The pulmonary innate immune system responds to various airborne microbes. Although its specificity is broad and based on the recognition of pathogen-associated molecular patterns, it is uniquely regulated to limit inflammation and thereby prevent damage to the gas-exchanging alveoli. Macrophages, critical cell determinants of this system, recognize microbes through pattern recognition receptors such as TLRs, which typically mediate proinflammatory responses. The lung collectin, surfactant protein A (SP-A), has emerged as an important innate immune determinant that regulates microbe-macrophage interactions in this environment. In this study, we report the basal and SP-A-induced transcriptional and posttranslational regulation of TLR2 and TLR4 expression during the differentiation of primary human monocytes into macrophages. Despite SP-A’s ability to up-regulate TLR2 expression on human macrophages, it dampens TLR2 and TLR4 signaling in these cells. SP-A decreases the phosphorylation of IκBα, a key regulator of NF-κB activity, and nuclear translocation of p65 which result in diminished TNF-α secretion in response to TLR ligands. SP-A also reduces the phosphorylation of TLR signaling proteins upstream of NF-κB, including members of the MAPK family. Finally, we report for the first time that SP-A decreases the phosphorylation of Akt, a major cell regulator of NF-κB and potentially MAPKs. These data identify a critical role for SP-A in modulating the lung inflammatory response by regulating macrophage TLR activity.

Although the majority of inhaled microbes and particulates are trapped and cleared by the upper airways of the lung, some organisms are able to travel further to the terminal bronchioles and alveoli. The degree of inflammation initiated in this site by a stimulus is tightly regulated by several elements of the innate immune system, as even moderate inflammation could be harmful to the gas-exchanging alveolar structures (1). Two key components involved in this regulation are phagocytes, mainly alveolar macrophages (AM),3 and pulmonary surfactant. AM, which are bathed in surfactant, are the first professional phagocytes to encounter, internalize, and degrade invading microbes (2).

AM are considered alternatively activated, based upon their unique biological attributes. These include a greater phagocytic potential compared with other macrophages (2, 3) due to significant expression and activity of PRRs, such as the mannose receptor (MR) and scavenger receptor A (SR-A) (4, 5); and immunoregulation through production of proinflammatory cytokines (e.g., such as TNF-α and/or anti-inflammatory cytokines (e.g., TGF-β) (2, 3). They also produce less IL-1β (TNF-α), have a reduced oxidative response to pathogens compared with blood monocytes (6, 7, 8), and serve as poor APCs (9), all of which are mechanisms that serve to control alveolar inflammation.

TLRs are important cell-associated and intracellular PRRs. There are currently 12 identified mammalian TLRs, of which TLR2 and TLR4 are among the most widely studied and are considered the major transmembrane TLRs (10). TLR2 and TLR4 play roles in initiating immune responses against pathogens. TLR2 forms a heterodimer with TLR6 or TLR1 to recognize diacyl and triacyl lipopeptides, respectively. TLR2 binds to zymosan, a particle composed of yeast cell wall components; peptidoglycan, a component of the Gram-positive bacteria cell wall (11, 12), and Pam3Cys-Ser-(Lys)4 hydrochloride (Pam3Cys), a lipohexapeptide analog of the immunologically active N-terminal portion of bacterial lipoprotein (13, 14). TLR4 associates with MD2 and CD14, enabling recognition of LPS, a major component of the Gram-negative cell wall, and heat shock proteins 60 and 70, which are ubiquitously expressed (15, 16, 17, 18).

Because an unrestricted inflammatory response induced by TLR signaling can be potentially harmful to the host, this signaling is tightly regulated, detecting the presence of certain microbial determinants and responding differently depending on the stimulus. The signaling cascade typically produces proinflammatory cytokines; however, activation through TLRs can also generate negative regulators that inhibit a proinflammatory response (19).

Surfactant lipids and the surfactant-associated proteins A, B, C, and D are major components of pulmonary surfactant, which lines the lung alveolus and serves to decrease surface tension at the air-liquid interface of the lung (20, 21, 22). Surfactant protein A (SP-A) is a large, multimeric protein with trimers that are assembled into an 18-mer protein through interactions between the collagen-like domains (23, 24, 25). It is classified as a collectin because each monomer contains a linear collagen-like sequence, a short linking domain, and a globular Ca2+-dependent carbohydrate recognition domain (CRD) (22).

SP-A has been implicated as a key component of the lung innate immune response because it mediates host interactions with a variety of microbial pathogens (26) and regulates the nature of the inflammatory response. In this regard, SP-A can serve both as a microbial opsonin and as a direct activator of macrophage function. SP-A has been shown to up-regulate certain PRRs, such as SR-A and the MR, on macrophages (4, 5, 27). SP-A also regulates TNF-α production, either up or down, depending on which receptor it binds on the cell surface and the activation state of the cell (28). In human macrophages, SP-A down-regulates oxidant production to stimuli by decreasing NADPH oxidase activity (29).

