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The Journal of Immunology, 2008, 181, 2672 -2682
Copyright © 2008 by The American Association of Immunologists, Inc.
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Outer Membrane Protein A Expression in Escherichia coli K1 Is Required to Prevent the Maturation of Myeloid Dendritic Cells and the Induction of IL-10 and TGF-β1

Rahul Mittal* and Nemani V. Prasadarao2,*,{dagger}

* Division of Infectious Diseases, Saban Research Institute, Childrens Hospital Los Angeles, and {dagger} Keck School of Medicine, University of Southern California, Los Angeles, CA 90027


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Dendritic cells (DCs) are professional APCs that direct both cellular and humoral immune responses. Escherichia coli K1 causes meningitis in neonates; however, the interactions between this pathogen and DCs have not been previously explored. In the present study, we observed that E. coli K1, expressing outer membrane protein A (OmpA), was able to enter, survive, and replicate inside DCs, whereas OmpA– E. coli was killed within a short period. Opsonization of OmpA+ E. coli either with adult or cord serum did not affect its survival inside DCs. Exposure of DCs to live OmpA+ E. coli K1 prevented DCs from progressing in their maturation process as indicated by failure to up-regulate costimulatory molecules, CD40, HLA-DR, and CD86. The distinct DC phenotype requires direct contact between live bacteria and DCs. The expression of costimulatory molecules was suppressed even after pretreatment of DCs with LPS or peptidoglycan. Furthermore, the suppressive effects of OmpA+ E. coli on DCs were abrogated when the bacteria were incubated with anti-OmpA Ab. The inhibitory effect on DC maturation was associated with increased production of IL-10 as well as TGF-β and decreased production of IL-6, TNF-{alpha}, IL-1β, and IL-12p70 by DCs, a phenotype associated with tolerogenic DCs. These results suggest that the subversion of DC functions may be a novel strategy deployed by this pathogen to escape immune defense and persist in the infected host to reach a high degree of bacteremia, which is crucial for E. coli to cross the blood-brain barrier.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Bacterial meningitis is the most serious and frequently fatal infection of the CNS. It results in significant neurological sequelae in half of the survivors, including hearing loss, convulsive disorders, abnormal speech patterns, cortical blindness, and mental retardation (1). The incidence of bacterial meningitis is ~5 cases per 100,000 adults per year in developed countries and may be 10 times higher in underdeveloped countries (2, 3). Escherichia coli K1 is a predominant pathogen in neonatal meningitis and septicemia (4, 5). E. coli K1 meningitis, in particular, is associated with high mortality rates and poor outcome. Studies have shown that a high degree of bacteremia is required for E. coli to cross the blood-brain barrier, indicating that the bacterium must evade host defense mechanisms and survive in the blood stream (6, 7). Our studies in the newborn rat model of hematogenous meningitis have demonstrated that E. coli enters monocytes and macrophages and then multiplies in a time-dependent fashion (8). Therefore, the ability of E. coli to survive in host immune cells like monocytes, macrophages, and neutrophils may represent an important step in the vicious cycle of this organism. Survival of E. coli within monocytes may be crucial in the establishment of a replication permissive niche. In this regard, outer membrane protein A (OmpA)3 of E. coli has been demonstrated to play an important role in infectious process by interacting with immune cells (8). Several virulence factors of E. coli have been implicated in facilitating the onset of meningitis in humans. These include OmpA, fimbriae, IbeA, IbeB, cytotoxic necrotizing factor (CNF), and TraJ (9, 10, 11, 12, 13). However, lack of OmpA expression in E. coli profoundly prevents the incidence of meningitis in infant rats when compared with other virulence factors (8, 11). We have previously shown that OmpA of E. coli binds C4b-binding protein (C4bp), an inhibitor of complement activation, and that log phase OmpA+ E. coli avoids serum bactericidal activity more effectively than OmpA– E. coli, which failed to survive in serum (14). In addition, E. coli pretreated with either adult human serum or purified C4bp could not survive in macrophages efficiently when compared with the bacteria treated with neonatal serum, indicating that serum complement factors may contribute to the interaction between E. coli and phagocytic cells (15). The receptor analogs that bind OmpA also inhibited the occurrence of meningitis in newborn rats, clearly demonstrating that OmpA is a major factor that contributes to the pathogenesis (16).

The mononuclear phagocyte system is considered to be a continuum linking circulating pluripotent monocytes with differentiated effector cells such as tissue-based macrophages or specialized APCs. Dendritic cells (DCs) are the most potent APCs that play a crucial role in initiation and modulation of specific immune responses (17, 18). DCs are rare sentinel cells that provide an important line of defense and are key orchestrators of the immune responses against invading pathogens (19, 20). Immature DCs, in the periphery and submucosa, sample the external environment and capture Ag including whole bacteria. After successful sampling, DCs migrate to secondary lymphoid tissue where they present processed Ag to stimulate Ag-specific T cells (21, 22, 23). DC activation induces up-regulation of costimulatory molecules and abundant surface expression of MHC class II molecules resulting in so-called mature DCs, which are potent stimulators of naive T cells (24, 25, 26). The outcome of the interaction between DCs and lymphocytes is critically influenced by the release of both cytokines and chemokines by DCs and by the expression of critical costimulatory molecules (27, 28, 29). There is a paucity of literature regarding the interaction of E. coli with human DCs. Since DCs occupy center stage as the most efficient APC in the immune system, there is a clear need to understand how human DCs respond to E. coli. Herein, we demonstrate for the first time that OmpA+ E. coli prevents the maturation of DCs to promote its multiplication inside DCs.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Bacterial strains

