Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2013 Apr 29.
Published in final edited form as: J Immunol. 2010 Sep 1;185(7):3884–3894. doi: 10.4049/jimmunol.1000387

Surfactant Protein-A Inhibits Mycoplasma-Induced Dendritic Cell Maturation through Regulation of HMGB-1 Cytokine Activity

JG Ledford *, B Lo *, MM Kislan *, JM Thomas *, K Evans *, DW Cain †, M Kraft ‡, KL Williams *,†, JR Wright *
PMCID: PMC3638720  NIHMSID: NIHMS455123  PMID: 20810986

Abstract

During pulmonary infections, a careful balance between activation of protective host defense mechanisms and potentially injurious inflammatory processes must be maintained. Surfactant protein A (SP-A) is an immune modulator that increases pathogen uptake and clearance by phagocytes while minimizing lung inflammation by limiting dendritic cell (DC) and T cell activation. Recent publications have shown that SP-A binds to and is bacteriostatic for Mycoplasma pneumoniae (Mp) in vitro. In vivo, SP-A aids in maintenance of airway homeostasis during Mp pulmonary infection by preventing an overzealous pro-inflammatory response mediated by TNF-α. While SP-A has been shown to inhibit maturation of dendritic cells in vitro, the consequence of DC/SP-A interactions in vivo has not been elucidated. Here we show that the absence of SP-A during Mp infection leads to increased numbers of mature DCs in the lung and draining lymph nodes during the acute phase of infection and consequently, increased numbers of activated T and B cells during the course of infection. The findings that glycyrrhizin, a specific inhibitor of extracellular high-mobility group box-1 (HMGB-1) abrogated this effect and that SP-A inhibits HMGB-1 release from immune cells suggest that SP-A inhibits Mp-induced DC maturation by regulating HMGB-1 cytokine activity.

INTRODUCTION

Mycoplasma pneumoniae is recognized as one of the most common causes of community acquired pneumonia and greater than 50% of chronic stable asthmatics have evidence of airway infection with M. pneumoniae (1, 2). Mp are atypical bacteria that form strong attachments to ciliated airway epithelial cells where they release oxidative products that can cause airway tissue damage and contribute to exacerbations in chronic asthmatics (3). Infections with Mp may persist, with mild symptoms, for several weeks with manifestations in the upper, as well as the lower respiratory tract.

Because Mp is primarily an extracellular pathogen that invades and resides in the respiratory tract, it has the potential to encounter pulmonary surfactant proteins that are produced by alveolar type II cells, Clara cells and submucosal glands of the respiratory tract. Indeed, studies have demonstrated that surfactant protein A (SP-A) binds Mp though disaturated phosphatidylglycerols and through a specific surface binding protein, MPN372 (4, 5) and limits the growth of Mp in vitro (5). SP-A also helps maintain airway homeostasis and reduce hyperresponsiveness by curtailing an overly-ambitious pro-inflammatory immune response during Mp infection in mice in vivo (6).

Several immune functions have been ascribed to SP-A including inhibition of T cell proliferation, augmentation of pathogen phagocytosis by acting as an opsonin, and modulation of chemotaxis and cytokine production (reviewed in (7)). A further role for SP-A has been established in mediating adaptive immune responses through interactions with DCs. For example, SP-A binds to DCs and negatively regulates their maturation in vitro thereby reducing their T cell allostimulatory ability (8). The consequences of this interaction during an infection in vivo, as well as the mechanism by which SP-A modulates DC functional maturation have not been defined. Therefore, using mice deficient in SP-A, we tested the hypothesis that SP-A regulates recruitment, activation and maturation of adaptive immune cells in response to Mp by regulating expression of the endogenous stress factor HMGB-1, which, if released in the context of infection, can activate DCs and lead to their maturation (9).

We report here that Mp infection leads to increased numbers of exudative macrophages and DCs in the lung parenchyma, a response that is augmented by the absence of SP-A. Likewise, the total number and activation state of DCs that have migrated to the mediastinal draining lymph nodes (MLN) during the acute phase (three days) of infection is also increased in the absence of SP-A. Additionally, elevated numbers of activated T and B cells in both the lungs and MLN, as well as Mp-specific IgG in the serum, are observed in mice lacking SP-A nine days after Mp infection. Treatment with glycyrrhizin, a specific extra-cellular inhibitor of the potent pro-inflammatory cytokine HMGB-1, protects the DCs in the MLNs of mice lacking SP-A from Mp-induced maturation, suggesting that SP-A inhibits Mp-induced DC maturation by regulating HMGB-1 cytokine activity.

MATERIALS AND METHODS

M. pneumoniae culture

Mp from American Type Culture Collection (ATCC; cat. # 15531) was grown in SP4 broth (Remel) at 35°C until adherent. Mp concentration was determined by plating serial dilutions of Mp on PPLO agar plates (Remel). Colonies were counted under 10× magnification on plates after incubation for 14 days. For in vivo infection, adherent Mp were washed by centrifuging at 6000 rpm for 5 minutes and resuspended in sterile saline for infection at a concentration of 1 × 108 Mp/50 μl inoculum. Mp burden was assessed as previously described by plating BAL or by RT-PCR using primers against Mp-specific P1-adhesin gene relative to the housekeeper cyclophilin (6).

Mice

An inbred strain of SP-A deficient mice was generated by disrupting the murine gene encoding SP-A by homologous recombination as previously described (10). SP-A null mice were backcrossed for 12 generations onto the C57BL/6 background which were purchased from Charles River. Wild type C57BL/6 mice were purchased from Charles River and bred in-house to account for any possible effects of environmental conditions. All mice used in experiments were age (8–12 weeks) and sex (males) matched. Protocols were approved by the Institutional Animal Care and Use Committee (IACUC) at Duke University.

Mice 8–12 weeks of age were anesthetized via i.p. injections of 10 μl/g body weight of a 12% Ketamine (100 mg/ml) and 5% Xylazine (20 mg/ml) mix. Mice were infected with either 50 μl of sterile saline or 50 μl of ~1 × 108 Mp units in sterile saline by intranasal instillation. Some groups of mice received i.p injections of glycyrrhizin (10 mg/kg body weight) 2 hours prior to Mp infection and 24 hours post-infection, in order to neutralize HMGB-1 cytokine activity as previously described (11).

BAL collection, lung digestion and analysis of pulmonary dendritic cells

The lungs of mice were perfused with 10 ml of PBS and then lavaged with PBS containing 0.1 mM EDTA (warmed to 37°C). Analysis of cytokines and chemokines present in the cell free BAL of infected and uninfected mice was carried out by multiplex cytokine analysis (Luminex technology, Invitrogen). Lungs were removed and minced with a razor blade and resuspended in 5 ml of HBSS (containing calcium and magnesium) with 1.0 mg/ml collagenase A and 0.2 mg/ml DNAase I. The cell suspensions were incubated for 1 hour with shaking (200 rpm) at 37°C for enzymatic digestion to occur. Lung digests were then filtered through 40 μm strainers. Remaining red blood cells in the digests or BAL were lysed with Gey's lysis solution (0.83% NH4Cl, 0.1% KHCO3). The suspended cells were layered on top of a 4.0% solution of iodixanol (Optiprep™; Axis-Shield), placed above a 14.5% iodixanol solution, and centrifuged at 600 × g (no brake) for 20 min at room temperature. Low-density cells were isolated from the 4–14.5% interface and used for FACS analysis. Cells were tri-stained with APC-labeled anti-CD11c, FITC- (or PE-) labeled anti-MHC II, and either PE-anti-CD80 or PE-anti-CD86.

