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Published as doi: 10.1096/fj.06-5910fje.
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(The FASEB Journal. 2006;20:2153-2155.)
© 2006 FASEB

Cooperative molecular and cellular networks regulate Toll-like receptor-dependent inflammatory responses

Gavin E. Morris*, Lisa C. Parker*, Jon R. Ward*, Elizabeth C. Jones*, Moira K. B. Whyte*, Christopher E. Brightling#, Peter Bradding#, Steven K. Dower{dagger} and Ian Sabroe*,1

Academic Units of
* Respiratory Medicine and

{dagger} Cell Biology, Division of Genomic Medicine, University of Sheffield, Sheffield, UK; and

# Institute for Lung Health, Department of Infection, Inflammation and Immunity, Leicester-Warwick Medical School and University Hospitals of Leicester, Leicester, UK

1Correspondence: Academic Unit of Respiratory Medicine, Division of Genomic Medicine, University of Sheffield, M Floor, Royal Hallamshire Hospital, Sheffield, S10 2JF, UK. E-mail i.sabroe{at}sheffield.ac.uk

ABSTRACT

Viral and bacterial pathogens cause inflammation via Toll-like receptor (TLR) signaling. We have shown that effective responses to LPS may depend on cooperative interactions between TLR-expressing leukocytes and TLR-negative tissue cells. The aim of this work was to determine the roles of such networks in response to agonists of TLRs associated with antiviral and autoimmune responses. The TLR3 agonist poly(I:C) activated epithelial cells, primary endothelial cells, and two types of primary human smooth muscle cells (airway [ASMC] and vascular) directly, while the TLR7/8 agonist R848 required the presence of leukocytes to activate ASMC. In keeping with these data, ASMC expressed TLR3 but not TLR7 or TLR8. Activation of ASMC by poly(I:C) induced a specific cytokine repertoire characterized by induction of CXCL10 generation and the potential to recruit mast cells. We subsequently explored the ability of TLR agonists to cooperate in the induction of inflammation. Dual stimulation with LPS and poly(I:C) caused enhanced cytokine generation from epithelial and smooth muscle cells when in the presence of leukocytes. Thus, inflammatory responses to pathogens are regulated by networks in which patterns of TLR expression and colocalization of tissue cells and leukocytes are critical.—Morris, G. E., Parker, L. C., Ward, J. R., Jones, E. C., Whyte, M. K. B., Brightling, C. E., Bradding, P., Dower, S. K., Sabroe, I. Cooperative molecular and cellular networks regulate Toll-like receptor-dependent inflammatory responses.


Key Words: airway smooth muscle • TLR • asthma

TOLL-LIKE RECEPTOR (TLR) signaling is crucial to effective responses to pathogens, but there is increasing evidence that such responses may contribute to disease. Detrimental inflammation may arise as a result of excessive responses to pathogens or as a consequence of activation of TLRs by endogenous agonists (1) . TLR4 mediates responses to LPS, and also to a variety of endogenous agonists (2) . We have shown that responses mediated via TLR4 can be potently amplified when innate immune cells and tissue cells cooperate. Using primary human tissue cells and primary human leukocytes to model interactions likely to be relevant to human disease, we recently showed that airway smooth muscle cells (ASMCs) are unresponsive to bacterial stimuli such as LPS. Coculture of ASMCs and monocytic cells enabled a profound collaborative response to LPS that was dependent on interleukin-1ß (IL-1ß) generation by monocytes, which in turn is responsible for substantial proinflammatory responses in the ASMC (3) . Other evidence suggests that such cooperative systems are likely to be important in vivo (4) .

In this manuscript, we sought to determine the relevance of such cooperative networks in mediating inflammation in response to stimuli activating TLRs implicated in antiviral and autoimmune responses. TLR3 responds to double-stranded viral RNA (5 , 6) and mRNA (7) , and is implicated in autoimmune diseases (8 , 9) . TLR7 and TLR8 respond to single-stranded viral RNA (10 11 12) , and TLR9 (originally identified as a receptor for bacterial DNA; ref. 13 , 14 ) also detects viral DNA (15 , 16) . TLR4 also mediates responses to some viruses (17) . TLR3 is expressed by tissues including epithelial cells (18) , skeletal muscle cells (19) , and endothelial cells (20) , but by only a subset of leukocytes, principally dendritic cells (21) . In contrast, TLR7 and TLR8 expression has been described in cardiac muscle (22) but is perhaps more evident in leukocytes (23) . We hypothesized that optimal responses to pathogens mimicked by agonists of these TLRs would require cooperative signaling between TLRs, and between tissue cells and leukocytes.

