuraw.
2000
. fMet-Leu-Phe stimulates proinflammatory cytokine gene expression in human peripheral blood monocytes: the role of phosphatidylinositol 3-kinase.
J. Immunol.
164
:
404
-411.
46
Rawadi, G., J. L. Zugaza, B. Lemercier, J. C. Marvaud, M. Popoff, J. Bertoglio, S. Roman-Roman.
1999
. Involvement of small GTPases in Mycoplasma fermentans membrane lipoproteins-mediated activation of macrophages.
J. Biol. Chem.
274
:
30794
-30798.
47
Etienne-Manneville, S., A. Hall.
2002
. Rho GTPases in cell biology.
Nature
420
:
629
-635.
48
Bokoch, G. M., C. J. Vlahos, Y. Wang, U. G. Knaus, A. E. Traynor-Kaplan.
1996
. Rac GTPase interacts specifically with phosphatidylinositol 3-kinase.
Biochem. J.
315
: (Pt. 3):
775
-779.
49
Arbibe, L., J. P. Mira, N. Teusch, L. Kline, M. Guha, N. Mackman, P. J. Godowski, R. J. Ulevitch, U. G. Knaus.
2000
. Toll-like receptor 2-mediated NF-κB activation requires a Rac1-dependent pathway.
Nat. Immunol.
1
:
533
-540.
50
Lamont, R. J., H. F. Jenkinson.
1998
. Life below the gum line: pathogenic mechanisms of Porphyromonas gingivalis.
Microbiol. Mol. Biol. Rev.
62
:
1244
-1263.
51
Jain, A., E. L. Batista, Jr, C. Serhan, G. L. Stahl, T. E. Van Dyke.
2003
. Role for periodontitis in the progression of lipid deposition in an animal model.
Infect. Immun.
71
:
6012
-6018.
52
Li, L., E. Messas, E. L. J. Batista, R. A. Levine, S. Amar.
2002
. Porphyromonas gingivalis infection accelerates the progression of atherosclerosis in a heterozygous apolipoprotein E-deficient murine model.
Circulation
105
:
861
-867.
53
Watanabe, K., Y. Yamaji, T. Umemoto.
1992
. Correlation between cell-adherent activity and surface structure in Porphyromonas gingivalis.
Oral Microbiol. Immunol.
7
:
357
-363.
54
Neumann, F.-J..
2002
. Chlamydia pneumoniae-atherosclerosis link: a sound concept in search for clinical relevance.
Circulation
106
:
2414
-2416.
55
Belland, R. J., S. P. Ouellette, J. Gieffers, G. I. Byrne.
2004
. Chlamydia pneumoniae and atherosclerosis.
Cell. Microbiol.
6
:
117
-127.
56
Kalayoglu, M. V., B. N. Perkins, G. I. Byrne.
2001
. Chlamydia pneumoniae-infected monocytes exhibit increased adherence to human aortic endothelial cells.
Microbes Infect.
3
:
963
-969.
57
May, A. E., V. Redecke, S. Grüner, R. Schmidt, S. Massberg, T. Miethke, B. Ryba, C. Prazeres da Costa, A. Schömig, F.-J. Neumann.
2003
. Recruitment of Chlamydia pneumoniae-infected macrophages to the carotid artery wall in noninfected, nonatherosclerotic mice.
Arterioscler. Thromb. Vasc. Biol.
23
:
789
-794.
58
Krüll, M., A. C. Klucken, F. N. Wuppermann, O. Fuhrmann, C. Magerl, J. Seybold, S. Hippenstiel, J. H. Hegemann, C. A. Jantos, N. Suttorp.
1999
. Signal transduction pathways activated in endothelial cells following infection with Chlamydia pneumoniae.
J. Immunol.
162
:
4834
-4841.
59
Giacona, M. B., P. N. Papapanou, I. B. Lamster, L. L. Rong, V. D. D’Agati, A. M. Schmidt, E. Lalla.
2004
. Porphyromonas gingivalis induces its uptake by human macrophages and promotes foam cell formation in vitro.
FEMS Microbiol. Lett.
241
:
95
-101.