American Journal of Respiratory and Critical Care Medicine

Rationale: Toll-like receptors 2 and 4 (TLR2, TLR4) enable cellular responses to bacterial lipoproteins, LPS, and endogenous mediators of cell damage. They have an established role in the activation of leukocytes, endothelial cells, and some smooth muscle cell types, but their roles in airway smooth muscle are uncertain. Objectives: To determine the roles of TLRs in activation of airway smooth muscle. Methods: Airway smooth muscle cells were cultured with TLR agonists, in the presence or absence of mononuclear leukocytes. Measurements and Main Results: We observed expression of TLR2 and TLR4 mRNAs, which could be upregulated by treatment with proinflammatory cytokines in primary human airway smooth muscle, but no important functional responses to agonists of these TLRs were seen. Coincubation of airway smooth muscle with peripheral blood mononuclear cells, at concentrations as low as 250 mononuclear cells/ml, resulted in a marked cooperative response to TLR stimuli, and synergistic production of cytokines, including chemokines (interleukin [IL-]-8) and IL-6. This cooperative response was greater when monocytes were enriched and was transferable using supernatants from LPS-stimulated peripheral blood mononuclear cells. Activation of cocultures required IL-1 generation from mononuclear cells, and was blocked by IL-1 receptor antagonist, though IL-1 generation alone was not sufficient to account for the magnitude of mononuclear cell–dependent coculture activation. Conclusions: These data indicate that potent amplification of inflammation induced by TLR agonists, such as LPS, may be achieved by cooperativity between airway smooth muscle and leukocytes involved in immune surveillance or inflammation.

Microbial-derived products, such as LPS, are potent initiators of inflammation in lung disease, with potential roles in a range of lung diseases, including asthma, acute respiratory distress syndrome, and chronic obstructive pulmonary disease (1, 2). Toll-like receptors (TLRs) are crucial for detection of these pathogen-associated molecules, with LPS signaling being dependent on TLR4, whereas signaling of gram-positive cell wall products, such as lipopeptides, is dependent on TLR2 (1). TLRs are expressed on cells undertaking immune surveillance roles in the lungs, such as alveolar macrophages (3), but there is also an increasing appreciation of the importance of lung tissue cells in the initiation and orchestration of inflammatory responses. For example, pulmonary epithelial cells express TLRs and are involved in lung responses to LPS (46). There is also growing evidence that TLRs can respond to a range of endogenous agonists present or generated at sites of inflammation, such as fibrinogen (7) and matrix breakdown products (8), providing these receptors with potentially important roles in inflammation, even in the absence of infections.

Airway smooth muscle (ASM) also has considerable potential to contribute to inflammatory lung disease (913). Exposure of the whole lung in vivo to TLR agonists, such as endotoxin, has a proinflammatory effect on multiple cell types, including ASM (14). ASM is capable of generating many cytokines, resulting in amplification of leukocyte recruitment and in the development of specific phenotypes of disease, such as the Th2-type inflammatory response seen in asthma (9, 10).

Recent studies have revealed the complexities of the mechanisms involved in TLR signaling. The long-used biological reagent LPS typically contains both LPS-associated protein, which activates TLR2, and LPS itself, which activates TLR4 (15). These receptors activate distinct signaling pathways (16, 17) and also have the potential to modify expression and signaling of themselves and each other (1820). In addition, TLRs, such as TLR4, can be active on the cell surface, or in specific cell types may function intracellularly (21), and signaling is modified by a range of accessory molecules, including CD14, MD-2, and CD11b/CD18 (1). In light of the identification of the TLRs as the crucial proteins enabling signaling in response to LPS (22, 23) and LPS-associated proteins (15), we examined the potential of TLR2 and TLR4 to mediate direct and indirect activation of ASM.