MATERIALS AND METHODS

Cells and reagents
Reagents were purchased from Sigma-Aldrich (Poole, UK) or Invitrogen (Paisley, UK), unless specified. The TLR3 agonist poly(I:C) was from both Invivogen (San Diego, CA, USA) and Amersham Biosciences (Chalfont St. Giles, UK). The TLR7/8 imidazoquinoline agonists R848 and gardiquimod (23 24 25) were from Invivogen, and purified LPS (from E. coli serotype R515) from Alexis (Nottingham, UK). Matched ELISA antibody (Ab) pairs were from the National Institute for Biological Standards and Controls (Potters Bar, UK) and R&D Systems (Abingdon, UK). The complete human IFNß ELISA kit was also from R&D systems. TNF{alpha} and IL-1ß were from Peprotech EC (London, UK). Primary cells were maintained in the manufacturer’s recommended media. Human umbilical vein endothelial cells (HUVECs) were obtained from Promocell (Heidelberg, Germany) and used between passages 3 and 5. HUVECs were plated in gelatin-coated 12-well plates; monolayers were used when confluent and stimulated with the indicated agonists in the presence of 2% FCS. Human aortic vascular smooth muscle cells (AoVSMCs) were obtained from Cascade Biologics (Mansfield, Nottinghamshire, UK) and used between passages 2 and 6. AoVSMC were plated in 24-well plates, grown to ~70% confluence, then cultured for 48 h in Dulbecco’s modified Eagle medium (DMEM) with reduced serum (0.2%) before use. Stimulation of AoVSMC with the indicated agonists was performed in the presence of 2% serum. The immortalized epithelial cell line BEAS-2B was maintained and stimulated in 6- or 24-well plates in RPMI + 10% FCS and antibiotics. Primary human airway smooth muscle cells (ASMCs) from four donors were purchased from Cambrex Bioscience (Wokingham, UK); cells from a fifth donor, purchased from Promocell, were maintained as described (3) and used between passages 2 and 7 only. ASMCs were plated in 24-well plates and confluent monolayers were starved of serum for 48 h prior to stimulation except where noted (3) . Stimulation of airway smooth muscle cells was typically carried out in the absence of serum unless otherwise noted, such as those experiments comparing responses in the presence or absence of PBMC [where FCS was present at a concentration of 2% (Fig. 4) or 0.2% (Fig. 6) , in accordance with our previous methodologies (3) ].


Figure 1
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  		Figure 1. Airway smooth muscle cells and the airway epithelial cell line BEAS-2B respond to the TLR3 agonist, poly(I:C). ASMCs were grown to confluence in 24-well plates and starved of serum before stimulation. BEAS-2B cells were grown to confluence in 24-well plates. Cells were then stimulated with poly(I:C) or poly(dI:dC) at the indicated concentrations. After 24 h, generation of CXCL8 and IL-6 was measured by ELISA. A, B) CXCL8 and IL-6 generation from ASMCs; C) CXCL8 generation from BEAS-2B cells. Data are mean ±SE from 3–4 replicates from one ASMC donor, each replicate being performed on a separate passage and from 4 passages of BEAS-2B cells.


Figure 2
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  		Figure 2. Proinflammatory cytokines and poly(I:C) cooperate to activate ASMCs and BEAS-2B cells. A, B) ASMCs were grown to confluence in 24-well plates and starved of serum before stimulation. BEAS-2B cells were grown to confluence in 24-well plates. Cells were stimulated with poly(I:C) or poly(dI:dC) (25 µg/ml) in the presence of media, IL-1ß (10 ng/ml), or TNF{alpha} (10 ng/ml). After 24 h, cytokine generation was measured by ELISA. Data shown in panel A are mean ±SE for 4–5 replicates from one ASMC donor, each from a separate passage. Similar data were obtained using ASMCs from another donor (data not shown). Data in panel B are mean ±SE for 4 passages of BEAS-2B cells. *P < 0.05, **P < 0.01 and ***P < 0.001 for poly(I:C) vs. poly(dI:dC) samples stimulated with media or the indicated cytokine (e.g., samples stimulated with poly(I:C) and IL-1ß produced significantly more CXCL8 than samples stimulated with poly(dI:dC) and IL-1ß). Data are analyzed by 2-way ANOVA with Bonferroni’s post test. C, D) Subsequently, ASMCs were grown to confluence, starved of serum, and stimulated with poly(I:C) (25 µg/ml), IL-1ß (10 ng/ml), TNF{alpha} (100 ng/ml), or combinations of these. After 24 h, ICAM-1 expression on the cell surface was determined by flow cytometry. C) Representative histograms of control IgG binding, vs. binding of anti-ICAM-1 Ab to cells stimulated with media alone, poly(I:C), TNF{alpha}, or poly(I:C) + TNF{alpha}. Mean data (n=4±SE, each replicate from a separate passage) are shown in panel D. ***Significant differences (P < 0.001) for the indicated comparisons analyzed by ANOVA and Tukey’s post test. E) The effects of supernatants from poly(I:C) -activated tissue cells on mast cell chemotaxis. ASMCs were grown to confluence, starved of serum, and stimulated with media or poly(I:C) (25 µg/ml). Supernatants were prepared and tested at a 1:3 dilution in DMEM + 2% FCS for their ability to induce chemotaxis of HMC-1 mast cells as described. Data were normalized to the appropriate control (HMC-1 migration in response to poly(I:C) diluted in appropriately in ASMC medium, but that had not been applied to ASMC cultures), and expressed as fold change in migration. Samples were tested in the presence of a blocking anti-CXCR3 Ab or a matched control. Data shown are mean ±SE of three supernatants, each generated from a separate passage of ASMCs, tested on one passage of HMC-1 cells, and analyzed by paired Student’s t test, where *P < 0.05.