All strains used in this study were derived from a cerebrospinal fluid isolate of E. coli K1 strain RS 218 (serotype O18:K1:H7). E44 is a spontaneous rifampin-resistant mutant of E. coli K1 strain RS 218 (designated as OmpA+ E. coli) and invades human brain microvascular endothelial cells in a cell culture model (9, 30). E91 is a noninvasive derivative of E44 that expresses no OmpA, since the ompA gene is disrupted. E91 (OmpA– E. coli) was transformed with pUC19 containing the entire ompA gene and the pUC19 plasmid alone to obtain pOmpA+ E. coli and pOmpA– E. coli, respectively (9). E. coli K1 strains that lack S-fimbriae ({Delta}sfa), type 1 fimbriae ({Delta}fimH), ibeA gene ({Delta}ibeA), and cytotoxic necrotizing factor 1 ({Delta}cnf1) are described elsewhere (11, 12, 13). E. coli{Delta}sfa and E. coli{Delta}fimH are invasive in human brain microvascular endothelial cells, whereas E. coli{Delta}ibeA and E. coli{Delta}CNF1 are noninvasive. All bacteria were grown in brain-heart infusion broth with appropriate antibiotics as necessary. All bacterial media were purchased from Difco Laboratories. For studies using chloramphenicol, optimization of time kill curves was performed in our bacterial culture system. At doses of 3–10 µg ml–1 and at concentrations over 105–107 CFU ml–1, the bacterial concentration remained static over an incubation period of up to 24 h, as determined by serial dilution, plating on blood agar, and enumerating the bacteria after overnight incubation. For preparation of bacterial lysates, E. coli was suspended in 0.5% Triton X-100 in PBS and sonicated briefly (30-s pulses) over a period of 3 min on ice. Cellular debris were removed by centrifugation followed by filtration through 0.2-µm pore size filters. Lysis of live bacteria was confirmed by plating the lysates on blood agar plates and incubation overnight at 37°C. All general chemicals were obtained from Sigma-Aldrich unless otherwise stated.

DC culture and activation

DCs were generated from human PBMCs as described previously (31, 32) and obtained from AllCells, LLC. Briefly, monocytes were prepared from PBMCs by positive selection using CD14 immunomagnetic beads (Miltenyi Biotec). CD14+ isolated cells were then cultured in RPMI 1640 supplemented with 10% FCS, 2.4 mM L-glutamine (Invitrogen), 50 ng ml–1 human recombinant GM-CSF, and 20 ng ml–1 human rIL-4 (PeproTech). DCs were used after 7 days of culture and phenotype was determined by FACSCalibur flow cytometer (BD Biosciences). Immature DCs were CD3neg, CD14low, CD19neg, CD83neg, and CD25neg and expressed low levels of HLA-DR, CD40, CD86, and CD1a. For stimulation experiments, DCs (5 x 104 ml–1) were cultured with live or killed E. coli at a multiplicity of infection of 10 (cell:bacteria ratio, 1:10) for 24 and 48 h. In some experiments, 3–10 µg ml–1 chloramphenicol was added to the DC bacteria coculture either at the start of stimulation or at various time points after addition of bacteria. DCs were also stimulated with LPS (Sigma-Aldrich) or peptidoglycan (PGN; Calbiochem) at a concentration of 10 ng ml–1 as well as with a maturation mixture (MM) containing TNF-{alpha} (10 ng ml–1), IL-1β (10 ng ml–1), and PGE2 (1 µg ml–1) (33). In separate experiments, DCs were incubated with supernatants from DC and E. coli K1 cocultured for 24 h. To study the effect of serum and complement components, bacteria were incubated with adult or cord serum (40–100%), C4bp (10 µg ml–1). and factor H (10 µg ml–1) for 15 min, then washed three times and used in phagocytosis assay as well as for stimulating DCs. Before Ab staining, an aliquot of DC culture was stained with trypan blue, propidium iodide (PI), or annexin-V to assess the amount of cell death in the coculture. For Transwell experiments, DCs were added to 24-well tissue culture plates at a density of 5 x 104 cells ml–1. Transwell inserts (0.1-µm pore size, Falcon; BD Biosciences) were placed in contact with medium and the bacteria were added to the top chamber. After overnight culture, DCs were removed and cell surface markers were analyzed by flow cytometry. Lack of any transmigration of live bacteria to the lower chamber was proven by culturing an aliquot of the stimulated cells on blood agar plates.

Phagocytosis and detection of intracellular and extracellular bacterial growth

DCs (5 x 104 cells ml–1) were washed three times in culture medium without antibiotics and then placed in 500 µl of culture medium in 12 x 75-mm polystyrene snap-cap tubes (Falcon; BD Bisociences). Varying concentrations of bacteria (in 10 µl) were added to the tubes. DC and bacteria were then incubated for 1 h at 37°C. Gentamicin was added to DC-bacteria coculture tubes at a final concentration of 100 µg ml–1 and incubated for an additional 60 min at 37°C. DC-bacteria cocultures were washed three times in RPMI 1640 containing no antibiotics and reconstituted with antibiotic-free culture medium. Tubes were then assessed immediately for intracellular bacteria (time period 0) or placed again at 37°C in culture medium containing 30 µg ml–1 gentamicin. To assess intracellular bacteria at different times postexposure, DC-bacteria cocultures (described above) were washed twice and cells were lysed with 100 µl of 0.5% Triton X-100. Percentage uptake of bacteria was assessed by plating 20 µl each of DC-bacteria coculture supernatant onto blood agar plates. Count in supernatant was subtracted from the initial count to calculate the uptake of bacteria by DCs. Results are expressed as percentage of viable bacteria taken up by the DCs at respective sampling time intervals.