Despite a large body of literature on the importance of TLR2 and TLR4 in pathogen recognition and host response, there is limited information about TLR2 and TLR4 expression on primary human mononuclear phagocytes and the response of these cells to TLR agonists, particularly during monocyte differentiation into macrophages. SP-A has emerged as a key regulator of the phagocyte response in the alveolar compartment and contributes to the characteristic alternative activation state of the macrophages in this environment. In this study, we report the discovery that SP-A differentially regulates the expression of TLR2 and TLR4 during primary human monocyte differentiation into macrophages. Despite up-regulation of TLR2 expression, SP-A markedly diminishes the macrophage proinflammatory response generated by both TLR2 and TLR4 agonists. The underlying mechanism is related to altered phosphorylation of a central regulator of cellular function, Akt, as well as downstream intermediates in the MAPK pathway and the activation of NFκB. These studies underscore the important role of SP-A in shaping the biology of macrophages in the lung alveoli.

Dulbecco’s PBS with and without Ca2+ and Mg2+ ions and RPMI 1640 medium with l-glutamine (RPMI) were purchased from Invitrogen. PBF buffer (PBS without Ca2+ and Mg2+ (Invitrogen), 5 mg/ml BSA (Sigma-Aldrich), and 10% heat-inactivated FBS (HyClone)) was used as a blocking agent for confocal microscopy experiments. RHH medium (RPMI 1640 plus 10 mM HEPES plus 0.4% human serum albumin) was used for cell culture experiments. Pam3Cys (Calbiochem) and Escherichia coli LPS (Sigma-Aldrich) were used for functional assays involving Western blotting and ELISAs.

Allophycocyanin-labeled anti-human TLR2 (clone T2.1), PE-labeled anti-human TLR4 (clone HTA125), and mouse IgG2a were purchased from eBioscience for flow cytometry experiments. Unconjugated mouse anti-human TLR2 (clone TL2.1; Novus Biologicals), mouse anti-human TLR4 (clone HTA125; Gene Tex), mouse IgG2a (clone 20102; R&D Systems), mouse anti-human MR (clone 19.2; BD Biosciences), and mouse IgG1 (R&D Systems) were used for confocal microscopy experiments in which Alexa Fluor 488-conjugated goat anti-mouse IgG (Molecular Probes and Invitrogen Detection Technologies) was used as a secondary Ab. Phosphorylated Akt (phospho-Akt Ser473), phosphorylated p38 (phospho-p38 MAPK Thr180/Tyr182), phosphorylated ERK (phospho-p44/42 MAPK Thr202/Tyr204), phosphorylated JNK (phospho-SAPK/JNK T183/Y185), phosphorylated IκBα (Ser32), and IκBα Abs were purchased from Cell Signaling Technology, while actin, total Akt, goat anti-rabbit IgG-HRP (secondary Abs for primary Abs to phosphorylated proteins), and donkey anti-goat IgG-HRP (actin and total Akt secondary Ab) Abs were purchased from Santa Cruz Biotechnology and used for Western blotting experiments. TNF-α ELISA kits were purchased from R&D Systems.

The SP-A proteins used in this study were purified as previously described (4). In brief, bronchoalveolar lavage (BAL) from alveolar proteinosis patients (APP) was used to obtain APP-SP-A (4). Purity of the SP-A preparation was assessed by SDS-PAGE. Bacterial endotoxin levels were determined using the Limulus amebocyte lysate kit (BioWhittaker). Endotoxin levels in SP-A preparations ranged from undetectable to 0.2 pg/μg protein. Two SP-A functional assays, oxidative burst and liposome aggregation (29, 30), were performed as quality control experiments.

Blood was obtained from healthy adult volunteers using an approved protocol by the Ohio State University Institutional Review Board. PBMC from single donors were isolated from heparinized blood on Ficoll-Paque (Amersham Biosciences) and cultured in Teflon wells (Savillex) for 1 (monocytes) through 5 (monocyte-derived macrophages (MDMs)) days in the presence of RPMI 1640 medium containing 20% autologous serum (2.0 × 106 PBMC/ml) at 37°C (4). On the day of each experiment, PBMC were removed from Teflon wells and washed extensively. Monocytes and MDMs were further purified by adherence in tissue culture plates in some assays. Human AM were isolated from BALs of healthy human donors as previously described (31) using an approved protocol by the Ohio State University Institutional Review Board. Briefly, the BAL was centrifuged (200 × g, 4°C, 10 min), supernatant removed, and the pellet resuspended in RPMI and washed two more times with RPMI.

PBMC were incubated with APP-SP-A (10 μg/ml) or human serum albumin (control) in Teflon wells for specific time periods. PBMC were harvested from Teflon wells, centrifuged (200 × g, 4°C, 10 min), and resuspended in RPMI. After the cells were counted, they were centrifuged, resuspended in FACS buffer (2% BSA), and incubated with allophycocyanin- or PE-conjugated Abs to TLR2 or TLR4 (20 μl/million cells), respectively, for 20 min. PBMC incubated with the appropriate allophycocyanin- or PE-conjugated subtypic control mAb served as negative controls.