Reagents were purchased from Sigma-Aldrich (Poole, UK) or Invitrogen (Paisley, UK), except where specified. Pam3CSK4 was from EMC Microcollections (Tübingen, Germany). Commercial LPS (cLPS; Escherichia coli 0111:B4) was from Sigma; purified LPS (pLPS; E. coli K235) was a gift from Professor Stephanie Vogel (University of Maryland). Primary ASM cells from three donors were purchased from Cambrex Bioscience (Wokingham, UK) and cultured in the manufacturer's media. They were used when fully confluent between Passages 2 through 7 in 24-well plates and after 48 hours' starvation in serum-free media. Low concentrations of serum were added during stimulation with TLR agonists to facilitate LPS responsiveness of ASM or leukocytes; similar results were seen with a final concentration of fetal calf serum (FCS) of either 2 or 0.2% (see figure legends). Final volumes in each well of a 24-well plate always totalled 400 μl during experiments. Transwell inserts (0.4-μm pore size) were purchased from Millipore (Watford, UK). Reagents for real-time polymerase chain reaction (PCR) were purchased from Eurogentec (Romsey, UK). Matched ELISA antibody pairs were from the National Institute for Biological Standards and Controls (Potters Bar, UK), except for CCL2, which were from R&D Systems (Abingdon, UK). Cytokines were from Peprotech EC (London, UK), except for interleukin (IL-)-1β, which was from ImmunoKontact (Abingdon, UK). Antitotal and anti–phospho p38 were from Cell Signaling Technology (Beverly, MA) and Promega (Southampton, UK), respectively.

Peripheral Blood Mononuclear Cells

Leukocyte preparations enriched for peripheral blood mononuclear cells (PBMC) were prepared from healthy volunteer donors by centrifugation over density gradients of sterile, endotoxin-free Histopaque 1077 or Percoll, as described (24, 25), after written, informed consent and in accordance with a protocol approved by the local research ethics committee. In some experiments, T cells or monocytes were enriched further from PBMC by negative magnetic selection using EasySep cocktails from StemCell Techologies (Vancouver, BC, Canada).

RNA Studies

Total RNA was prepared using RNeasy kits (Qiagen, Crawley, UK), genomic DNA was removed, and the sample reverse-transcribed as described (25). Reverse transcriptase–PCR was performed using HotStar Taq (Qiagen) over 35 cycles (25). Real-time PCR was performed on duplicate samples using an ABI7900 cycler (PE Applied Biosystems, Foster City, CA) and quantified using individual standard curves for glyceraldehyde-3-phosophate dehydrogenase (GAPDH) and the target gene to establish relative units of gene expression, with no–reverse transcriptase samples studied to confirm lack of genomic DNA contamination. Primers and probe sets for reverse transcriptase–PCR and real-time PCR are listed in the online supplement (Table E1).

Cytokine Generation

Cell-free supernatants from cultured cells were prepared and stored at –70°C. Analysis of cytokine concentrations was by ELISA using matched pairs of antibodies at optimized concentrations, as described (24).

Western Blot Analysis

ASM were grown in six-well plates, and protein from duplicate wells pooled before loading into electrophoresis gels. Equal amounts of protein were analyzed by gel electrophoresis, blotted onto nitrocellulose, and probed with anti–phospho p38 and anti–total p38, detected using a horseradish peroxidase–conjugated secondary antibody and enhanced chemiluminescence. Quantitative signals were derived by densitometric analysis using NIH Image (version 1.62f; National Institutes of Health, Bethesda, MD) and data displayed as the ratio of phospho to total p38, providing correction for loading (26).


Data from more than two groups were examined using analysis of variance with an appropriate post test. Statistical testing and calculation of ELISA data were performed using the Prism 4.0 program from GraphPad, Inc. (San Diego, CA).

LPS Is a Poor Activator of ASM

Preliminary experiments showed the presence of mRNAs for TLRs 1, 2, and 4 in primary human ASM in exponential growth phase (data not shown). Primary human ASM cells were then grown to confluence and serum-starved for 48 hours before stimulation with cLPS (activating TLR2 and TLR4) or the cytokines IL-1β or tumor necrosis factor α (TNF-α). Induction of generation of CXC chemokine ligand (CXCL)8 (IL-8) was determined by real-time PCR. IL-1β and TNF-α induced a potent upregulation of CXCL8, consistent with previous observations (27). In contrast, cLPS induced a comparatively small upregulation of CXCL8 mRNA expression (Figure 1A)

. In keeping with these data, and using ASM from a different donor, IL-1β and TNF-α were able to induce phosphorylation of p38 mitogen-activated protein kinase (MAPK), whereas cLPS did not stimulate this marker of proinflammatory signaling (Figures 1B and 1C).