Mediastinal lymph node digests and dendritic cell analysis

Mediastinal lymph nodes collected from each mouse were placed in individual wells of a 6-well plate containing 5 ml of PBS with 5% FCS, 1.0 mg/ml collagenase A, and 0.2 mg/ml DNAase I. Lymph nodes were minced using a scalpel and then incubated at 37°C for ~35 min. Digestion was stopped by adding 1 ml of 120 mM EDTA in PBS and incubating another 5 min at room temperature. Lymph node digests were pushed through a 40 μm strainer to obtain single cell suspensions. Some lymph node digests were centrifuged through a density gradient (Nycodenz, Axis-Shield) to enrich for DCs. Red blood cells were lysed with Gey's solution (0.83% NH4Cl, 0.1% KHCO3). Cells were then resuspended in HBSS with 5% FCS, 2 mM EDTA, and 100 U/ml penicillin–streptomycin and stained for FACS analysis. Cells were tri-stained with APC-labeled anti-CD11c, FITC- labeled anti-MHC II, PE-labeled anti-CD86, and PE-Texas Red labeled anti-CD80.

Flow cytometry analysis

Flow cytometry was performed in the Duke Human Vaccine Institute Flow Cytometry Core Facility, which is supported by the National Institutes of Health award AI-51445. Initially, cells were examined by forward-scatter (FSC) versus side-scatter (SSC) to separate those smaller non-granular lymphoid populations from the larger granular myeloid populations. The percentage of cells within each of these gates was used to calculate the total number of lymphoid cells and myeloid cells examined from the total number of cells obtained from the lung digests as counted on a hemacytometer. Based on similar gating strategies published by Lin et al. and Pecora et al., alveolar macrophages were defined as CD11bneg-low/CD11cmid/MHCIImid, DCs were defined as CD11bhigh/CD11chigh/MHCIIhigh and inflammatory lung macrophages (also known as exudate macrophages) CD11bhigh/CD11cneg-mid/MHCIImid (12, 13).

Generation of bone marrow-derived dendritic cells (BMDCs)

BMDCs were generated as described previously by Inaba et al. (14) and as modified by Brinker et al. (15). Briefly, marrow from the tibia, femur and humerus of mice were harvested, cultured in RPMI 1640 supplemented with 5% FCS, antibiotics, and 50 μM 2-mercaptoethanol plus 5% GM-CSF conditioned media for 6 days. Loosely attached cells were harvested and negatively selected with biotinylated-Gr-1 antibodies (Pharmingen) and streptavidin paramagnetic microbeads (Miltenyi). Dendritic cells were matured for 24 hours in 24-well plates at 5 × 105 cells/ml in the presence of 100 ng/ml LPS (serotype 055:B5) as previously described (8).

Cell culture stimulation experiments

Undifferentiated THP-1 cells (ATTC) were maintained in RPMI 1640 with supplements (10% fetal calf serum, 2 mM L-glutamine, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, and streptomycin-penicillin) at 37°C at 5% CO2. Clonetics® normal human bronchial epithelial cells (NHBE) purchased from Lonza and grown to approximately 80% confluence in bronchial epithelial growth media (Clonetics®) with supplements and growth factors (BPE, hydrocortisone, hEGF, epinephrine, insulin, triiodothyronine, transferrin, gentamicin/amphotericin-B and retinoic acid) at which point cells were placed in bronchial epithelial basal medium (no supplements) for stimulation experiments. THP-1 cells were resuspended in RPMI 1640 (no supplements). DC, NHBE and THP-1 cells were resuspended in the respective medium in the presence or absence of purified human SP-A (50 μg/ml), which was purified as described in published methods (16) and approximately 5 × 105 cells were seeded per well into 12-well plates. After 2 hrs, Mp was added at a concentration of 10 Mp cfu/cell (5 × 106/well) to some wells, while others received saline as a control buffer for the Mp. Some sample wells received neutralizing anti-TLR2 andibody (eBioscience, #16-9024) or isotype control prior to stimulation. After Mp stimulation for approximately 16 hrs, the cell-free supernatant was collected and pelleted cells were lysed for Western analysis.

Western blot analysis

Cell-free lavages from uninfected and Mp-infected WT and SPA−/− mice, stimulated DC, THP-1 or NHBE cell-free supernatants, and DC, THP-1 or NHBE lysates were denatured by heating at 95°C for 5 min in buffer containing DTT prior to gel electrophoresis on 12.5% polyacrylamide gels (Protogel). For cell-free BAL analysis, 25 μl from 1 ml lavages was loaded for each sample. For the DC, THP-1 and NHBE cell-free supernatants, approximately 25 μl from a total well volume of 250 μl was loaded for each sample and 25 μl was loaded from 250 μl of the cell lysates. Gels were transferred onto Nitrocellulose membrane and blocked with 5% block (milk in TBS) and labeled with an HMGB-1 antibody (Abcam, ab12029 or ab65003) overnight at 1:1500. The secondary antibody, HRP-conjugated anti-mouse IgG or anti-rabbit IgG (Cell Signaling), was used at 1:10,000 for 1 hr. The SP-A antibody used was rabbit anti-sheep SP-A (cross reacts with mouse SP-A), which was used with anti-rabbit HRP secondary. GAPDH antibody was purchased from Millipore (MAB374). SuperSignal West Fempto (Thermo Scientific) was used to visualize protein expression.

Cytokine/chemokine and IgG analysis

Cytokines and chemokines were analyzed by Luminex technologies using a mouse 20-plex kit (Invitrogen) and GM-CSF was analyzed by ELISA (eBioscience) according to manufacturer's instructions. Mp-specific IgG was analyzed by ELISA as described previously (17). Briefly, sonicated heat-killed Mp dissolved in coating buffer (0.05M Carbonate-bicarbonate buffer, pH10) was used at a concentration of 10 μg/ml and was incubated overnight at 4°C. Wells were then blocked for 1 hr with 2% BSA/PBS/0.05% Tween-20 mix after which time serum samples were diluted 1:10,000 in 1% BSA/PBS buffer and added from uninfected and Mp-infected mice. A biotin-goat anti-mouse IgG antibody (Jackson ImmunoResearch) and streptavidin-HRP (BDPharmingen) were used for the detection. Samples were analyzed at 450 nm absorbance on the FLUOstar Optima plate reader (BMG Labtech).

Microscope Image Acquisition

Images of the lung were taken on a Nikon Eclipse 50i light microscope at either 10× (aperture 0.3) or 40× (aperture 0.75) at room temperature by digital photography of bright field images using a Nikon Infinity 2 camera. Infinity Capture software was used for image acquisition and Adobe Photoshop was used for color contrast settings and figure presentation.

Statistical analysis

All data measurements were analyzed with PRISM (GraphPad) software, first to determine if data were normally distributed, followed by t-test to determine significance. Data sets with significant variance between comparison groups were analyzed by t-test using Welch's correction as assessed in PRISM. *p<.05 and **<.01 unless otherwise noted.

RESULTS

Increased numbers of antigen presenting cells in lungs of SP-A null mice during Mp infection

In order to assess the role of SP-A in regulating immune cell responses in vivo, the cells present in the lungs due to Mp infection and the resulting inflammation were analyzed in WT and SP-A null mice. Although the total numbers of myeloid cells present in the lung digests were not significantly different in the SP-A null infected mice as compared to WT infected mice, the cellular composition, which was analyzed by flow cytometry, showed differences in certain cell populations. Inflammatory monocytes and PMNs increased in number in response to Mp challenge, although levels were equivalent in the lungs of both WT and SP-A null mice (data not shown). Among the other cell populations, dendritic cells were examined from the lung digests of infected and uninfected mice. Lung DCs were defined based on their pattern of side scatter (SSC) versus forward scatter (FSC) and high levels of expression of both CD11c and MHC II cell surface markers (12, 18). Very few DCs (<1% of total cells) were present in the lung digests of the saline treated mice (Fig. 1A). After Mp infection, the number of DCs observed in WT mice was only slightly elevated over those observed in the saline treated. However, the number of cells observed within the DC gate was significantly increased in the SPA null mice (Fig. 1A).