Figure 3
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  		Figure 3. Airway smooth muscle cells express TLR3 intracellularly, and BEAS-2B cells express TLR3 both intracellulary and extracellularly. A) Airway smooth muscle cells were analyzed for expression of TLR3 by polymerase chain reaction (PCR). Lanes 1, 5, and 9 show a 100 bp MW marker ladder. Lanes 2, 6 show TLR3 expression; lanes 3 and 7, TLR7 expression, and lanes 4 and 8, TLR8 expression in two different ASM donors (the first analyzed from mRNA prepared from growing cells, the second from mRNA from cells in stationary growth phase after serum deprivation). In additional experiments, ASMCs (grown to confluence and starved of serum) or BEAS-2B cells were stimulated with media alone, poly(I:C) 25 µg/ml, or poly(I:C) + TNF{alpha} (100 ng/ml). Cells were harvested 24 h after stimulation and stained for TLR3 cell surface expression or permeabilized and stained for total extracellular + intracellular TLR3 expression. Ab binding was determined by flow cytometry and calculated as specific geometric mean (D, ASMCs) or specific mean (H, BEAS-2B cells) fluorescence by subtraction of the fluorescence of the isotype control. Representative histograms of intracellular and extracellular staining respectively for the IgG control, for anti-TLR3 binding in cells stimulated with media alone, and cells stimulated with poly(I:C) + TNF{alpha} in B, C) ASMCs, F, G) BEAS-2B cells. B) Three histograms are superimposed on each other. D, H) Mean data (n=5–6 replicates, each from a separate passage, of one ASMC donor representative of results from two independent ASMC donors; n=3–5 from BEAS-2B cells). E, I) The effects of chloroquine on poly(I:C) signaling in ASMCs (E) and BEAS-2B cells (I) were determined. ASMCs were starved of serum for 24 h and stimulated in the presence of 2% FCS. BEAS-2B cells were stimulated as described in Materials and Methods. Chloroquine was applied 60 min prior to poly(I:C) and CXCL8 generation measured 24 h later by ELISA. Data shown are n = 4 for BEAS-2B cells and n = 3 replicates, each from a separate passage, of one ASMC donor. Significant differences between the indicated comparisons are shown by *P < 0.05 (Student’s t test).


Figure 4
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  		Figure 4. Activators of TLR7/8 stimulate ASMCs indirectly via PBMCs. A, B) ASMCs were grown to confluence in 24-well plates, starved of serum, and stimulated with the indicated concentrations of R848 in the presence or absence of PBMCs (30,000 cells per well). After 24 h, levels of CXCL8 and IL-6 in the media were determined by ELISA. Data shown are n = 3–4 ±SE, each replicate being performed at a separate passage with freshly prepared PBMCs from separate donors. Significant differences in cytokine production between stimulated ASMC monolayers and ASMC/PBMC cocultures are indicated by **P < 0.01 and ***P < 0.001, measured by 2-way ANOVA and Bonferroni’s post test. Similar results were obtained using cells from an independent ASMC donor (data not shown). C) ASMCs from another donor. PBMCs (30,000 cells/well) or cocultures of ASMCs and PBMCs were grown to confluence in 24-well plates, starved of serum for 24 h, and stimulated with the indicated concentrations of an alternative imidazoquinoline, gardiquimod. After 24 h, levels of CXCL8 in the media were determined by ELISA. Data shown are n = 3 ±SE, each replicate being performed at a separate passage with freshly prepared PBMCs from separate donors. Significant differences in cytokine production between stimulated monocultures and ASMC/PBMC cocultures are indicated by **P < 0.01, measured by 2-way ANOVA and Bonferroni’s post test.