Proinflammatory Cytokines Upregulate TLR mRNA in ASM, but Only Modestly Upregulate Functional TLR Protein

Our initial experiments found that ASM responses to cLPS were modest compared with responses to IL-1β and TNF-α. We therefore investigated whether treatment of ASM with proinflammatory mediators could upregulate expression of TLR2 and TLR4, potentially enabling responses to microbial-derived agonists. The data in Figure 2

show that TLR2 expression could be moderately upregulated by TNF-α and IL-1β stimulation, whereas TLR4 expression was upregulated principally by IL-1β. Despite upregulation of TLR mRNAs by treatment of ASM with these cytokines, we were unable to see reliable expression of TLR2 or TLR4 protein on the surface of these cells by flow cytometry (data not shown). In further experiments, ASM were treated with IL-1β, TNF-α, or cLPS for 24 hours, the media changed, and the cells stimulated for a further 24 hours with the selective TLR2 agonist, Pam3CSK4, or the selective TLR4 agonist, pLPS. Accumulation of the proinflammatory mediators CXCL8 and IL-6 in the media of the second phase of stimulation was determined by ELISA (Figure 3). Pretreatment of cells with either buffer or cLPS resulted in only very low levels of cytokine production, which were not increased by secondary stimulation with the TLR2 or TLR4 agonist. Pretreatment of ASM with IL-1β or TNF-α resulted in a persistent induction of generation of these cytokines. The TLR4 agonist, pLPS, was unable to further enhance cytokine generation. In contrast, a second stimulation with Pam3CSK4 resulted in an approximate doubling in the amount of IL-6 protein detected in the culture media of cells pretreated with TNF-α.

PBMC Synergize with ASM and TLR Agonists in Initiating a Proinflammatory Response

In other work, we have shown that small numbers of PBMC may significantly amplify TLR responses observed in cells, such as neutrophils (25, 28, 29), and other groups have reported a similar dependence of eosinophils on monocytes for TLR-mediated effects (30). The accumulation of leukocytes in the airway wall, including monocytes and lymphocytes, is a marked feature of inflammatory lung diseases, such as asthma, and monocyte-derived alveolar macrophages are an important immune surveillance cell in the lung. We therefore determined if small numbers of PBMC prepared from the blood of healthy volunteers could enhance ASM responses to TLR agonists. PBMC were enriched from leukocytes by density gradient centrifugation. PBMC alone, although capable of generating the cytokines studied, did not cause detectable cytokine generation when stimulated with TLR agonists, because they were present at too low a number (Figure 4)

. In parallel, PBMC were added to wells containing ASM, and these cocultures were stimulated with medium, Pam3CSK4, or pLPS. The addition of PBMC to ASM resulted in additional generation of CXCL8 and IL-6 from the cocultures, even without costimulation with TLR agonists. The addition of either TLR agonist to the cocultures resulted in a marked synergistic increase in cytokine generation (Figure 4). This was particularly evident with pLPS. Costimulation of ASM with pLPS in the 24-well plates (containing ∼ 48,000 ASM/well, manufacturer's data), in the presence of just 100 PBMC, resulted in a greater induction of CXCL8 and IL-6 than seen in the absence of PBMC. In contrast, generation of CCL5 was virtually undetectable in the stimulated cocultures (data not shown).

Cooperation between PBMC and ASM Is Inhibited by IL-1 Receptor Antagonist

To determine the mechanism resulting in synergistic induction of ASM cytokine generation by TLR agonists and PBMC, cocultures were treated with antagonists of likely cytokines involved in the cooperative response. Pretreatment of cocultures with soluble TNF receptor 1 (sTNFR1) (Figure 5)

or antagonists of NO generation (NG-methyl-l-arginine [LNMA]; data not shown) did not inhibit PBMC-dependent, TLR-induced cytokine generation from ASM. In contrast, IL-1 receptor antagonist (IL-1Ra) pretreatment of the cocultures resulted in effective inhibition of TLR2- or TLR4-mediated induction of cytokine generation at three different PBMC densities (Figure 5).