Surface marker expression by flow cytometry

Expression of surface molecules associated with DC maturation and activation (CD40, CD86, and HLA-DR) was detected by staining with appropriate FITC, PE, PE-Cy5.5, or allophycocyanin mouse mAbs and mouse IgG isotype-matched controls (eBioscience). Cells were first preincubated for 20 min with IgG-blocking buffer to mask nonspecific binding sites and then further incubated with the indicated Abs or an isotype-matched control Ab for 30 min at 4°C. After incubation, the cells were washed three times with PBS containing 2% FBS and subsequently fixed with BD Cytofix (BD Biosciences). Cells were then analyzed by four-color flow cytometry using FACSCalibur and CellQuest Pro software (BD Biosciences). DCs form a distinct population when separated by side and forward scatter parameters for which CD1a was used as a DC gating marker; this population formed the collection gate and at least 5000 events within this gate were collected for analysis. The data are presented as geometric mean fluorescence intensity of logarithmic data after subtracting the values obtained with isotype-matched control Abs.

Fluorescence microscopy

DCs were infected with either OmpA+ or OmpA– E. coli for varying periods, washed with PBS, and preincubated with IgG-blocking buffer to prevent nonspecific binding. Cells were then fixed with BD Cytofix (BD Biosciences), washed, and incubated with 5 µg ml–1 anti-HLA-DR Ab (eBioscience) followed by Alexa Fluor 488 goat anti-mouse IgG (Invitrogen). Cells were then allowed to adhere to poly-L-lysine-coated slides for 10 min (Paul Marienfeld) and mounted in an antifade Vectashield solution containing 4',6-diamidino-2-phenylindole (Vector Laboratories) The cells were viewed with a Leica DMRA microscope with Plan-apochromat oil immersion objective lenses. Images were acquired with a SkyVision-2/VDS digital charge-coupled device camera (12-bit; 1280 x 1024 pixels) in unbinned or 2 x 2 binned models into the EasyFISH software, saved as 16-bit monochrome images, and merged as 24-bit RGB TIFF images (Applied Spectral Imaging). The images were then assembled using Adobe Photoshop 7.0.

Viability

The percentage of viable DCs was assessed by trypan blue, PI staining, and an annexin V-FITC apoptosis kit (BD Biosciences). In all culture conditions, a proportion of cells (ranging from 5 to 15%) were trypan blue, PI, and/or annexin V positive. However, there was no significant difference observed in the proportion in cultures stimulated with medium, live, LPS, PGN, or killed E. coli.

Cytokine measurements

Cytokine (TNF-{alpha}, IL-1β, IL-6, IL-12p70, IL-10, and TGF-β) production in cell culture supernatants of DC-bacteria coculture experiments collected after 24 and 48 h of incubation was conducted using Biosource ELISA kits (Invitrogen) according to the manufacturer’s instructions.

Statistical analysis

Statistical significance was determined by a paired, two-tailed Student’s t test. Values of p < 0.05 were considered to be statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
E. coli K1 enters and survives in myeloid DCs for which OmpA expression is critical and preincubation with serum or C4b-binding protein does not affect the interaction

To examine the entry and survival of E. coli K1 in DCs derived from myeloid cells, OmpA+ or OmpA– E. coli were incubated with DCs at a multiplicity of infection (moi) of 10 (DC:bacteria ratio, 1:10) or 100. In addition, we investigated the role of other important factors required to invade brain microvascular endothelial cell by using E. coli K1 strains lacking S-fimbriae (sfa), type 1 fimbriae (fimH), CNF, and IbeA. As shown in Fig. 1A, 100% of all strains entered DCs at a moi of 10 and no bacteria were left in the solution at 2 h postinfection. However, 60% of bacteria entered the cells at a moi of 100 after 2 h of infection, which was increased to 100% by 6 h (Fig. 1B). Of note, 100% of OmpA– E. coli was taken up by DCs within 2 h postinfection at a moi of 100. The entry of other mutant strains of E. coli was similar to OmpA+ E. coli, suggesting that lack of any of the virulence factors does not prevent the uptake of bacteria by DCs. Faster uptake of OmpA– E. coli could be due to efficient killing of the bacteria by DCs within a short period of time. In case of OmpA+ E. coli, the entry might be mediated through a receptor on DCs, whose recycling is necessary for subsequent uptake of the bacteria at higher inoculum sizes. In agreement with this concept, the intracellular survival of E. coli inside DCs as measured by gentamicin protection assays demonstrated that OmpA– E. coli were killed within 2 h postinfection at both inoculum sizes, whereas OmpA+ E. coli as well as {Delta}cnf, {Delta}fimH, {Delta}ibeA, and {Delta}sfa strains enter and survive inside DCs equally well (Fig. 1, C and D). Viable counts of OmpA+ E. coli increased with an increase in postinfection time from 2 to 6 h when used at a moi of 10, indicating the multiplication of bacteria inside DCs. The number of OmpA+ E. coli that survived inside DCs was greater when the assays were conducted at a moi of 100 (Fig. 1D). These data suggest that the expression of OmpA, but not other virulence factors significantly contribute to the survival of E. coli inside DCs. To further confirm the role of OmpA in survival of bacteria inside DCs, OmpA– E. coli were transfected with a plasmid containing an entire ompA gene (pOmpA+ E. coli) or plasmid alone (pOmpA– E. coli). The pOmpA+ E. coli expressed equal amounts OmpA on its surface as determined by Western blot analysis (9). Similar to that of OmpA+ E. coli, the OmpA complemented strain pOmpA+ E. coli survived inside DCs, whereas plasmid alone transformed pOmpA– E. coli were killed within 2 h postinfection (Fig. 1, A–D). Taken together, these data suggest that OmpA expression is critical to the survival of E. coli inside DCs. All subsequent experiments were performed with bacterial strains at a moi of 10.