Figure 1. Analysis of antigen presenting cells in the lungs.

Figure 1

Open in a new tab

Mice were instilled with 1 × 108 Mp and 72 hours post infection samples were collected. Lungs were perfused and cells harvested from lung tissue after enzymatic digest and density gradient centrifugation. Cells were labeled with fluorescent antibodies against cell surface markers and analyzed by flow cytometry. A, Dendritic cells (DCs) were identified as MHCIIhi CD11c+ cells and B, Macrophages were identified as MHCIIint CD11c+ cells by flow cytometry. Total numbers of dendritic cells, macrophages and CD11b+ macrophages were determined by multiplying the percentage of each respective cell population by hemacytometer counts per sample. Results shown are the mean ± sem from three independent experiments, n=12/group, **p<.01, *p<.05.

The numbers of the macrophages (MHCIIint, CD11c+) isolated from lungs of saline treated mice were not significantly different between the WT control and the SP-A null mice. However, 72 hours after Mp infection there were significantly more macrophages (including resident and inflammatory) in both groups of infected mice as compared to saline treated animals in each group (Fig. 1B). Whereas the number of macrophages increased in the WT infected mice by two-fold, the number of macrophages in the lungs of the infected SP-A null mice increased almost four-fold over their saline controls and was significantly greater in comparison to the WT infected mice (*p<0.01).

Expression of the cell surface marker CD11b was also assessed on macrophage populations in the Mp infected mice in order to differentiate inflammatory newly-recruited macrophages (CD11b+), also known as exudate macrophages, that had migrated into the lung during pulmonary infection from the resident alveolar macrophages (CD11b−) (13, 19). Similar to the alveolar macrophages analyzed in the bronchoalveolar lavages (BAL) of saline treated mice, very few of the tissue macrophages expressed CD11b in the saline treated mice suggesting those were primarily resident macrophages (Fig. 1B). However, CD11b expression was significantly elevated on macrophages in both the WT and SP-A null mice after Mp infection suggesting these were predominantly newly recruited or differentiated inflammatory macrophages. Additionally, there were significantly more CD11b+ macrophages in Mp-infected mice deficient in SP-A as compared to infected WT mice, suggesting that SP-A is important in inhibiting the influx of these inflammatory macrophages into the lung during Mp infection.

Increased chemoattractant signals in BAL during Mp infection in the absence of SP-A

The function of the pulmonary DC network is altered dramatically during inflammatory conditions due to signals produced by pulmonary epithelium and myofibroblasts of the airways that attract immature DCs from the bloodstream into areas of pulmonary challenge (20). To determine if any such chemokine or cytokine mediators were differentially expressed in response to Mp infection in the absence of SP-A, BAL fluid was harvested after 3 days of infection and analyzed. As shown in Table I, neither monocyte chemotactic protein-1 (MCP-1) nor macrophage inflammatory protein-1α (MIP-1α), factors known to be chemotactic for immature DCs (21, 22), were detectable in BAL fluid of saline treated mice. However, MCP-1 was detected at 15.6 pg/ml (not detectable in WT infected) and MIP-1α at 21.9 pg/ml (compared to 9.7 pg/ml in WT infected mice) in BAL from Mp infected animals lacking SP-A. The amount of GM-CSF, a factor known to enhance the differentiation of monocytes into immunostimulatory DCs in the lung vasculature, was increased in BAL from both WT and SP-A null Mp infected mice as compared to levels detected in untreated control sample.

Increased T-lymphocytes in Mp infected SP-A null mice

Analysis of cell surface markers from the lymphoid panel revealed a dramatic influx of CD3+ lymphocytes (mean=1.6 × 106 per total lung cells) into the lung tissue of SP-A−/− mice after three days of Mp infection in comparison to both their respective saline controls (**p<0.001) or WT infected mice (**p<0.001) (Fig. 2A). Interestingly, the number of CD3+ lymphocytes present in the WT Mp infected mice at day three (mean=0.7 × 106) was not significantly elevated over non-infected mice (mean=0.7 × 106). T-lymphocytes were also analyzed nine days after infection to determine if the inflammatory response was diminished, maintained or augmented over levels detected after three days of infection. Despite no significant findings of T cell infiltration in WT mice three days after infection, they did present with significantly enhanced numbers of CD3+ T lymphocytes in the lungs nine days after infection as compared with non-infected controls (Fig. 2A). Additionally, CD3+ T cells detected in the lungs of SP-A null mice after nine days of infection were significantly increased over the numbers present at three days of infection in mice lacking SP-A (**p<0.001), as well as over the numbers present in the WT mice after nine days of infection (**p<0.001). Interestingly, the T-cell chemoattractant, monokine induce by gamma interferon (MIG), was also significantly increased in Mp-infected mice lacking SP-A as compared to infected WT mice (Table I).

Figure 2. Increases in T lymphocytes in the lungs and mediastinal lymph nodes of Mp challenged mice.

Figure 2

Open in a new tab

WT and SP-A−/− mice were instilled with 1 × 108 Mp and 3 or 9 days post infection samples were collected. Lungs were perfused and cells harvested from lung tissue or MLNs after enzymatic digest and density gradient centrifugation. Cells were labeled with fluorescent antibodies against cell surface markers and analyzed by flow cytometry. A, CD3+ T cells and B, CD3+CD4+, and CD3+CD8+ cells detected by flow cytometry from the total lung cell digest at the indicated times. C, The activation status of CD3+CD4+ and CD3+CD8+ T cells from the total lung cell digest at day 9 as determined by the presence of CD25 and CD69 by flow cytometry. D, Histological representation taken at 10× and 40× magnification of lymphocytic foci in infected lungs from WT and SP-A−/− mice. E, The total number of CD3+CD4+ and CD3+CD8+ T cells harvested from the MLNs after 9 days of infection. F, The activation status of CD3+CD4+ and CD3+CD8+ T cells from MLN digest as determined by the presence of CD25 and CD69 by flow cytometry. Results shown are the mean ± sem from three independent experiments, n=12/group, **p<.01, *p<.05.

Those cells expressing CD3 were further analyzed by flow cytometry for their surface expression of CD4 and CD8. Mp-infected SP-A−/− mice had significantly elevated numbers of CD3+ CD4+, as well as CD3+ CD8+ lymphocytes compared to WT Mp-infected mice (Fig. 2B). The cell surface expression of CD25, which plays dual roles in lymphocyte differentiation and activation/proliferation, on CD4+ T cells was also significantly higher in the SP-A−/− Mp infected mice as compared to infected control mice (data not shown).

In order to determine the activation status of those T lymphocytes present in the lung after nine days of infection, we analyzed CD69 expression, a very early activation antigen, on CD3+CD4+ and CD3+CD8+ T cells by flow cytometry. CD69 was not upregulated in WT mice nine days after infection on either CD4+ or CD8+ populations. However, there was significant expression of CD69 on both CD4+ and CD8+ cells in the SP-A null mice as compared to WT infected mice and to saline controls (Fig. 2C). In support of these findings, expression of IL-12, which is involved in the differentiation of T cells and is associated with T cell activation, was also significantly increased in the BAL of infected SP-A mice as compared to infected WT mice (Table I).

T cells present in the MLN after 9 days of Mp infection were also examined. There were no appreciable increases in the total number of CD4+ or CD8+ T cells recovered in MLN from the infected WT mice. However, in Mp infected SP-A deficient mice, there were significantly more CD4+ and CD8+ T cells as compared to WT infected (Fig. 2E). The state of T cell activation was determined by the presence of CD25 and CD69 on the cell surface. The numbers of CD8+ activated cells were quite low in all groups of mice examined (Fig. 2F). However, as shown in Fig. 2F, the numbers of CD4+ T cells expressing these activation markers were significantly increased in the MLN of infected SP-A deficient over infected WT mice, as well as saline control mice.