IL-1 Is Necessary but Not Sufficient for the Activation of ASM

The addition of exogenous IL-1β also resulted in a concentration-dependent generation of CXCL8 from ASM (Figure 6A)

. To investigate the role of IL-1 in more detail, PBMC (1 × 106/ml) were stimulated for 24 hours with media alone or pLPS, and the resulting supernatants were harvested and stored for later addition to ASM monocultures. Supernatants from unstimulated PBMC did not activate ASM (Figure 6B), and contained undetectable levels of IL-1β by ELISA. Supernatants from pLPS-stimulated PBMC contained IL-1β (3.5 ± 0.63 ng/ml, n = 4 ± SEM) and, when applied diluted 1:4 to ASM, resulted in a major induction of CXCL8 release that was again blocked by IL-1Ra (Figure 6B). However, cytokine induction from these stimulated ASM cultures was many-fold greater than explicable on the basis of the IL-1 concentrations in the supernatant, implying that IL-1β was necessary to achieve activation of ASM, but that other mediators present in the supernatants from activated PBMC played an important role in inducing cytokine generation.

Preparations Enriched for Monocytes, but Not T Cells, Support Coculture Responses to pLPS

To determine the cell type or types within PBMC responsible for synergistic interactions with ASM, we studied cells further purified by negative magnetic selection. Starting populations of gradient-enriched PBMC contained a mean ± SEM (n = 4) of 42 ± 5.8% CD3+ T cells and 5.6 ± 1.04% CD14+ cells. T-cell preparations further enriched by negative magnetic selection contained a mean ± SEM (n = 4) of 92 ± 1.7% CD3+ T cells and 0.7 ± 0.3% CD14+ cells, whereas enriched monocytes contained a mean ± SEM (n = 4) of 0.2 ± 0.1% CD3+ T cells and 26.2 ± 5% CD14+ cells (mean fold enrichment vs. starting PBMC was 2.3 for CD3+ T cells and 5.2 for CD14+ monocytes). When stimulated with pLPS at high concentrations (1 × 106/ml), CXCL8 generation was significantly greater from monocultures of PBMC and enriched monocytes than from T cells (Figure 7A)

. In cocultures where low numbers of each cell type were added to ASM and stimulated with pLPS, purified T cells (10,000/well = 25,000 cells/ml) supported only minimal activation of the coculture, whereas enriched monocytes (1,000 cells/well = 2,500 cells/ml) resulted in similar activation to that seen in wells containing 10,000 PBMC.

Local Release of IL-1β Is Likely to Be Important in Initiating ASM Responses

IL-1β is released in microvesicles from monocytic cells (31). We hypothesized that, in our cocultures where PBMC are present at low numbers, close contact of PBMC and ASM would be necessary to achieve full cell activation, through the generation of high concentrations of mediators, such as IL-1β, directly adjacent to the ASM. Figure 8

shows that, where separation of PBMC from ASM was achieved using a cell culture insert with a 0.4-μm pore size, a substantially greater number of PBMC were needed to support coculture activation. Again, our data showed substantial inhibition of coculture activation in the presence of IL-1Ra.

PBMC/ASM Cocultures Result in Cytokine Generation That May Amplify PBMC Recruitment

The coculture of ASM with PBMC and TLR agonists resulted in generation of cytokines, such as CXCL8, that could enable neutrophil recruitment. In further experiments, we found that these cocultures also generated CCL2 (monocyte chemotactic protein 1 [MCP-1]), thus providing a pathway that could result in further monocyte recruitment in vivo, and therefore further enhancement of the ASM/PBMC cooperative response. Generation of CCL2 in these cocultures was also largely inhibited by pretreatment with IL-1Ra (Figure 9)