Figure 5
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  		Figure 5. Human umbilical vein endothelial cells and human aortic vascular smooth muscle cells respond to the TLR3 agonist, poly(I:C). HUVECs were grown to confluence in 12-well plates and AoVSMCs were grown to confluence before being starved in 0.2% serum prior to use. Cells were then stimulated with poly(I:C) (25 µg/ml) or LPS (1000 ng/ml) in the presence or absence of monocytes enriched from PBMC by negative magnetic selection at a concentration of 2 x 104/ml. After 24 h, generation of CXCL8 and IL-6 was measured by ELISA. A) CXCL8 release from HUVECs stimulated with the indicated agonist, and B) IL-6 release from HUVECs stimulated with the indicated agonists. C) CXCL8 release from AoVSMCs stimulated with the indicated agonists; D) IL-6 release from AoVSMCs stimulated with the indicated agonists. Data are mean ±SE from 4–5 replicates from one donor, each replicate being performed on a separate passage and using enriched monocytes from separate donors. Data were analyzed using 2-way ANOVA and Bonferroni’s post test; relevant significant differences are indicated; ***P < 0.001.


Figure 6
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  		Figure 6. Cooperative production of cytokines in cocultures. ASMCs or BEAS-2B cells were grown to confluence in 24-well plates and stimulated with poly(I:C) (25 µg/ml), LPS (10 ng/ml), or both in the presence or absence of PBMCs (10,000 cells per well). After 24 h, levels of CXCL8 and CXCL10 in the media were determined by ELISA. A) The effects of the indicated agonists on CXCL8 release from ASMCs in the presence and absence of PBMCs; B) the effects of the indicated agonists on the release of CXCL10 from ASMCs in the presence or absence of PBMCs. C) the effects of the indicated agonists on the release of CXCL8 from BEAS-2B cells in the presence or absence of PBMCs. In each graph, open bars show cytokine generation from monolayers of the relevant tissue cell (ASMCs or BEAS-2B cells), shaded bars show cytokine generation from wells containing 10,000 PBMCs only, and filled bars show cytokine generation from cocultures of PBMCs and the indicated tissue cell. Data shown are from 4 passages of BEAS-2B cells, and from 5 passages of one ASMC donor. Selected relevant comparisons were performed using Student’s t test and the results indicated on the figure.

ELISA
Cell-free supernatants were prepared from each experiment and stored at –80°C until cytokine generation was determined by ELISA. Samples were appropriately diluted to ensure analysis within the linear portions of a log (concentration)/linear (optical density) standard curve. Samples in which no cytokine could be detected were assigned values equivalent to the lower limit of assay detection in each assay (typical values being CXCL8=15 pg/ml, IL-6=20 pg/ml, CCL5=78 pg/ml, CXCL10=39 pg/ml, IFNß=250 pg/ml).

Flow cytometry
PE-conjugated antibodies to TLR3 and ICAM-1, and their matched isotype controls, were from eBioscience (San Diego, CA, USA). Cells were detached by trypsinization (optimized in separate experiments to allow removal of cells from plasticware with minimal trypsin exposure, and found not to cause loss of cell surface receptors such as TLR4 from peripheral blood monocytes). Trypsin was immediately neutralized, and the cells were washed and resuspended in buffer (PBS without Ca2+/Mg2+, 10 mM HEPES, 0.25% BSA). Staining with antibodies (5 µg/ml anti-TLR3, 5 µg/ml anti-ICAM-1) was performed for 40 min. Intracellular staining was quantified after fixation and permeabilization using a commercial fix/permeabilize buffer system (eBioscience) according to the manufacturer’s instructions. Ab binding was detected using a FACSCalibur flow cytometer using CellQuest software (Becton-Dickinson, Cowley, UK) and quantified as specific mean fluorescence after subtraction of the isotype-matched control Ab fluorescence.

Preparation of leukocytes
Peripheral blood mononuclear cells (PBMCs) were prepared fresh for each experiment from venous blood of healthy volunteers taken in accordance with a protocol approved by the local research ethics committee. PBMCs were enriched by centrifugation over density gradients of Histopaque 1077 or plasma/Percoll as described (3 , 26) . In some experiments, monocytes were further enriched from PBMCs by negative magnetic selection. Briefly, PBMCs were incubated with the monocyte enrichment negative selection Ab cocktail from StemCell Technologies (Vancouver, Canada) according to the manufacturer’s instructions, and cells were separated in a magnetic field as described (27) . Purity of resulting monocyte populations was determined by CD14+ staining and flow cytometry, and found to be typically between 60 and 80%.