Increased Mp burden does not correlate with increased T lymphocytes

Mycoplasma burden in SP-A deficient mice has previously been described in our studies after 3 days of infection, which showed that while there was no difference in Mp burden from BALs at day 3 of infection, significantly more Mp were colonizing the airway by binding to epithelial cells (6). While both wild type and SP-A deficient mice were clearing Mp by day 9, interestingly, only 20% of WT mice tested positive for Mp in the BAL compared to greater than 80% of the SP-A deficient mice (Fig. 3A). Likewise, Mp burden in association with lung tissue as determined by RT-PCR was also significantly increased in SP-A deficient mice at day 9 of infection (Fig. 3B). Although some mycoplasmas are believed to be mitogenic for T cells, we did not find a correlation between Mp burden and T cell number in our system (Fig. 3C).

Figure 3. Mp burden in WT and SP-A−/− infected mice.

Figure 3

Open in a new tab

WT and SP-A−/− mice were infected with 1 × 108 Mp intranasally and the Mp burden assessed after 9 days from A, BAL plated on PPLO agar or B, Lung tissue by RT-PCR for the Mp specific P1-Adhesin gene relative to the housekeeper gene, cyclophilin. Data is from 2 experiments combined with n=8 per group, *p<.05, **p<.01. C, WT and SP-A−/− mice were infected with 1 × 108 Mp intranasally and after 3 days the Mp burden was determined in the lung tissue by RT-PCR of the Mp specific P1-adhesin gene relative to the housekeeper gene, cyclophilin and the number of CD3+ T cells were determined by flow cytometry using specific cell surface markers. Data are from 2 experiments combined and were statistically assessed using Prism software.

Heightened B cell responses in Mp-infected SP-A null mice

Cells that expressed the cell surface marker B220 were also significantly increased in mice lacking SP-A, both in the saline treated mice, as well as in infected mice at both three and nine days after infection (Fig. 4A). The number of B cells did not significantly increase in WT mice during the infection. However, B cells present in the lungs of SP-A null mice not only significantly increased over their saline treated controls, but also over each time point examined as compared to WT infected controls (Fig. 4A). Although there were significantly more B220+ cells in saline treated SP-A null mice, the total number of B220+ cells considered to be activated, as determined by coexpression of IgM and CD69, were equivalent in the saline treated groups (Fig. 4B). There was also no increase in the total number of activated cells (B220+ IgM+ CD69+) cells isolated from Mp-infected WT mice after 9 days of infection as compared to their saline controls. In striking contrast, Mp-infected SP-A null mice had significantly greater numbers of activated cells isolated from the lungs after 9 days of infection as compared to their saline controls and to the WT infected mice at the same day of infection (Fig. 4B).

Figure 4. Increased B lymphocytes in lungs and mediastinal lymph nodes of Mp infected mice.

Figure 4

Open in a new tab

WT and SP-A−/− mice were instilled with 1 × 108 Mp and 3 or 9 days post infection samples were collected. Lungs were perfused and cells harvested from lung tissue or MLNs after enzymatic digest and density gradient centrifugation. Cells were labeled with fluorescent antibodies against cell surface markers and analyzed by flow cytometry. A, B220+ cells and B, B220+ IgM+ CD69+ cells from the digested lung tissue and C, B220+ cells and D, B220+ CD69+ cells from the digested MLNs as determined by flow cytometry. E, Mp-specific IgG as determined by ELISA from serum of infected mice shown as the fold over the serum level of non-infected mice. Results shown are the mean + sem from three independent experiments, n=12/group, **p<.01, *p<.05.

B cells were also analyzed from the MLN of infected mice after 9 days of infection. The total numbers of B220+ cells harvested from the MLN were unchanged in infected WT mice as compared to saline treated WT mice. However, significantly more B220+ cells were collected from infected SP-A deficient mice as compared to both saline treated control mice and WT infected mice (Fig. 4C). Additionally, the number of B220+ activated cells were significantly increased in both infected WT and SP-A deficient mice (Fig. 4D). However, in infected SP-A deficient mice, the total number of activated B cells was significantly enhanced over infected WT mice.

Since the Mp-infected SP-A deficient mice displayed a strikingly enhanced B cell response as compared to infected WT mice, we sought to determine if Mp-specific IgG antibodies were produced by day 9 of the pulmonary infection. Serum was collected from WT and SP-A null mice that were Mp-infected and Mp-specific IgG was determined by ELISA. The amount of Mp-specific IgG is expressed as the fold over the baseline levels measured from the serum of non-infected mice. As shown in Fig. 4E, Mp-specific IgG levels were significantly elevated in the serum of infected SP-A null mice as compared to infected WT mice.

More mature DCs in MLNs in Mp infected mice lacking SP-A

Given that increased numbers of activated CD4+ T cells were discovered in the MLNs of infected SP-A deficient mice at the later time point (day 9), we examined the activation state of DCs in the MLNs at an earlier time point (day 3), since their early activation state could directly influence the conversion of naïve T cells into activated T cells at the later time point. WT and SP-A null mice were instilled with Mp or saline and the MLNs were harvested after 3 days. Lymph nodes were enzymatically digested to aid in the release of DCs, a single cell suspension was obtained, and cells were then stained and analyzed by flow cytometry. In the lymph nodes, there are at least 2 populations of CD11c+ cells that can be differentiated. The more mature population expressing higher levels of MHC II and CD86 are predicted to be the DCs that have migrated from the lungs (23, 24). Mp-infected SP-A null mice had a greater percentage of MHC IIhi CD11c+ cells in the mediastinal lymph nodes than infected WT mice (data not shown). The percentage of cells that were MHC IIhi CD11c+, which was determined by gating the flow cytometry measurements, was then used to calculate the total number of MHC IIhiCD11c+ cells based on hemacytometer counts from individual lymph node preparations. As shown in Fig. 5A, the total number of MHC IIhi CD11c+ cells was also significantly greater in infected mice lacking SP-A, suggesting that in the absence of SP-A more DCs in the infected pulmonary environment migrate into the draining lymph nodes. The number of DCs in the MLNs and lungs at day 9 were back to levels observed in saline treated animals and likewise, the lymphocytic inflammation had subsided by day 15 in both WT and SP-A−/− mice (data not shown). Expression of the cell surface markers associated with DC maturation, CD80 and CD86 were also analyzed. Since the total number of MHC IIhi CD11c+ cells was higher in the infected SP-A deficient mice, CD80 and CD86 levels are shown as fold increase in expression within the DC populations. Both CD80 and CD86 cell surface expression were significantly greater on the MHC IIhi CD11c+ cells of the SP-A null mice as compared to levels detected on cells harvested from the infected WT mice (Fig. 5B).

Figure 5. Dendritic cells from mediastinal lymph nodes of M. pneumoniae infected mice 3 days post-infection.

Figure 5

Open in a new tab

A, Total number of MHCIIhi CD11c+ DCs present in the MLNs of uninfected and Mp-infected mice on day 3 of infection. **p<.01, n=8/group, representative of 3 experiments. B, The fold expression, relative to WT Mp-infected, of the maturation markers CD80 and CD86 on the MHC IIhi, CD11c+ DCs. *p<.05, n=15–17/group, combination of 4 experiments. C, BMDCs were cultured for 6 days in the presence of GM-CSF, purified and stimulated with Mp for 16 hours in the presence or absence of SP-A (40 μg/ml). The percentage of DCs undergoing maturation was assessed by flow cytometry for expression of MHC II, CD11c, and CD86. *p<.05, **p<.01, n=3 experiments.