Figure 1
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  		FIGURE 1. Phagocytosis of E. coli by DCs. Various strains of E. coli were incubated with DCs at a moi of 10 (A and C) or 100 (B and D) for the indicated time points. The uptake of bacteria (A and B) or intracellular bacteria (C and D) was determined as described in Materials and Methods. In separate experiments, OmpA+ E. coli were untreated or treated with heat-inactivated serum (HIS), adult serum (AS), cord serum (CS), C4bp, or factor H (FH), washed, and used for phagocytosis assays. Percentage uptake (E) and intracellular survival (F) of OmpA+ E. coli in DCs at moi of 10 was determined. Data are presented as log10 CFU per 5 x 104 DCs. The data represent mean ± SD of triplicate samples from three independent experiments.

 
Our previous studies demonstrated that OmpA+ E. coli incubated with adult human serum could not efficiently enter macrophages and human brain microvascular endothelial cells when compared with bacteria treated with newborn serum (15). The serum component that prevented the interaction of E. coli with host cells has been identified as C4bp, a classical complement pathway regulator, which also helps evade serum bactericidal activity (34). These studies indicate that the deposition of serum components contribute to the interaction between E. coli and host cells. To examine whether the differential deposition of various complement components affect the survival of OmpA+ E. coli with DCs, the bacteria were treated with adult or cord serum (40–100%) for 15 min, washed, and then added to DCs. In addition, OmpA+ E. coli were also treated with heat-inactivated serum as a control. Neither adult or cord serum affected the entry nor the survival of OmpA+ E. coli inside DCs (Fig. 1, E and F). Bacteria treated with either C4bp or factor H (an alternative pathway regulator) also entered and survived similar to that of untreated OmpA+ E. coli, suggesting that complement deposition is not playing a significant role in the interaction of OmpA+ E. coli and DCs.

Immature DCs are optimally phagocytic and pinocytose Ags; however, DCs must be activated to mature before serving as efficient APCs. The HLA-DR molecules are critical for the initiation of the Ag-specific immune response, because they guide the development and activation of CD4+ Th cells. Therefore, to examine whether DCs were activated upon infection with OmpA+ E. coli and the morphology of internalized E. coli after incubation for 6 and 24 h, immunocytochemistry was performed. DCs were stained with PE-conjugated HLA-DR Ab and the intracellular bacteria were stained with anti-S-fimbria Ab or anti-OmpA Ab followed by Alexa Fluor 488-conjugated secondary Ab. The internalized OmpA+ E. coli, which were able to survive and multiply within DCs at 6 h (Fig. 2), still maintained the rod shape structure at 24 h, indicating that pathogens can exploit host cells as replication-permissive niches. The majority of OmpA+ E. coli were localized in cytoplasm of DCs, indicating cytoplasm as a preferential site of replication. In addition, DCs infected with OmpA+ E. coli for 6 h revealed the expression of HLA-DR on its surface, which appeared to be clustered in certain areas and also appeared to be colocalized at the bacterial staining. The surface expression of HLA-DR was completely suppressed by 24 h postinfection with OmpA+ E. coli. In contrast, OmpA– E. coli stained with anti-S-fimbria Ab showed no bacteria inside DCs (Fig. 2H), suggesting that OmpA expression in E. coli is important for the survival of the bacterium inside DCs. However, the OmpA– E. coli-infected DCs showed increased intensity of HLA-DR staining, although the cells showed clustered staining in several areas.


Figure 2
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  		FIGURE 2. Fluorescence microscopy of DCs infected with E. coli. DCs were cocultured with OmpA+ (A–F) or OmpA– E. coli (G–I) at a moi of 10. At 6 (A–C) and 24 h (D–I) postinfection, the cells were washed, DCs were allowed to adhere to poly-L-lysine slides, and then stained with either anti-S-fimbria Ab (B, E, and H) or anti-HLA-DR Ab (C, F, and I) followed by respective secondary Abs coupled to fluorophores. The slides were counterstained with 4',6-diamidino-2-phenylindole (DAPI) and visualized by fluorescence microscopy.

 
OmpA+ E. coli K1 suppresses the maturation of DCs

Since OmpA+ E. coli survived efficiently inside DCs when compared with OmpA– E. coli and suppressed the expression of HLA-DR, maturation of DCs was also characterized by analyzing the cell surface expression of CD40, CD86, and HLA-DR after 24- and 48-h stimulation by flow cytometry. LPS and PGN were used as positive controls in these studies. The surface expression of all of these molecules was significantly greater in OmpA– E. coli-infected DCs when compared with OmpA+ E. coli even after 48 h postinfection (Fig. 3, A–C; p < 0.01 by Student’s t test). LPS treatment of DCs alone stimulated the expression of CD40, CD86, and HLA-DR similar to that of OmpA– E. coli, whereas the levels stimulated by PGN were significantly higher than the levels induced by OmpA– E. coli. Some enhancement in the expression of cell surface costimulatory molecules was also detected in unstimulated medium-treated cells, which is likely a result of DC handling procedures and/or IL-4 and GM-CSF used in DC culture. Manipulation of DCs during tissue culture could alter E-cadherin-mediated DC-DC adhesion triggered by activation of the β-catenin pathway (35). However, OmpA+ E. coli-induced expression was relatively low compared with even unstimulated cells.