TLR2 and TLR4 have been identified only recently, but it is clear from recent studies, and knowledge of the biology of their principal agonists, that their contribution to respiratory disease will be significant. Their most important roles are probably in mediating responses to bacterial lipoproteins and lipoteichoic acids (TLR2) and LPS (TLR4) (15, 22, 23, 32), but endogenous molecules associated with inflammation and cell damage may also activate these receptors. For example, the transcription factor high-mobility group box 1, or HMGB1, may activate either receptor if released extracellularly during inflammation (33), and TLR4 responds to tissue breakdown products, such as hyaluronan oligosaccharides (8), and to proteins present at inflammatory sites, such as fibrinogen (7). (The recent realization that activation of TLR4 induced by heat shock proteins 60 and 70 preparations is perhaps caused by LPS contamination [34, 35] provides a cautionary note when considering the biology of this receptor.) In addition to their obvious role in leukocyte activation, TLRs 2 and 4 have also been shown to mediate activation of cardiac myocytes (36, 37), endothelial cells (38, 39), and pulmonary epithelial cells (5, 40). We therefore investigated the potential for these receptors to influence the activation of primary human ASM. We observed a striking synergy between PBMC and ASM, driven by TLR signals, that has the potential to be of considerable biological significance.

In contrast to data generated studying cardiac myocytes (36, 37), we could find little evidence of a significant role for TLR2 and TLR4 in the proinflammatory function of isolated ASM cells. The mRNAs for these receptors were expressed, but compared with actions of IL-1β and TNF-α, responses to cLPS were an order of magnitude smaller (Figure 1), and upregulation of TLR mRNAs by these cytokines was again of modest levels compared with effects on CXCL8 mRNA. ASM are starved of serum to arrest growth of the cells, but serum provides sCD14 and LPS binding protein (LBP), which play important roles in enhancing LPS responses. In these experiments, however, the presence of serum did not allow ASM to respond to TLR agonists. TLRs can be functional when expressed at very low numbers (41), and there was some evidence that ASM responses to TLR2 could be primed by TNF-α (Figure 3). Even this effect was relatively modest and consistent with a cell type with only minimal responses to TLR2 and TLR4 agonists. Over the course of the experiments, ASM from three different subjects were used, suggesting that these data are likely to be broadly applicable. We cannot exclude the possibility that in vivo cells may show some alteration of phenotype, and particularly in diseases of long duration, such as asthma and chronic obstructive pulmonary disease, it is possible that functional TLR expression may become apparent on ASM. However, our data would suggest that, even if such upregulation of function should occur, the effects of TLR2 and TLR4 agonists on ASM function would be dominated by the synergy observed between PBMC and ASM.

In further experiments, low numbers of PBMC (range, 100–30,000 cells/well = 250–75,000 cells/ml) were added to ASM cultures (48,000 cells/well, manufacturer's data). Even very low numbers of PBMC resulted in a dramatic enhancement of the ability of TLR agonists to cause the generation of inflammatory cytokines from these cocultures (Figures 4, 5, and 7). The levels of CXCL8 observed were similar to those seen from other groups when ASM were directly activated by proinflammatory cytokines (27, 42). PBMC themselves can make the cytokines studied here (16, 24), but the numbers of cells per well (30,000 PBMC/well [i.e., 75,000 cells/ml]) were not sufficient to result in detectable cytokine when cultured with TLR agonists in the absence of ASM. PBMC contain several TLR-expressing cell types that might contribute to coculture effects, such as dendritic cells (41) and monocytes, and there is an increasing realization that TLR expression occurs on some lymphocyte populations (43, 44). Monocytes make significant quantities of IL-1β in response to TLR agonists, are relatively numerous, and are recruited actively to sites of pulmonary inflammation (45). In other studies, we have found that PBMC markedly enhance neutrophil responses to LPS, and we and others have shown that removal of contaminating CD14+ cells (which are principally monocytes) from cell preparations reduces LPS-mediated granulocyte survival (25, 30). In keeping with these published results, we observed that purified T cells had only a very limited capacity to support activation of the cocultures, whereas enriched monocytes showed enhanced coculture activation compared with PBMC. Furthermore, CCL2 (MCP-1) is not released by LPS-stimulated ASM (12) but was induced from the ASM/ PBMC cocultures, providing a mechanism for further recruitment of monocytes in vivo and therefore amplification of this inflammatory cascade. Chemokines, such as CXCL8 and CCL2, generated in the coculture will favor leukocyte recruitment, amplifying innate immune responses that could contribute to diseases, such as chronic obstructive pulmonary disease, asthma, and acute respiratory distress syndrome. Recent data have shown that IL-6 generation can overcome the inhibitory effects of T-regulatory cells on allergen-specific T-cell proliferation, facilitating adaptive immunity (46). Thus, the elaboration of IL-6 by these cocultures provides TLR- and monocyte-dependent mechanisms enhancing not just innate but also adaptive immunity, with clear relevance to the amplification of inflammation in diseases such as asthma. Of note, monocyte–smooth muscle interactions have been observed previously concerning elaboration of proteins causing destruction of tissue matrix (47), which could release further TLR4-agonistic moieties.