Mast cell chemotaxis
Mast cell chemotaxis assays were performed using the HMC-1 mast cell line as described (28) . Briefly, 1 x 105 cells were placed in a fibronectin-coated, 8 µm pore-sized Transwell insert (Becton-Dickinson) and cells were allowed to migrate in the presence of media or the indicated tissue culture supernatants, in the presence or absence of a blocking CXCR3 Ab (R&D Systems) or a matched control for 4 h prior to counting (28) .

Statistics
Data were tested statistically and graphed using GraphPad Prism v4.0 (GraphPad Inc, San Diego, CA, USA), using the statistical tests indicated in the figure legends.

RESULTS

Poly(I:C) induces proinflammatory cytokine generation by a panel of tissue cells
The ability of the TLR3 agonist poly(I:C), a mimic of double-stranded RNA, to cause proinflammatory responses in a panel of tissue cells was investigated. Figure 1 shows for the first time that primary human ASMCs responded to poly(I:C) as determined by the induction of CXCL8 and IL-6, but did not respond to the control agonist, poly(dI:dC). Induction of cytokines in response to poly(I:C) was observed in cells from each of 5 independent donors (data from all donors used in this study are summarized in Table 1 ). In three of the four other donors, levels of CXCL8 generated were similar to those shown in Fig. 1 , and in one donor levels of cytokine generation were approximately an order of magnitude higher. Poly(I:C) induced similar amounts of CXCL8 production from the BEAS-2B immortalized human lung epithelial cell line (Fig. 1) . Induction of IL-6 from poly(I:C)-treated BEAS-2B cells was modest and failed to reach significance [164±119 (SEM) pg/ml in poly(dI:dC)-treated cells vs. 606±151 pg/ml in cells treated with 25 µg/ml poly(I:C), P = 0.06 Student’s t test]. Primary human aortic smooth muscle cells (AoVSMCs) and HUVECs also released CXCL8 and IL-6 in response to poly(I:C) (Fig. 5) .


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  		Table 1. Variations in cytokine generation between donorsa

In other experiments, the inflammatory response induced by poly(I:C) was characterized in more detail using primary human ASMCs. Production of the pleiotropic cytokine IL-6 and the Th1-associated chemokine CXCL10 was also measured in 3 (IL-6) and 2 (CXCL10) of the 5 donors, and paralleled the amounts of CXCL8 generated (summarized in Table 1 ). Control reagents, including poly(dI:dC) (a double-stranded DNA mimic) and poly(C) (a single strand of a synthetic RNA mimic), failed to induce cytokine generation from monocultures of ASMCs (Fig. 1 and data not shown), and poly(dI:dC) did not induce cytokine generation from BEAS-2B epithelial cells (Fig. 1) .

There is increasing evidence that apical cytokines such as IL-1 and TNF{alpha} play central roles in the regulation of inflammation in a range of diseases and across a range of tissues. Figure 2 A, B shows that stimulation of ASMCs or BEAS-2B cells with either IL-1ß or TNF{alpha} resulted in a marked induction of poly(I:C)-mediated production of proinflammatory cytokines. The control poly(dI:dC) was inert in these assays, as responses to poly(dI:dC) ± IL-1ß or TNF{alpha} were similar to those seen in cells stimulated with media ± IL-1ß or TNF{alpha} (data not shown). Activation of tissue cells can also be shown by their up-regulation of adhesion molecules such as ICAM-1, the molecule by which rhinovirus gains access to airway cells and that is up-regulated on viral-infected ASMCs (29) . Up-regulation of ICAM-1 in response to poly(I:C) or cytokines, including IL-1ß and TNF{alpha}, was observed in ASMCs (Figs. 2C, D) . Dual stimulation with poly(I:C) and IL-1ß caused a greater induction of ICAM-1 expression compared with stimulation with each agonist individually, an effect not seen in the poly(I:C)/TNF{alpha} combination, though the effect of TNF{alpha} alone was very marked and it is possible that synergy between poly(I:C) and TNF{alpha} would have been observed at lower TNF{alpha} concentrations (Fig. 2D ).

CXCL10 has a central role in the regulation of mast cell recruitment to the airway smooth muscle (30) . We therefore investigated the potential of poly(I:C) to stimulate recruitment of the CXCL10-responsive HMC-1 mast cell line (28) to ASMCs. Exposure of HMC-1 cells to media containing poly(I:C) in the lower well of a chemotaxis plate reduced their baseline migration levels by ~70% compared with cells exposed to media alone (data not shown), consistent with a direct effect of poly(I:C) on mast cell function (31 32 33) . Exposure of HMC-1 cells to media transferred from ASMCs treated with poly(I:C) resulted in an increase in mast cell chemotaxis compared with cells exposed to the relevant control (media containing poly(I:C) that had not been applied to ASMCs). This increase in chemotaxis was inhibited by an Ab to the CXCL10 receptor, CXCR3, but not by an isotype control (Fig. 2E ).