Figure 3
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  		FIGURE 3. Expression of maturation markers on the surface of DCs infected with E. coli. OmpA+ (E44), pOmpA+, pOmpA–, {Delta}ibeA, {Delta}sfa, {Delta}fimH, {Delta}CNF1, or OmpA– E. coli (E91), LPS, or PGN were incubated with DCs for 24 and 48 h. In separate experiments, anti-OmpA Abs were preincubated with OmpA+ E. coli (OmpA Ab-pretreated E44) or added to DC culture before coculture (E44 + OmpA Ab). Bacteria were also treated with control IgG and then incubated with DCs (E44 + control IgG). DCs were then washed, stained with Abs to CD40 (A and D), CD86 (B and E), and HLA-DR (C and F), fixed, and analyzed by flow cytometry. The data are presented as geometric mean fluorescence intensity (MFI) of logarithmic data subtracted from isotype-matched controls. The error bars represent SDs from the means of triplicate samples. The results are representative of three independent experiments.

 
In addition, DCs were also infected with E. coli{Delta}ibeA, E. coli{Delta}cnf, E. coli{Delta}sfa, and E. coli{Delta}fimH mutant strains, which did not induce up-regulation of maturation marker expression again substantiating the role of OmpA in the suppression of DC maturation (Fig. 3. D–F). To further confirm the role of OmpA, anti-OmpA Abs were preincubated with OmpA+ E. coli before stimulating DCs. The surface expression of costimulatory molecules significantly increased similar to the levels of OmpA– E. coli-induced markers (p < 0.01). The same stimulation effect was also observed when anti-OmpA Ab was added in the DC culturing medium and then challenged with OmpA+ E. coli (E44 plus OmpA Ab in Fig. 3, D–F). Treatment of the bacteria with a control Ab (E44 plus control IgG) failed to show any effect. In support of the OmpA role in the suppression of costimulatory molecules, OmpA– E. coli that was complemented with pUC19 containing the ompA gene (pOmpA+ E. coli) failed to up-regulate the expression of cell surface markers, whereas pOmpA– E. coli induced the expression of HLA-DR, CD40, and CD 86. Since the observed differences could be related to the excessive necrosis or apoptosis of DCs in coculture with proliferating bacteria, we investigated DC viability. Despite the exposure of DCs to continually proliferating bacteria, there was no difference in the proportion of dead DCs (ranging from 5 to 15%) for all culture conditions as determined by trypan blue staining (data not shown). To address the proportion of DCs that were undergoing apoptosis, uninfected and infected DCs were stained with annexin V or PI. No significant difference between the number of DCs that was apoptotic or necrotic when infected with live E. coli and the number of uninfected controls was observed (data not shown). Therefore, the distinct DC surface phenotype observed is not due to excessive cellular necrosis or apoptosis in live cocultures.

LPS, PGN, or maturation mixture-induced up-regulation of DC costimulatory markers was blocked by infection with OmpA+ E. coli

The lack of significant induction of surface markers on DC by live OmpA+ E. coli raised the question of whether the bacteria could inhibit activation by DC agonists. Previous studies demonstrated that the MM containing IL-1β and TNF-{alpha} as well as LPS and PGN induce the maturation of DCs (33, 36). Therefore, to examine whether these molecules exhibit any effect on OmpA+ E. coli-induced suppression of DC maturation, DCs were pretreated with LPS, PGN, or MM for 24 h followed by infection with OmpA+ E. coli for another 24 h. DCs stimulated with LPS or PGN failed to show expression of cell surface markers in the presence of OmpA+ E. coli (Fig. 4, A–C). Similarly, the surface expression of CD40, CD86, and HLA-DR molecules on the surface of DCs was also suppressed by infecting the cells with OmpA+ E. coli before treatment with the MM. These results demonstrate that OmpA expression in E. coli K1 contributed to the suppression of DC maturation even after challenging with LPS, PGN, or MM. Several pathogens have shown to secrete proteins into host cells via the type III secretion system to control the signaling mechanisms of the cells and thereby the innate immune response (37). Recently, uropathogenic E. coli has been shown to secrete a protein, TcpC, after interaction with macrophages, which binds to MyD88 protein to inhibit TLR-mediated signaling (38). However, there is no type III secretion system identified in E. coli K1 so far. To examine whether the inhibition of DC maturation by E. coli K1 is due to any secreted protein, the supernatants of OmpA+ E. coli previously incubated with DCs for 6 and 24 h were added to a fresh batch of DCs. The supernatants from both treatments also showed an inhibitory effect on DC maturation markers similar to that of OmpA+ E. coli K1 (Fig. 4, A-C, data from 6 h is only shown). These results suggest that a bacterial protein or other factors that are being secreted, after interaction of OmpA with its cognate receptor on DCs, might be controlling the signaling to prevent the maturation of DCs.


Figure 4
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  		FIGURE 4. Analysis of DC maturation following pre- or posttreatment with LPS, PGN, or MM (MC). DCs were either first stimulated with LPS, PGN, or MM for 24 h and then infected with OmpA+ E. coli K1 for an additional 24 h or vice versa. In some experiments, DCs were first incubated with E44 for 24 h, supernatants were collected, and added to fresh DCs for additional 24 h (Sup. + DCs). DCs from all of these treatments were then washed separately, stained with Abs to CD40 (A), HLA-DR (B), and CD86 (C), fixed, and analyzed by flow cytometry. The error bars represent SDs from the means of triplicate samples. The results are representative of three independent experiments.