Inhibition of Mp-induced DC maturation by SP-A in vitro

Previous studies from our laboratory demonstrated a role for SP-A in inhibiting LPS-induced DC maturation in vitro (8). Our in vivo studies examining DC maturation in Mp infected SP-A null mice as compared to control WT mice supported a role for SP-A in limiting DC maturation to Mp, as well. To determine if SP-A directly carries out this protective role in response to a clinically relevant pulmonary pathogen, DC maturation in response to Mp was examined in the presence or absence of SP-A. Bone marrow derived dendritic cells were plated and some samples were pre-incubated with SP-A, after which live Mp were added to the DCs and Mp-induced maturation in the presence or absence of SP-A was determined after 24 hours of culture by flow cytometry. MHC IIhi CD11c+ cells were analyzed for expression of the maturation marker, CD86.

As described previously, addition of SP-A to non-stimulated DCs leads to decreased expression of maturation markers as compared to media alone. These results were repeated as controls and are in agreement with published reports (8) (Fig. 5C). Addition of Mp to DCs resulted in increased expression of CD86 as compared to those DCs receiving media alone or those pre-incubated with SP-A alone (no Mp stimulation). Interestingly, those samples that had been pre-incubated with SP-A prior to stimulation with Mp were significantly protected from the up-regulation of the maturation marker CD86 as compared to the Mp-stimulated DCs that were treated with vehicle only (Fig. 5C).

Elevated HMGB-1 expression in Mp-infected SP-A deficient mice

HMGB-1, although most commonly associated with necrotic cells and used as a marker of tissue damage, has more recently been identified as a potent pro-inflammatory cytokine released by activated macrophages and monocytes in the lung during acute inflammation (9, 25, 26). Since HMGB-1 cytokine activity has been shown to be integral for DC maturation and is necessary for antigen-presentation leading to the activation of T cells (27), we sought to determine if levels of HMGB-1 were elevated in Mp-infected SP-A null mice. BAL fluid from uninfected and Mp-infected mice was collected and the presence of HMGB-1 was examined by Western blot analysis. Viability of the cells recovered in BAL was assessed using Trypan blue and microscopy and no significant differences in cell death were noted between groups. As shown in Fig. 6A, levels of HMGB-1 in BAL of WT and SP-A null mice were similar in the uninfected mice. However, levels of HMGB-1, albeit increased in WT infected mice over uninfected controls, were dramatically increased in infected mice lacking SP-A (Fig. 6A).

Figure 6. Inhibition of HMGB-1 cytokine activity limits DC maturation to Mp.

Figure 6

Open in a new tab

A, Western analysis of cell free BAL harvested 3 days after Mp infection of WT and SP-A−/− mice for HMGB-1 expression and compared to HMGB-1 levels measured in saline treated mice. The same total volume was loaded for each sample. Control (C) is brain lysate. Representative of 3 separate experiments. B, WT and SP-A−/− mice were treated with Glycyrrhizin (10 mg/kg body weight) or vehicle (saline) prior to and during infection with Mp. Three days after infection, MLNs were harvested, cells collected after enzymatic digestion and density gradient centrifugation to enrich for DCs, and labeled with fluorescent antibodies for flow cytometry analysis. The number of MHCIIhi CD11c+ DCs from the MLNs that were CD86+ are shown. WT and SP-A−/− uninfected (U) mice were included as baseline controls. *p<.05, **p<.01, n=10–12/group, 3 combined experiments.

In order to determine if the heightened HMGB-1 secretion, and its potential cytokine activity, observed in the absence of SP-A could be influencing that maturation of dendritic cells in the Mp-infected mice, we used a direct inhibitor of HMGB-1 cytokine activity, glycyrrhizin (28). Cells from the MLNs were collected and analyzed from mice that were Mp-infected and treated with vehicle (saline) and compared to those that were Mp-infected and treated with glycyrrhizin. The total number of cells collected from the MLNs, which were enriched for DCs by gradient centrifugation, of uninfected WT and SP-A null mice were not significantly different. However, the total number of cells collected from the MLNs of both the WT and SP-A deficient Mp-infected mice were elevated over their uninfected controls (data not shown). When the DCs (MHC IIhi CD11c+) isolated from the MLNs were analyzed for the presence of the maturation marker CD86, again we detected significantly more cells expressing CD86 in the infected mice lacking SP-A that were treated with vehicle as compared to WT mice (Fig. 6B). In contrast, SP-A null mice that were treated with the HMGB-1 inhibitor, glycyrrhizin, had significantly fewer DCs that were expressing CD86 as compared to those receiving vehicle (Fig. 6B), indicating that SP-A can inhibit Mp-induced DC maturation by modulating HMGB-1 cytokine activities.

SP-A regulates HMGB-1 expression from Mp-stimulated human cells

In order to determine if exogenously added SP-A inhibits the release of HMGB-1 from Mp-activated cells into the culture supernatant, additional experiments were carried out in vitro using a human acute monocytic cell line (THP-1) as well as normal human bronchial epithelial cells (NHBE). The levels of HMGB-1 in the supernatant from THP-1 cells treated with either saline or SP-A alone were below the level of detection by Western analysis. However, when cells were infected with 10 Mp cfu per cell, secreted HMGB-1 was readily detected in the supernatants after 16 hours of stimulation (Fig. 7A). Pre-incubation of the cells prior to infection with exogenous human SP-A (50 μg/ml) inhibited secretion of HMGB-1 into the culture supernatant. Importantly, cellular viability (greater than 85%) was not significantly altered in stimulated conditions as determined by Trypan-blue exclusion and by lactate dehydrogenase assays (supplemental data figure 1).

Figure 7. SP-A inhibition of HMGB-1 secretion from human cells.

Figure 7

Open in a new tab

A, THP-1 cells or B, NHBE cells were incubated with 50 μg/ml purified human SP-A (+) or an equivalent volume of sterile Tris buffer (−) for 2 hours prior to stimulation with 10 Mp cfu/cell or with saline only (0). Cell free supernatants and lysates were harvested 16 hours post stimulation and HMGB-1 and GAPDH expression were assessed by Western analysis. Picture is representative of 2 experiments.

While monocytes are the most likely source of the secreted HMGB-1 present in the inflamed pulmonary environment, human epithelial cells could also potentially produce HMGB-1 upon stimulation. Therefore, we also examined the ability of SP-A to regulate HMGB-1 release from Mp-stimulated NHBE cells. Similar to observations with the THP-1 cells, levels of HMGB-1 were also undetectable in samples with the addition of either saline or SP-A alone. However, stimulation of cells with 10 Mp cfu per cell also induced HMGB-1 secretion from NHBE cells, while the addition of exogenous SP-A attenuated this response in Mp-stimulated samples (Fig. 7B).

SP-A binding to Mp limits HMGB-1 secretion from dendritic cells

While SP-A could be acting directly on DCs to inhibit Mp-induced maturation, it could also be functioning by binding to the Mp, which is known to occur through a binding protein (MPN372) and surface lipids (4, 5), and thereby protecting the cells from interactions with Mp. SP-A inhibited DC maturation when immature (Fig. 8A) or mature (Fig. 8B) DCs were pre-incubated with human SP-A. In order to determine which aspect of these DC/SP-A/Mp interactions was key to limiting DC maturation, we pre-incubated DCs with SP-A and washed away unbound SP-A prior to the addition of Mp. In parallel, we pre-coated Mp with SP-A, washed away unbound SP-A prior to addition to DCs. Interestingly, pre-incubation of DCs with SP-A prior to stimulation did not lead to reduced levels of HMGB-1 secretion, in fact HMGB-1 levels were still quite high (Fig. 8C). However, in the parallel experiment, Mp that had been pre-coated with SP-A was not able to elicit HMGB-1 secretion from DCs versus Mp that was not coated with SP-A (Fig. 8C).

Figure 8. SP-A interaction with Mp-stimulated DCs.