TLR3 can be expressed intracellularly and/or extracellularly
TLR3, but not TLRs 7 or 8, were detected in ASMCs by RT-polymerase chain reaction (RT-PCR) (Fig. 3 A). TLR3 is typically, though not exclusively, expressed intracellularly (34) . Using flow cytometry, we observed no expression of TLR3 on the surface of ASMCs, but a marked binding of anti-TLR3 in permeabilized cells (Fig. 3) . Culture of ASMCs with poly(I:C) or TNF{alpha}, or combinations of poly(I:C) and IL-1ß or TNF{alpha} resulted in minimal changes in TLR3 expression in permeabilized cells, which failed to reach statistical significance in mean data (Figs. 3C, D) . In contrast, BEAS-2B cells expressed TLR3 on the cell surface and showed a further increase in total anti-TLR3 staining in permeabilized cells, indicating the presence of additional TLR3 intracellularly (Figs. 3 F, G). In BEAS-2B cells, extracellular TLR3 expression was modestly up-regulated by poly(I:C) + TNF{alpha} stimulation alone (Fig. 3H ).

TLR3 signaling is generally held to require endosomal acidification (35 , 36) , though there is evidence that in some cells, signaling may occur from the cell surface (32 , 37) . We found that an inhibitor of endosomal acidification, chloroquine, inhibited CXCL8 generation in response to poly(I:C) treatment in ASMCs but not BEAS-2B epithelial cells (Fig. 3) .

ASMCs do not themselves respond to TLR7/8 activation, but respond to imidazoquinolines in coculture with PBMCs
The work above showed that ASMCs express TLR3 and respond to its agonist, poly(I:C). We also showed that ASMCs do not express mRNAs for TLR7 or TLR8. In keeping with this observation, we found that the TLR7/8 agonist R848 did not cause the generation of IL-6 or CXCL8 from ASMCs at the concentrations tested (Fig. 4) . We therefore explored the potential of leukocytes and tissue cells to cooperate in signaling mediating antiviral and autoimmune responses. We constituted cocultures that contained 30,000 PBMC and 50,000 ASMCs per well as described previously (3) . When these cocultures were stimulated with R848, marked production of CXCL8 and IL-6 was observed that was not seen in ASMC monocultures (Fig. 4 ). In separate experiments, we observed that PBMCs stimulated with R848 at the cell density used in these cocultures (30,000 cells/well) generated only low amounts of CXCL8 (2.6±0.3 ng/ml, mean±SE, n=3). These actions were shared by gardiquimod, another member of the imidazoquinoline family, which also caused substantial CXCL8 generation from cocultures but not ASMC or PBMC monocultures (Fig. 4C ).

Activation of tissue cells by poly(I:C) is not enhanced by coculture with PBMCs, but can be enhanced by coculture with PBMCs and LPS
We observed above that activation of tissue cells by poly(I:C) could be mediated directly, while agonists of TLRs 7&8 (these data) or TLR4 (3) required PBMC to initiate cytokine production from ASMC. The activating cell type within PBMC populations was previously shown to be the monocyte (3) . In keeping with these results, we also found that responses of HUVEC and AoVSMC to LPS were substantially amplified in the presence of purified monocytes (Fig. 5 ), where the numbers of monocytes were sufficiently low (20,000 cells/ml) that detectable CXCL8 and IL-6 production from the monocytes alone was minimal. We therefore determined whether responses to poly(I:C) were entirely independent of monocytes or could be amplified by their presence in coculture with tissue cells. Figures 5 and 6 show that activation of ASMC, AoVSMC, HUVEC, and BEAS-2B cells by direct stimulation with poly(I:C) was not enhanced by the presence of PBMC as a source of monocytes (for ASMC and BEAS-2B cells, Fig. 6 ) or purified monocytes themselves (for HUVECs and AoVSMCs, Fig. 5 ). Thus, our data reveal two patterns of tissue cell responses to TLR agonists: direct activation by TLR agonists that is not amplified by coculture with PBMC (TLR3), and indirect activation that is PBMC dependent (TLR4 and TLR7/8). Strikingly, LPS-activated PBMC/ASMC cocultures did not result in generation of CXCL10 (Fig. 6B ), demonstrating specific and compartmentalized responses to individual TLR agonists in mono- and cocultures.