We were also able to define the molecular basis for the cooperativity seen between ASM and PBMC, because almost all the cytokine generation in the cocultures was inhibited by pretreatment with IL-1Ra. IL-1Ra did not ablate direct induction of cytokine release from ASM stimulated with TNF-α (data not shown), and when high numbers of PBMC were present in cocultures but separated by a transwell (Figure 8), IL-1Ra was a less efficient antagonist of cytokine production. PBMC cultured at 1 × 106/ml in the absence of ASM produced CXCL8 when activated with pLPS, which was only modestly inhibited by IL-1Ra (CXCL8 from pLPS-stimulated PBMC = 109 ± 28 ng/ml [mean, n = 4 ± SEM] vs. pLPS-stimulated cells in the presence of IL-1Ra = 76 ± 14 ng/ml; p = 0.04, paired t test). These data suggest that IL-1Ra did not result in a global functional paresis of PBMC or ASM, and that IL-1β is important particularly when released in close proximity to ASM. These data fit with the range of known important effects of IL-1β on ASM, including its potent ability to cause CXCL8 generation (27, 42, 4850), although we saw little CCL5 generation in the cocultures, which is in contrast to data showing generation of this cytokine by IL-1β–stimulated ASM (11). Furthermore, the much greater efficacy of pLPS versus Pam3CSK4 in the induction of cytokine generation in the cocultures parallels the relative abilities of these agonists to induce the release of IL-1β from PBMC (24). It is interesting to note that a similar interaction was recently observed between dermal epithelial cells, PBMC, and LPS (51), which was also found to be IL-1–dependent. The reduction in efficiency of activation of the cocultures when PBMC were separated from ASM by transwells could also be consistent with a role for cell:cell contact in induction of responses, as has been observed between ASM and activated T cells (52), although our studies place secreted IL-1β as a central mediator of coculture activation. Our data also reveal that amounts of IL-1β as measured by ELISA in supernatants from pLPS-stimulated PBMC were clearly unable to account for the total CXCL8 generated when the supernatants were applied to ASM. These data suggest that IL-1β is a crucial permissive factor in the induction of inflammation, but that other monocyte-derived mediators also play major roles in amplifying cytokine generation. Synergistic interactions between IL-1β and Th2-type cytokines resulting in enhanced generation of cytokines from ASM have been demonstrated previously (11). Monocyte–epithelial interactions in the lung have been shown to be at least in part dependent on cell–cell contact signaling mediated by intercellular adhesion molecule 1 (53). However, in asthma and chronic obstructive pulmonary disease, the submucosal leukocytic infiltration is in close proximity to ASM, but there is little evidence for direct infiltration of the ASM by the leukocytes themselves, favoring local release of cytokines as a likely mechanism of cooperativity. In keeping with this, IL-1Ra can decrease neutrophil infiltration and lung damage in inflammatory models in vivo (54).

In summary, we found that primary ASM cells do not appear to express significant levels of functional TLR2 and TLR4. However, these highly IL-1–responsive cells generate marked levels of biologically relevant cytokines when stimulated with TLR agonists in the presence of very low numbers of PBMC, a cooperativity that was blocked by IL-1Ra. Monocytes have crucial roles as the initiators and regulators of inflammation, and their interaction with tissue cells, such as ASM, provides a major mechanism by which their actions may be amplified effectively.

The authors thank Dr. Lynne Prince, Ms. Elizabeth Jones, and Dr. Stephen Bianchi for help in preparation of PBMC, and Dr. Lisa Parker for assistance with the Western blotting.

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Correspondence and requests for reprints should be addressed to Ian Sabroe, Ph.D., Academic Unit of Respiratory Medicine, Division of Genomic Medicine, University of Sheffield, M Floor, Royal Hallamshire Hospital, Sheffield, S10 2JF, UK. E-mail:


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