Figure 8

Open in a new tab

Bone marrow derived dendritic cells were cultured for 6 days in the presence of GM-CSF, purified and stimulated with Mp (10 cfu/cell) for 16 hours in the presence or absence of SP-A (50 μg/ml). Supernatants and lysates were harvested for analysis by SDS-PAGE and western immunoblotting for either HMGB-1, SP A or GAPDH. A, immature dendritic cells were used immediately after purification on day 6 or B, mature dendritic cells were stimulated for 24 hours in the presence of LPS prior to studies with Mp. C, dendritic cells were either treated for 16 hours with nothing (lane 1), pre-incubated with SP-A, washed, then stimulated with media (lane 2) or Mp (lane 3), or stimulated with SP-A coated Mp (lane 4) or non-coated Mp (lane 5). D, dendritic cells were pre-incubated with a TLR-2 neutralizing antibody or isotype control antibody prior to stimulation with Mp for 16 hours. Blots are representative from experiments conducted from 3 independent BMDC cultures.

Since Mp is known to work almost exclusively through TLR-2, additional experiments were conducted to determine if this receptor was a key receptor in Mp-induced HMGB-1 secretion from these cells. A TLR-2 neutralizing antibody or an isotype control was incubated with DCs prior to stimulation with Mp. Interestingly, inhibition of TLR-2 resulted in no detectable HMGB-1 secretion upon Mp stimulation (Fig. 8D).

DISCUSSION

Our findings describe a novel role for SP-A in limiting Mp-induced DC maturation via inhibition of HMGB-1 cytokine activity. The absence of SP-A in mice during Mp infection leads to increased numbers, as well as the activation state, of antigen presenting cells in the lung and draining lymph nodes during the acute phase of infection and consequently, increased numbers of activated T and B cells later during the course of infection. These findings are consistent with reports that describe SP-A as a vital component of the pulmonary innate immune system that limits inflammation and inhibits LPS-induced DC maturation in vitro. In contrast, reports have only recently begun to focus on the extracellular cytokine activity of HMGB-1 as an important mediator of inflammation that is actively secreted from stimulated myeloid cells and induces DC activation. While no association between HMGB-1 and SP-A has previously been described, several phenotypic parameters measured in Mp-infected SP-A null mice, suggested that SP-A may play role in regulating HMGB-1 extracellular cytokine activity. Using a specific inhibitor of HMGB-1 cytokine activity, glycyrrhizin, our research describes a novel role for SP-A in regulating HMGB-1 activity during Mp pulmonary infection.

Mp infection of mice lacking SP-A induced a dramatic and significant increase in the total number of CD3+ T cells present in both the lung and MLNs at both the early time point (3 days) and the later time point (9 days) as compared to WT infected mice. Mp is not likely acting as a mitogen in this response since the levels of Mp detected at 9 days of infection is significantly decreased in the lungs of both WT and SP-A null mice as compared to the burden at 3 days, while the numbers of T cells are continuing to increase (data not shown). Cell surface markers on T cells indicative of activation were also significantly increased in both the lung and MLNs of infected mice lacking SP-A. Additionally, compared to WT infected mice, SP-A null Mp infected mice had approximately four-fold more BAL IL-12, which is involved in T cell differentiation.

Although one of the many known roles of SP-A is to inhibit T cell proliferation and attenuate the initial Ca2+ spike (29), an additional likely explanation for the increase in activated T cells in infected SP-A null mice can be described by the role of SP-A in DC maturation. Previous work from our laboratory has determined that SP-A inhibits basal and LPS-induced DC maturation in vitro. Therefore, it was important to determine if this role would be carried out in vivo to a respiratory pathogen, such as Mp. Not only were more DCs found in the lung of Mp-infected mice that lacked SP-A, but more functionally mature DCs were also detected in the MLNs, where they would then act as potent T cell stimulators. Additionally, MCP and MIP-1, factors known to be chemotactic for immature DCs (21, 22), were present in BAL fluid of infected mice at much higher levels in mice lacking SP-A, suggesting that more DCs migrated into the lung from the bloodstream of Mp infected mice in the absence of SP-A due to greater chemokine production. The amount of GM-CSF, a factor vital to enhancing the differentiation of monocytes into immunostimulatory DCs in the lung vasculature, was significantly increased in BAL from both WT and SP-A null Mp infected mice. Collectively, these findings suggest that, in the absence of SP-A, more DCs infiltrate the airways due to increases in chemoattractant signals, however, since GM-CSF is increased comparably in both infected WT and SP-A null mice, it is not likely that the increase in GM-CSF accounts for the difference in DC maturation observed.

Since differential increases in GM-CSF in the SP-A null mice were not observed, we investigated the possibility that other mediators may be responsible for the SP-A mediated inhibition of DC maturation. Recent studies have suggested that endogenous stress factors, such as HMGB-1, released during infection can aid in maturing DCs that will then further contribute to the initiation and maintenance of an immune response against an invading pathogen (9). We thought that HMGB-1 was a good candidate for mediating the SP-A dependent modulation of DC and T cells because HMGB-1 can be actively secreted from activated monocytes and macrophages, is actively involved in regulating the maturation of DCs (30), and is also thought to be necessary for the proliferation and polarization of naïve CD4+ T cells (27). Indeed, we found that the level of HMGB-1 was dramatically increased in the BAL of Mp-infected mice when SP-A was absent. There was no indication of increased cell death in these mice, as determined by cell staining from the BAL and lung tissue, suggesting the increased HMGB-1 was released from activated, but not necrotic, airway monocytes or macrophages.

In order to determine if the increased HMGB-1 present in the lungs after Mp infection was responsible for the increased DC maturation observed in the absence of SP-A, we used an inhibitor of HMGB-1 cytokine activity, glycyrrhizin. Glycyrrhizin, a product produced by the licorice plant, has been shown to bind HMGB-1 directly and block its extracellular functions (28, 31). Taking advantage of the ability of glycyrrhizin to functionally inhibit HMGB-1 cytokine activity, we were able to examine the maturation state of those DCs that had migrated to the MLNs following Mp infection and determine if HMGB-1 was a key mediator in this process. In mice lacking SP-A, more mature DCs were again observed in the Mp infected vehicle treated mice as compared to WT infected vehicle treated mice. However, in mice lacking SP-A treated with glycyrrhizin, the level of DC maturation of migrated cells was significantly reduced to those levels measured in the WT infected mice. These novel findings suggest that SP-A inhibits DC maturation in vivo to Mp infection at least in part by limiting HMGB-1 extracellular cytokine activity, which can directly influence initiation of an adaptive immune response.

While we used cell surface markers most commonly used to distinguish macrophages from DCs, we acknowledge that phenotypic distinction of these two populations of APCs is increasingly nebulous. Although, the total numbers of phenotypically activated DCs were significantly increased in Mp-infected SP-A null mice, the percent of activated T lymphocytes was similar. This profile suggests that the increase in APCs maybe due to increases in the pool of activated macrophages rather than differentiated DCs, the latter of which should induce activation of T-cells. Therefore, more detailed studies were conducted using a purified population of BMDCs in which we found that exogenous SP-A added at physiologic levels could inhibit DC maturation induced by Mp stimulation. However, functional assays examining antigen specific lymphocyte proliferation should be conducted in the future to strengthen our understanding of the inhibitory role of SP-A in DC differentiation independent of any macrophage participation.

Additional experiments were also carried out in vitro using a human monocyte cell line (THP-1) and human bronchial epithelial cells to determine if exogenously added SP-A inhibits the release of HMGB-1 from Mp-activated cells into the culture supernatant. The levels of HMBG-1 in the supernatant from THP-1 cells treated with either saline or SP-A alone were below the level of detection by Western analysis. However, when cells were infected with 10 Mp cfu per cell, secreted HMGB-1 was detected in the supernatants as has been previously described for human monocytes (32). Pre-incubation of the cells prior to infection with exogenous human SP-A indeed inhibited secretion of HMGB-1 into the supernatant in both the human monoyte cells and bronchial epithelial cells. Importantly, cellular viability and membrane permeability were not significantly altered in stimulated conditions, indicating the increased HMGB-1 was not from increased cell death or membrane leakage within those samples.