Inflammatory sites are often exposed to more than one TLR agonist. For example, during respiratory tract viral infections patients will continue to inhale environmental endotoxin and may develop superadded bacterial infections. Likewise, local tissue damage during inflammatory responses can generate endogenous agonists of TLR3 [mRNA (7) ] and TLR4 [HMGB1 (2) , fibrinogen (38) ]. To determine the consequences of exposure to multiple TLR agonists on inflammatory sites, we stimulated ASMC/PBMC and BEAS-2B/PBMC cocultures with poly(I:C) and LPS in combination. Figure 6 demonstrates that BEAS-2B/PBMC cocultures showed a modest cooperative production of CXCL8 when stimulated with poly(I:C) + LPS, while ASMC/PBMC cocultures showed a more marked cooperative production of CXCL10, though not CXCL8, when activated with poly(I:C) + LPS.

DISCUSSION

In this study we have identified cooperative mechanisms by which signals acting via TLRs can regulate inflammatory responses to viral infections and endogenous mediators of inflammation that have the potential to contribute substantially in a range of inflammatory diseases.

We have shown for the first time that TLR3 is expressed by ASMCs, and confirmed other studies showing TLR3 expression in BEAS-2B epithelial cells (18) and that the TLR3 agonist poly(I:C) drives proinflammatory cytokine generation from these cells. We also observed that poly(I:C) activated vascular smooth muscle; together with data showing expression of TLR3 by skeletal muscle (19) , these data suggest widespread functional expression of this receptor. We have also demonstrated, again in keeping with recent literature, that poly(I:C) can stimulate HUVEC (20) .

Activation of ASMC or BEAS-2B monolayers by poly(I:C) caused the generation of CXCL10, which was not seen when these cells were stimulated with IL-1ß or TNF{alpha} alone or in ASMC stimulated with LPS-activated PBMCs. These data demonstrate selective responses to TLR agonists in tissue cells. CXCL10 (IP-10) is a Th1-associated chemokine whose generation in response to TLR agonists is typically dependent on production of type 1 interferons (39 , 40) . This chemokine has the potential to recruit Th1-type T cells to the lung (41 , 42) , and has recently been identified as the chemokine responsible for the recruitment of mast cells to airway smooth muscle bundles (28 , 30) , a specific feature of asthmatic airways (43) . In keeping with these studies, media from ASMCs treated with poly(I:C) caused CXCR3-dependent mast cell chemotaxis with an efficacy similar to that observed for supernatants generated from ASMCs activated with cytokines such as IL-1ß (28) . We also observed that media controls containing poly(I:C) reduced basal mast cell migration; although the exact mechanism behind this effect of poly(I:C) remains to be determined, these data are consistent with the ability of mast cells to be activated via TLR3 (32 , 33 , 44) . Thus, viral infection and activation of ASMCs may contribute to asthma exacerbations not just through the production of acute inflammation, but also through the regulation of mast cell trafficking.

TLR3 expression is typically intracellular (34) , and was recently observed in lung epithelial cell lines and endothelial cells, demonstrating a wide tissue expression of this receptor (20 , 45) . Signaling via TLR3 is generally thought to occur in acidified endosomes (35 , 36 , 46) . In keeping with these observations, we found that an inhibitor of endosomal acidification, chloroquine, reduced levels of CXCL8 generated from poly(I:C) activated ASMCs. In contrast, chloroquine treatment failed to inhibit poly(I:C) responses in BEAS-2B cells, which express cell surface TLR3. Other studies have demonstrated a potential ability of TLR3 to signal at the cell surface (37 , 47) that would be in keeping with our results, but not in keeping with a requirement for a low pH for TLR3 to signal. Poly(I:C) can also signal via other innate immune recognition systems, such as protein kinase R (PKR) (48) and recently identified cytosolic signaling systems involving MDA5 and IPS-1 (46 , 49) . The close correlation of TLR3 expression patterns with the activation of ASMC monocultures and ASMC/PBMC cocultures indicates a major role for TLR3 in the responses we observed, but additional contributions may also arise from IPS-1/MDA5 or PKR that could provide an alternative explanation for the chloroquine-resistant signaling in BEAS-2B cells. These possibilities are the subject of ongoing work in our group.