 
Distinct DC surface phenotype induced by OmpA+ E. coli K1 requires bacterial protein synthesis and dividing bacteria

Since our experiments demonstrated that the supernatant obtained from the coculture of OmpA+ E. coli and DCs block the maturation of DCs, we speculate that bacterial proteins that were being released into the system might be responsible for the suppression of DC maturation. Therefore, to examine whether bacterial protein synthesis is required for the distinct DC phenotype observed when cultured with live bacteria, experiments were conducted using OmpA+ E. coli treated with bacteriostatic doses of chloroamphenicol (Cm; 10 µg ml–1) before and during incubation with DCs. The dose of Cm used was shown to inhibit the multiplication of E. coli by inhibiting the microbial protein synthesis and impairing peptidyl transferase activity. However, Cm does not alter the viability of the bacteria. Significant enhancement in the expression of CD40, CD86, and HLA-DR was observed in DCs cocultured with Cm-treated OmpA+ E. coli (Fig. 5, A–C) (p < 0.01). Cm had no effect on DC maturation when added alone, nor did it alter the induction of maturation markers in response to either killed bacteria, LPS, or PGN (data not shown). On the other hand, a delay in addition of Cm by 6 h after coculturing of E. coli and DCs lead to suppression of DC surface phenotype as observed with live bacteria. We also observed an increase in up-regulation of these markers when tetracycline, a cell-permeable antibiotic, was added to DCs after 2 h of bacterial coculture, thus preventing the multiplication of internal bacteria (Fig. 5, A–C). These findings suggest that poor maturation of DCs requires brief exposure to actively dividing OmpA+ E. coli and protein synthesis by the bacteria. It is possible that OmpA expressed in E. coli interacts with a DC receptor to induce virulence genes or factors that suppress the maturation of DCs.


Figure 5
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  		FIGURE 5. Bacteriostatic doses of Cm abrogate OmpA+ E. coli K1 induced a suppressive effect on maturation of DCs. OmpA+ E. coli K1 (moi of 10) pretreated with Cm (10 µg ml–1) were added to DCs for 24 and 48 h as described in Materials and Methods. In some experiments, Cm was added to DC-bacteria coculture immediately (E44 + Cm + 0 h) or after 6 h (E44 +Cm + 6 h). Similarly, tetracycline (5 µg ml–1) was added after a 2-h incubation of DCs with OmpA+ E. coli (E44 + Tet-2 h). The expression of CD40 (A), HLA-DR (B), and CD86 (C) was determined by flow cytometry. Data are expressed as the mean ± SD of triplicate samples and are representative of three independent experiments with similar results.

 
OmpA+ E. coli K1 induced suppression of DC maturation phenotype precludes stimulus of killed bacteria and requires direct contact between bacteria and DCs

Immune responses to live and killed bacteria will be different. During infection there is often a combination of both viable and dead bacteria; thus, we investigated the effect of killed bacteria alone or in the presence of live E. coli on the surface phenotype of DCs. As shown in Fig. 6, A–C, DCs when cocultured with killed E. coli exhibited the surface phenotype similar to that of DCs infected with OmpA– E. coli. In the presence of live OmpA+ E. coli, killed bacteria (ratio 10:100) were unable to induce DC markers CD40, CD86, and HLA-DR to levels observed when incubated with killed bacteria alone. Similar results were also observed when the ratio of live:killed bacteria was varied (1:100). These data show that live OmpA+ E. coli have primacy to take over the control of normally potent inducers of DC maturation. We next examined whether development of the distinct DC surface phenotype resulting from direct physical contact between DCs and bacteria. To explore this possibility, we used a Transwell system in which DCs were physically separated from bacteria by 0.1-µm porous membranes. Up-regulation of HLA-DR, CD86, and CD40 was observed following separation of bacteria from DCs after 24 h (Fig. 6, A–C). These results indicate that bacterial components released into the medium are capable of inducing DC maturation. To confirm these findings, DCs were incubated with supernatants from live bacteria grown to mid-log phase. We found that bacterial culture supernatant from live OmpA+ E. coli was a potent stimulator of DC maturation. Furthermore, DCs were also cocultured with lysates obtained after sonication and centrifugation of E. coli, which also induced up-regulation of DC maturation markers. These experiments demonstrate that inhibition of DC maturation requires close contact between live bacteria and the cells.


Figure 6
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  		FIGURE 6. OmpA+ E. coli K1-induced suppression of DC maturation phenotype dominates the stimulus of killed bacteria and requires direct contact between bacteria and DCs. Live or killed OmpA+ E. coli K1 at a moi of 10 were cocultured with DCs for 24 and 48 h. In separate experiments, DCs were cultured in the lower chamber of a Transwell unit and the bacteria (moi of 10) were added to the upper chamber. This was compared with DC responses in the absence of Transwell, but otherwise identical culture conditions. In parallel with this experiment, DCs were stimulated with the supernatant equivalent to that released from moi of 10 of live OmpA+ E. coli (supernatant). In addition, DCs were also stimulated with lysates obtained by sonicating live OmpA+ E. coli equivalent to moi of 10 (lysate). DCs were analyzed for expression of CD40 (A), CD86 (B), and HLA-DR (C) by flow cytometry. The data represent mean ± SD of three independent experiments performed in triplicate.