The amount of HMGB-1 detected in BAL from infected SP-A null mice is increased compared to infected WT mice, which suggests that SP-A may regulate the amount of HMGB-1 secreted from the activated cells. While it is possible that SP-A directly binds extracellular HMGB-1, co-immunoprecipitation experiments with BALF from infected WT mice showed no detectable binding, while both HMGB-1 and SP-A were readily observed. Further studies determined that the interaction and binding of SP-A to Mp was a vital component in limiting Mp-induced HMGB-1 secretion from cultured DCs. Additionally, when a neutralizing antibody was used to inhibit TLR-2, Mp was unable to elicit HMGB-1 secretion from these cells. Taken together, these findings suggest that the binding of SP-A to Mp is critical in curtailing HMGB-1 secretion from DCs by restricting the interaction of Mp to its primary receptor, TLR-2.

Mp is known to colonize the respiratory tract where it initiates a cascade of immune response amplification, including proliferation of lymphocytes, proinflammatory cytokine release and production of immunoglobulins. Several studies have reported increased IgG serum levels in Mp-infected individuals (33, 34). Our findings show that in the absence of SP-A, more Mp-specific IgG is present in the serum of infected mice as compared to WT infected mice. The production of IgG antibodies is predominantly associated with the secondary immune response. This finding further supports an indirect role for SP-A in limiting the advancement of an adaptive immune response by regulating the initiation of the antibody response to Mp that could be a direct result of increased DC maturation and migration in the innate phase of the response.

In summary, our findings support an inhibitory role for SP-A in Mp-induced DC maturation, which is a key step in the initiation of an immune response. Additionally, our studies show that one mechanism by which SP-A inhibits Mp-induced DC maturation is by regulating HMGB-1 secretion both in vivo and in vitro in human cells. Previous studies have shown that the receptor for advanced glycation end products (RAGE) and HMGB-1 are required for maturation of human DCs (30). Since SP-A inhibits DC maturation in the presence and absence of stimulation, further experiments testing whether SP-A interferes with HMGB-1 binding via RAGE and limiting DC maturation through this interaction may be of interest. These findings could potentially be of value in designing therapies, since HMGB1, as well as other proinflammatory ligands, and the RAGE are present in a myriad of acute and chronic inflammatory diseases such as sepsis, diabetes, atherosclerosis and renal failure (35). Additionally, chronic inflammation and inflammatory diseases in the lung often result in epithelial damage and airway remodeling, therefore blocking HMGB-1 by pharmacological interventions or potentially with surfactant treatment, may alleviate lung damage.

Supplementary Material

text for SF1
SF1

ACKNOWLEDGEMENTS

We would like to thank Charles Giamberardino for technical assistance, Pamela Hesker for HMGB-1 antibody advice, and Sambuddho Mukherjee and Amy Pastva for helpful discussions.

Research Support: F32HL091642, HL084917 and AI81672.

Abbreviations

BAL

bronchoalveolar lavage

FSC

forward scatter

HMGB-1

high-mobility group box-1

MIP-1α

macrophage inflammatory protein-1α

MLN

mediastinal lymph node

MCP-1

monocytes chemotactic protein-1

MIG

monokine induced by gamma interferon

Mp

Mycoplasma pneumoniae

NHBE

normal human bronchial epithelial cell

SSC

side scatter

SP-A

surfactant protein-A

Footnotes

The authors have no conflicting financial interests.