Tissari et al. failed to find TLR7 and TLR8 expression in A549 epithelial cells and found no response of these cells to the TLR7/8 agonist, R848 (20) . We observed that ASMCs also failed to respond to R848, but found that stimulation of TLR7 and/or TLR8 could activate ASMCs through indirect mechanisms. We previously showed that cocultures of ASMCs and PBMCs enabled a profound cooperative response to LPS, with marked activation of cytokine generation from the LPS-unresponsive ASMCs (3) . In keeping with the response of PBMCs to TLR7 agonists (23) , we found that cocultures of ASMCs and PBMCs stimulated with R848 showed a cooperative response resulting in the generation of CXCL8 and IL-6. Given the low numbers of PBMCs/well in coculture, these cytokines are again likely to have originated almost entirely from the ASMCs, as observed in cocultures activated with LPS (3) . An additional imidazoquinoline, gardiquimod, markedly activated cocultures of ASMCs and PBMCs but not monocultures of each cell type (Fig. 4C ). These data reinforce the potential importance of cellular networks in the regulation of responses to TLR agonists in tissues and provide both direct and indirect mechanisms by which tissue cells, which often have a profound synthetic ability with respect to cytokine production, may contribute significantly to the induction of a proinflammatory response when the host is subject to microbial attack. Whereas LPS-induced signaling in cocultures of ASMCs and PBMCs is highly IL-1 dependent (3) , where signaling is being driven by poly(I:C) or imidazoquinolines there is marked potential for involvement of type I interferons in autocrine or paracrine amplification of signaling. We were unable to detect significant IFNß protein from ASMCs activated with poly(I:C) or cocultures of ASMCs and PBMCs activated with gardiquimod (data not shown). These data do not exclude roles for type I IFNs in these settings since local concentrations of released cytokines in culture may be much greater than are seen in the culture supernatant as a whole, and the role of IFNs is the subject of ongoing work.

We observed a synergistic interaction between IL-1ß or TNF{alpha} and poly(I:C) in their ability to stimulate ASMCs and BEAS-2B cells to release cytokines including IL-6, CXCL8, and CXCL10, and between IL-1ß and poly(I:C) in their ability to up-regulate expression of the ICAM-1 integrin on ASMCs. Poly(I:C), IL-1ß and TNF{alpha} (alone or in combination) induced only minimal change in TLR3 expression in BEAS-2B and airway smooth muscle cells over the time courses studied that failed to reach significance in all but one condition, suggesting that the synergy between IL-1ß, TNF{alpha}, and poly(I:C) was not mediated at the level of TLR3 expression. Our previous work firmly established a central role for IL-1ß in the cooperative response to PBMC/ASMC cocultures activated with LPS (3) . TLR3 is not widely expressed by PBMC (being present on some DCs only; ref. 21 , 50 ), and thus shows an inverse expression pattern to TLR4, which we showed in previous work is expressed on many PBMCs but less so on ASMCs (3) . We therefore hypothesized that stimulation of tissue cell/PBMC cocultures by poly(I:C) and LPS would result in greater enhancement of the inflammatory response. We observed that this dual stimulation of cocultures did indeed result in a modest enhancement of CXCL8 generation from BEAS-2B/PBMC cocultures and in a more significant induction of CXCL10 from ASMC/PBMC cocultures.

These data provide mechanisms by which tissue damage at sites of inflammation might cooperatively activate inflammatory responses in networks dependent on multiple TLRs and multiple cell types. Within the lung, these data suggest that a viral infection might have the potential to sensitize an asthmatic individual such that they then respond to normal levels of inhaled LPS present in their environment, products of tissue damage, or superadded bacterial infection, with an exaggerated, PBMC-dependent inflammation resulting in further recruitment of inflammatory cells such as mast cells. The correlation of environmental endotoxin levels and asthma/risk of wheezing illness (51) (this is a complicated area; see ref. 52 53 54 ), together with the potential associations between respiratory virus infection and wheezy illness early in life, point toward mechanisms that are likely to be linked pathologically, and the mechanisms we describe here may be a component of this link. These data are also relevant to the pathological mechanisms of viral-induced wheeze in childhood and have relevance to viral exacerbations of chronic obstructive pulmonary disease. Such mechanisms of cooperative signaling between cell types and TLRs are likely to be of broad importance in the regulation of inflammation within a broad range of tissues and diseases, and the models illustrated here provide systems in which to dissect inflammatory mechanisms and develop novel therapies.

ACKNOWLEDGMENTS

The authors thank Amanda Sutcliffe for helpful assistance with the mast cell chemotaxis assays, and Kathryn Vaughan and Brenka McCabe for assistance in the preparation of PBMCs. This work was funded by a UCB Pharma (Belgium) Ph.D. studentship to G.E.M., the Medical Research Council UK via a Senior Clinical Fellowship (G116/170) to I.S., the British Heart Foundation to J.R.W. (grants FF/02/030 and FS/06/004), and through a Department of Health Clinician Scientist Award to C.E.B.

Received for publication February 24, 2006. Accepted for publication May 8, 2006.

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