 
Infection of DCs with OmpA+ E. coli K1 prevents the production of proinflammatory cytokines but induces the production of IL-10 and TGF-β

DCs regulate immune responses that are crucial for microbial eradication through the production of stimulatory and suppressive cytokines. Th1 cytokines are crucial for host antimicrobial immune responses, while Th2 cytokines may inhibit the development of these effector mechanisms. IL-12 produced by DCs can bias for the development of Th1 responses. IL-10 and TGF-β production by DCs in contrast is associated with tolerogenic DCs. Therefore, we examine the cytokine profile of DCs following interaction with E. coli. DCs stimulated with OmpA+ E. coli as well as with mutant strains that express normal levels of OmpA did not produce proinflammatory cytokines TNF-{alpha}, IL-1β, IL-6, and IL-12p70, but showed higher production of IL-10 and TGF-β (Fig. 7, A–F). On the contrary, OmpA– and pOmpA– E. coli-, LPS-, or PGN-stimulated DCs produced higher levels of proinflammatory cytokines and lower or no production of IL-10 and TGF-β, indicating activation and maturation of DCs. Similar to the DC maturation marker experiments, infection with OmpA+ E. coli after treatment with either LPS or PGN significantly prevented the production of proinflammatory cytokines and stimulated higher levels of IL-10 and TGF-β. As shown in DC maturation marker experiments, anti-OmpA Abs either pretreated or added along with OmpA+ E. coli during the coculture with DCs significantly enhanced the production of proinflammatory cytokines TNF-{alpha}, IL-1β, IL-6, and IL-12, but not IL-10 and TGF-β. Next, we examined the effect of opsonization on the expression of costimulatory molecules and cytokine production in DCs infected with OmpA+ E. coli. The analyses also revealed that opsonized OmpA+ E. coli did not significantly affect the cytokine production. Similarly, OmpA+ E. coli pretreated with either C4bp or factor H had no effect on the suppression of proinflammatory or anti-inflammatory cytokines induced by the bacteria (data not shown). These results suggest that OmpA+ E. coli induces anti-inflammatory cytokine production in DCs and the deposition of complement factors does not alter the effect of the bacteria on DCs.


Figure 7
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  		FIGURE 7. Cytokine production by DCs infected with various strains of E. coli. DCs were exposed to medium alone, treated with control (E44 + control IgG), anti-OmpA Abs, and OmpA+ E. coli together (E44 + OmpA Ab), OmpA+ E. coli pretreated with anti-OmpA Abs (OmpA Ab-Pre E44), or various E. coli strains for 24 and 48 h. Culture supernatants were collected and the levels of TNF-{alpha} (A), IL-6 (B), IL-10 (C), IL-1β (D), IL-12p70 (E), and TGF-β (F) were assessed by ELISA. In addition, cytokine assays were also performed of the supernatants of DCs treated with LPS or PGN. The error bars represent SDs from the means of triplicate samples from four individual experiments.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
DCs are highly specialized APCs that form a gateway between the innate and adaptive immune systems. It has been shown that DCs are crucial in the capture of bacteria in vivo, and it is believed that this function is critical for initiating adaptive immunity against bacterial Ags (39, 40). A role for DCs in antipathogen immunity is supported by the observation that deletion of DCs from mice renders them susceptible to infection by pathogens such as rodent malaria (41). However, pathogens hijack normal host protective immune responses to subvert immune clearance. Deregulation or premature termination of a clearing immune response can lead to long-term pathogen persistence. In the present study, we examined the ability of monocyte-derived myeloid DCs to ingest and kill live E. coli. We observed that 100% of OmpA+ E. coli was ingested by DCs after 2 h at a moi of 10. However, DCs failed to kill ingested bacteria, which then multiply and increase their number within these cells. Of note, our data indicate that OmpA expression in E. coli is critical for survival since OmpA– E. coli is unable to survive even for a short period of time. Serum opsonization also failed to affect the survival of OmpA+ E. coli within DCs. Immunocytochemistry revealed a large number of OmpA+ E. coli distributed throughout DCs after 24 h postinfection without killing the cell, suggesting that the bacteria might be using cytoplasm as a highly permissive environment. This may represent an important step for OmpA+ E. coli during the initial stages of infection to multiplication and allow them to produce high degree of bacteremia. However, this phenomenon is in contrast to the survival of OmpA+ E. coli in macrophages, in which the bacterial multiplies in a phagosome very rapidly and burst opens the cell (8). Billard et al. (42) demonstrated that DCs can serve as a reservoir for Brucella and may play an important role in dissemination of this pathogen into host tissues. Banks et al. (33) also observed that although Haemophilus ducreyi is ingested by DCs, killing is incomplete, as 5% of the internalized H. ducreyi (vs 95% of E. coli) were still recoverable from DCs for up to 48 h, which corroborates with our findings in this study (33).