REFERENCES

  • 1.Waites KB, Talkington DF. Mycoplasma pneumoniae and its role as a human pathogen. Clin Microbiol Rev. 2004;17:697–728. doi: 10.1128/CMR.17.4.697-728.2004. table of contents. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Martin RJ, Kraft M, Chu HW, Berns EA, Cassell GH. A link between chronic asthma and chronic infection. J Allergy Clin Immunol. 2001;107:595–601. doi: 10.1067/mai.2001.113563. [DOI] [PubMed] [Google Scholar]
  • 3.Kraft M, Cassell GH, Henson JE, Watson H, Williamson J, Marmion BP, Gaydos CA, Martin RJ. Detection of Mycoplasma pneumoniae in the airways of adults with chronic asthma. Am J Respir Crit Care Med. 1998;158:998–1001. doi: 10.1164/ajrccm.158.3.9711092. [DOI] [PubMed] [Google Scholar]
  • 4.Kannan TR, Provenzano D, Wright JR, Baseman JB. Identification and characterization of human surfactant protein A binding protein of Mycoplasma pneumoniae. Infect Immun. 2005;73:2828–2834. doi: 10.1128/IAI.73.5.2828-2834.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Piboonpocanun S, Chiba H, Mitsuzawa H, Martin W, Murphy RC, Harbeck RJ, Voelker DR. Surfactant protein A binds Mycoplasma pneumoniae with high affinity and attenuates its growth by recognition of disaturated phosphatidylglycerols. J Biol Chem. 2005;280:9–17. doi: 10.1074/jbc.M411570200. [DOI] [PubMed] [Google Scholar]
  • 6.Ledford JG, Goto H, Potts EN, Degan S, Chu HW, Voelker DR, Sunday ME, Cianciolo GJ, Foster WM, Kraft M, Wright JR. SP-A preserves airway homeostasis during Mycoplasma pneumoniae infection in mice. J Immunol. 2009;182:7818–7827. doi: 10.4049/jimmunol.0900452. [DOI] [PMC free article] [PubMed] [Google Scholar] [Research Misconduct Found]
  • 7.Wright JR. Immunoregulatory functions of surfactant proteins. Nat Rev Immunol. 2005;5:58–68. doi: 10.1038/nri1528. [DOI] [PubMed] [Google Scholar]
  • 8.Brinker KG, Garner H, Wright JR. Surfactant protein A modulates the differentiation of murine bone marrow-derived dendritic cells. Am J Physiol Lung Cell Mol Physiol. 2003;284:L232–241. doi: 10.1152/ajplung.00187.2002. [DOI] [PubMed] [Google Scholar]
  • 9.Telusma G, Datta S, Mihajlov I, Ma W, Li J, Yang H, Newman W, Messmer BT, Minev B, Schmidt-Wolf IG, Tracey KJ, Chiorazzi N, Messmer D. Dendritic cell activating peptides induce distinct cytokine profiles. Int Immunol. 2006;18:1563–1573. doi: 10.1093/intimm/dxl089. [DOI] [PubMed] [Google Scholar]
  • 10.Korfhagen TR, Bruno MD, Ross GF, Huelsman KM, Ikegami M, Jobe AH, Wert SE, Stripp BR, Morris RE, Glasser SW, Bachurski CJ, Iwamoto HS, Whitsett JA. Altered surfactant function and structure in SP-A gene targeted mice. Proc Natl Acad Sci U S A. 1996;93:9594–9599. doi: 10.1073/pnas.93.18.9594. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Sitia G, Iannacone M, Muller S, Bianchi ME, Guidotti LG. Treatment with HMGB1 inhibitors diminishes CTL-induced liver disease in HBV transgenic mice. J Leukoc Biol. 2007;81:100–107. doi: 10.1189/jlb.0306173. [DOI] [PubMed] [Google Scholar]
  • 12.Pecora ND, Fulton SA, Reba SM, Drage MG, Simmons DP, Urankar-Nagy NJ, Boom WH, Harding CV. Mycobacterium bovis BCG decreases MHC-II expression in vivo on murine lung macrophages and dendritic cells during aerosol infection. Cell Immunol. 2008 doi: 10.1016/j.cellimm.2008.07.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Lin KL, Suzuki Y, Nakano H, Ramsburg E, Gunn MD. CCR2+ monocyte-derived dendritic cells and exudate macrophages produce influenza-induced pulmonary immune pathology and mortality. J Immunol. 2008;180:2562–2572. doi: 10.4049/jimmunol.180.4.2562. [DOI] [PubMed] [Google Scholar]
  • 14.Inaba K, Inaba M, Romani N, Aya H, Deguchi M, Ikehara S, Muramatsu S, Steinman RM. Generation of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colony-stimulating factor. J Exp Med. 1992;176:1693–1702. doi: 10.1084/jem.176.6.1693. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Brinker KG, Martin E, Borron P, Mostaghel E, Doyle C, Harding CV, Wright JR. Surfactant protein D enhances bacterial antigen presentation by bone marrow-derived dendritic cells. Am J Physiol Lung Cell Mol Physiol. 2001;281:L1453–1463. doi: 10.1152/ajplung.2001.281.6.L1453. [DOI] [PubMed] [Google Scholar]
  • 16.McIntosh JC, Mervin-Blake S, Conner E, Wright JR. Surfactant protein A protects growing cells and reduces TNF-alpha activity from LPS-stimulated macrophages. Am J Physiol. 1996;271:L310–319. doi: 10.1152/ajplung.1996.271.2.L310. [DOI] [PubMed] [Google Scholar]
  • 17.Cartner SC, Simecka JW, Lindsey JR, Cassell GH, Davis JK. Chronic respiratory mycoplasmosis in C3H/HeN and C57BL/6N mice: lesion severity and antibody response. Infect Immun. 1995;63:4138–4142. doi: 10.1128/iai.63.10.4138-4142.1995. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Cook DN, Bottomly K. Innate immune control of pulmonary dendritic cell trafficking. Proc Am Thorac Soc. 2007;4:234–239. doi: 10.1513/pats.200701-026AW. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Kirby AC, Raynes JG, Kaye PM. CD11b regulates recruitment of alveolar macrophages but not pulmonary dendritic cells after pneumococcal challenge. J Infect Dis. 2006;193:205–213. doi: 10.1086/498874. [DOI] [PubMed] [Google Scholar]
  • 20.Lambrecht BN, Prins JB, Hoogsteden HC. Lung dendritic cells and host immunity to infection. Eur Respir J. 2001;18:692–704. [PubMed] [Google Scholar]
  • 21.McWilliam AS, Napoli S, Marsh AM, Pemper FL, Nelson DJ, Pimm CL, Stumbles PA, Wells TN, Holt PG. Dendritic cells are recruited into the airway epithelium during the inflammatory response to a broad spectrum of stimuli. J Exp Med. 1996;184:2429–2432. doi: 10.1084/jem.184.6.2429. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Sozzani S, Sallusto F, Luini W, Zhou D, Piemonti L, Allavena P, Van Damme J, Valitutti S, Lanzavecchia A, Mantovani A. Migration of dendritic cells in response to formyl peptides, C5a, and a distinct set of chemokines. J Immunol. 1995;155:3292–3295. [PubMed] [Google Scholar]
  • 23.Vermaelen KY, Carro-Muino I, Lambrecht BN, Pauwels RA. Specific migratory dendritic cells rapidly transport antigen from the airways to the thoracic lymph nodes. J Exp Med. 2001;193:51–60. doi: 10.1084/jem.193.1.51. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Wilson NS, El-Sukkari D, Belz GT, Smith CM, Steptoe RJ, Heath WR, Shortman K, Villadangos JA. Most lymphoid organ dendritic cell types are phenotypically and functionally immature. Blood. 2003;102:2187–2194. doi: 10.1182/blood-2003-02-0513. [DOI] [PubMed] [Google Scholar]
  • 25.Andersson U, Wang H, Palmblad K, Aveberger AC, Bloom O, Erlandsson-Harris H, Janson A, Kokkola R, Zhang M, Yang H, Tracey KJ. High mobility group 1 protein (HMG-1) stimulates proinflammatory cytokine synthesis in human monocytes. J Exp Med. 2000;192:565–570. doi: 10.1084/jem.192.4.565. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Erlandsson Harris H, Andersson U. Mini-review: The nuclear protein HMGB1 as a proinflammatory mediator. Eur J Immunol. 2004;34:1503–1512. doi: 10.1002/eji.200424916. [DOI] [PubMed] [Google Scholar]
  • 27.Dumitriu IE, Baruah P, Valentinis B, Voll RE, Herrmann M, Nawroth PP, Arnold B, Bianchi ME, Manfredi AA, Rovere-Querini P. Release of high mobility group box 1 by dendritic cells controls T cell activation via the receptor for advanced glycation end products. J Immunol. 2005;174:7506–7515. doi: 10.4049/jimmunol.174.12.7506. [DOI] [PubMed] [Google Scholar]
  • 28.Girard JP. A direct inhibitor of HMGB1 cytokine. Chem Biol. 2007;14:345–347. doi: 10.1016/j.chembiol.2007.04.001. [DOI] [PubMed] [Google Scholar]
  • 29.Borron P, McCormack FX, Elhalwagi BM, Chroneos ZC, Lewis JF, Zhu S, Wright JR, Shepherd VL, Possmayer F, Inchley K, Fraher LJ. Surfactant protein A inhibits T cell proliferation via its collagen-like tail and a 210-kDa receptor. Am J Physiol. 1998;275:L679–686. doi: 10.1152/ajplung.1998.275.4.L679. [DOI] [PubMed] [Google Scholar]
  • 30.Dumitriu IE, Baruah P, Bianchi ME, Manfredi AA, Rovere-Querini P. Requirement of HMGB1 and RAGE for the maturation of human plasmacytoid dendritic cells. Eur J Immunol. 2005;35:2184–2190. doi: 10.1002/eji.200526066. [DOI] [PubMed] [Google Scholar]
  • 31.Mollica L, De Marchis F, Spitaleri A, Dallacosta C, Pennacchini D, Zamai M, Agresti A, Trisciuoglio L, Musco G, Bianchi ME. Glycyrrhizin binds to high-mobility group box 1 protein and inhibits its cytokine activities. Chem Biol. 2007;14:431–441. doi: 10.1016/j.chembiol.2007.03.007. [DOI] [PubMed] [Google Scholar]
  • 32.Bonaldi T, Talamo F, Scaffidi P, Ferrera D, Porto A, Bachi A, Rubartelli A, Agresti A, Bianchi ME. Monocytic cells hyperacetylate chromatin protein HMGB1 to redirect it towards secretion. EMBO J. 2003;22:5551–5560. doi: 10.1093/emboj/cdg516. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Shimizu T, Mochizuki H, Kato M, Shigeta M, Morikawa A, Hori T. Immunoglobulin levels, number of eosinophils in the peripheral blood and bronchial hypersensitivity in children with Mycoplasma pneumoniae pneumonia. Arerugi. 1991;40:21–27. [PubMed] [Google Scholar]
  • 34.Tang LF, Shi YC, Xu YC, Wang CF, Yu ZS, Chen ZM. The change of asthma-associated immunological parameters in children with Mycoplasma pneumoniae infection. J Asthma. 2009;46:265–269. doi: 10.1080/02770900802647557. [DOI] [PubMed] [Google Scholar]
  • 35.Maillard-Lefebvre H, Boulanger E, Daroux M, Gaxatte C, Hudson BI, Lambert M. Soluble receptor for advanced glycation end products: a new biomarker in diagnosis and prognosis of chronic inflammatory diseases. Rheumatology (Oxford) 2009;48:1190–1196. doi: 10.1093/rheumatology/kep199. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

text for SF1
NIHMS455123-supplement-text_for_SF1.docx (43.4KB, docx)
SF1
NIHMS455123-supplement-Supplemental_Figure_1.jpg (23.9KB, jpg)

RESOURCES