In the present investigation, infection of DCs with live OmpA+ E. coli failed to up-regulate the expression of costimulatory molecules CD40, HLA-DR, and CD86. By affecting DC maturation, OmpA+ E. coli could alter DC functions, leading to increased bacterial persistence within the host. We also tested the ability of other E. coli mutants that were defective in the expression of various virulence factors, including FimH (a subunit of type I pili), CNF, S-fimbrial adhesion (sfa), and ibeA to stimulate DCs. All of these mutants showed a similar phenotype to the wild-type bacteria in survival and suppressing the up-regulation of costimulatory molecules in DCs. Interestingly, OmpA+ E. coli also inhibited DC maturation induced by potent agonists LPS, PGN, and DC MM. On the contrary, OmpA– E. coli activated myeloid DCs as indicated by enhanced expression of costimulatory molecules on the surface of DCs. Agrawal et al. (43) demonstrated that anthrax lethal toxin exerts a profound inhibitory effect on the DCs operative by disrupting the MAPK signaling pathway, ultimately resulting in impaired adaptive immunity (43). Salmonella has been reported to inhibit Ag presentation by DCs, which may help this pathogen to escape immune defense, colonize host organs, and persist in the infected host (44, 45, 46). Thus, pathogen-driven modulation of DC maturation may be one mechanism used by microorganisms to evade host immune responses and establish persistent infection. An important finding of this study is that the distinct DC phenotype that is observed only with live, proliferating OmpA+ E. coli and requires direct contact between bacteria and DCs. Treatment of OmpA+ E. coli with bacteriostatic doses of Cm lead to up-regulation of costimulatory markers, indicating that the OmpA interaction with DC receptor structures for subsequent synthesis of bacterial proteins that could directly interfere with processes leading to an increase in expression of activation markers on DCs. Furthermore, interaction of DCs with killed bacteria, lysates as well as supernatants from live OmpA+ E. coli, lead to enhanced expression of CD40, HLA-DR, and CD86, indicating that the effects of overproduction of bacterial products due to lysis of bacteria in humans would be suppressed by live OmpA+ E. coli. Previous studies by Jeannin et al. (47) demonstrated that recombinant OmpA from Klebsiella pneumoniae interacts with TLR2. However, this interaction induced the maturation of DCs and production of IL-12, which are in contrast to data obtained in this study. Therefore, the interaction of purified OmpA with DCs via TLR2 could be different from OmpA interaction when present in intact E. coli. In fact, the OmpA+ E. coli lysates, which also contain OmpA, induced the up-regulation of DC maturation markers. It is possible that OmpA of E. coli K1 interaction with an unknown cognate receptor on DCs might be responsible for the induction of a bacterial protein, which will be secreted into the medium or in the cytoplasm of DCs for subsequent control of DC maturation. Recent studies with uropathogenic E. coli CFT073 demonstrated that the bacteria secretes a TIR domain-containing protein, TcpC, which impedes TLR signaling by directly binding to MyD88 (38). On par with this concept, the supernatants of DC-E. coli coculture also prevented the maturation of DC; however, it remains to be determined whether E. coli K1 also produces such a protein secreted during the interaction.

DCs produce cytokines following exposure to microbes and its products at the site of infection, which determines whether an immunogenic or tolerogenic immune response will develop (48). The proinflammatory cytokines IL-6, TNF-{alpha}, IL-1β, IL-8, and IL-12 are crucial in the innate and adaptive immune response to infections. These cytokines are important in enhancing the bactericidal capacity of phagocytes, recruiting neutrophils to sites of infection, inducing DC maturation, and directing the specific immune response to invading microbes (49). However, cytokines like IL-10 and TGF-β can mediate potent immunosuppression. Our study revealed that exposure of DCs to live OmpA+ E. coli lead to decreased production of proinflammatory cytokines like IL-6, TNF-{alpha}, IL-1β, and IL-12 and enhanced production of IL-10 and TGF-β. It is well known that IL-10 and TGF-β have an important regulatory role in monocyte function and on DC maturation (50, 51). These cytokines have been shown to inhibit macrophage activation and production of reactive oxygen species, prevent the maturation of DC, induce T cell tolerance and development of T regulatory cells, and suppress TNF-{alpha} production (52, 53, 54). Furthermore, IL-10 and TGF- β have been documented to have a significant inhibitory effect on several aspects of APC function, e.g., the expression of costimulatory molecules and the ability to synthesize IL-12 (55, 56, 57). DCs secreting IL-10 exhibit minimal or no stimulatory properties in primary MLRs and are markedly inhibitory to T cell proliferation induced by polyclonal activators (58). McGuirk et al. (59) showed that DCs exposed to filamentous hemagglutinin from Bordetella pertussis secrete IL-10 and inhibit LPS-induced inflammatory cytokine production. Furthermore, McGuirk et al. (60) demonstrated the ability of these IL-10-secreting DCs to induce the clonal expansion of immunosuppressive T regulatory 1 cells capable of suppressing the Bordetella-specific Th1 immune response. Thus, OmpA+ E. coli can exploit IL-10- and TGF-β-producing DCs, which are functionally and phenotypically inhibitory accessory cells and putatively tolerogenic, for its survival and growth within the host, hence evading potent immune defense mechanisms.

In summary, our results demonstrate a unique aspect of interaction of live OmpA+ E. coli with DCs, which could play an important role in the pathogenesis and immune responses of neonatal meningitis. The ability of live OmpA+ E. coli to interfere with activation of DCs could enhance bacterial survival, promoting dissemination and systemic disease. Characterization of the mechanisms utilized by OmpA+ E. coli to suppress DC maturation is an important observation that could have an impact on the design of efficient vaccines directed to confer protective antimicrobial immunity. Our findings may help in developing alternative preventive approaches against meningitis, based on targeting OmpA epitopes that interact with DC receptors to suppress the DC activation.


   Acknowledgments
 
We sincerely thank Drs. Barbara Driscoll and Catherine Hunter for critical reading of this manuscript.


   Disclosures
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
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 National Institutes of Health Grant AI40567 (to N.V.P.). Back

2 Address correspondence and reprint requests to Dr. Nemani V. Prasadarao, Division of Infectious Diseases, MS 51, Childrens Hospital Los Angeles, 4650 Sunset Boulevard, Los Angeles, CA 90027. E-mail address: pnemani{at}chla.usc.edu Back

3 Abbreviations used in this paper: OmpA, outer membrane protein A; DC, dendritic cell; C4bp, C4b-binding protein; PGN, peptidoglycan; MM, maturation mixture; HBMEC, human brain microvascular endothelial cell; PI, propidium iodide; moi, multiplicity of infection; Cm, chloroamphenicol. Back

Received for publication April 10, 2008. Accepted for publication June 7, 2008.


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 Materials and Methods
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