For AM experiments, cells were counted, centrifuged, and resuspended in FACS buffer. Two × 105 AM were incubated with TLR2, TLR4, or subtype control Abs as described above for PBMC.

Ab-stained cells were fixed in 2% paraformaldehyde and 1 × 104 MDM or AM were analyzed for mean fluorescence intensity (MFI) and percentage of positive cells (95/5% cutoff) using the BD FACSCalibur System (BD Biosciences). Macrophages were distinguished by the side scatter vs forward scatter (4), and the MFI due to nonspecific binding (subtypic control Ab) was subtracted from each sample to obtain a specific MFI. The percent change in specific MFI in experiments with no treatment vs SP-A treatment were calculated as follows: (treated MFI − untreated MFI)/untreated MFI) × 100. Triplicate samples in each experiment were analyzed.

One- to 5-day-old PBMC in Teflon wells were harvested and mononuclear phagocytes adhered to a 12-well tissue culture plate with 10% autologous serum (3 × 106 PBMC/ml). After washing away lymphocytes, the monocytes or MDMs (3 × 105 cells) were treated with SP-A (10 μg/ml) or medium (untreated) over a time course. Finally, cells were lysed in TRIzol (Invitrogen) and total RNA was isolated by using the Qiagen RNeasy column method. The Experion (Bio-Rad) was used to determine the RNA quality and quantity of each sample.

RNA (550 ng) was converted to cDNA by reverse transcriptase enzyme and real-time PCR was performed with a human TLR2, TLR4, or MR TaqMan gene expression kit (Applied Biosystems). TLR2, TLR4, and MR amplification were normalized to the β-actin housekeeping gene (ΔCt). Relative copy number (RCN) and fold change were determined. RCN was calculated as follows: RCN = EΔCt × 100. E is the efficiency (2 = 100% efficiency) and ΔCt = Ct(target) − Ct(reference) (32). Fold change was calculated as 2−ΔΔCt. ΔΔCt = ΔCt (experimental cell group) − ΔCt (unstimulated cells). Duplicate samples were analyzed in each experiment.

Two × 105 MDMs were adhered to a glass coverslip in each well of a 24-well tissue culture plate for 2 h at 37°C, washed to remove lymphocytes, and incubated with APP-SP-A (10 μg/ml) or RHH for 2 h at 37°C. The cells were washed, fixed with 2% paraformaldehyde (10 min at room temperature), and then incubated overnight in blocking buffer (PBF). The cells were then stained with TLR2 (8 μg/ml), TLR4 (8 μg/ml), MR (1 μg/ml), or the appropriate subtype (8 μg/ml or 1 μg/ml) control Ab for 1 h at room temperature, washed with PBF buffer, and counterstained with an Alexa Fluor 488-conjugated secondary Ab (1/500) for 1 h at room temperature. The cells were next washed with PBF buffer and coverslips were mounted on glass slides. Slides were viewed using a confocal, Zeiss scanning laser microscope and fluorescence intensity of each cell slice was quantified using a pixel intensity measurement (NIH Image J program). An analytic box was placed on the cell membrane of each cell at four points (as represented on a clock at 12 a.m., 3 a.m., 6 a.m., 9 a.m.) and the MFI was calculated as the mean of the MFI of all four points. The MFI was determined for 20 cells per coverslip, with duplicate slides per experiment, and a minimum of two donors used for each treatment condition.

Three × 105 day 5 MDMs were adhered to wells of a 12-well plate for 2 h at 37°C, washed to remove lymphocytes, and repleted with RPMI containing 10% autologous serum overnight at 37°C. APP-SP-A in RHH medium was added to appropriate wells for 10 min and then cells were washed with RHH and incubated for another 10 min at 37°C to allow for internalization of all bound SP-A, as previously published (33). Dose (5–20 μg/ml) and time (5 min to 2 h) course experiments were conducted initially to determine the optimal concentration and time for SP-A incubation. Pam3Cys (100 or 500 ng/ml) or LPS (100 or 500 ng/ml) was added to appropriate wells for 5, 10, 15, or 30 min at 37°C, followed by the removal of medium in preliminary experiments to determine the optimal time point for analysis. Lysis buffer (TN1 buffer: 50 mM Tris (pH 8.0), 10 mM EDTA, 10 mM Na4PO7, 10 mM NaF, 1% Triton X-100, and 125 mM NaCl, 10 mM Na3VO4, 10 μg/ml aprotinin, and 10 μg/ml leupeptin (34)) was added to each well and the resultant lysates were added to Eppendorf tubes and incubated on ice, and then centrifuged at 13,000 × g for 10 min to remove cell debris. The cleared lysates containing soluble proteins were collected and protein content was calculated using a BCA kit (Pierce). Proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes. Finally, the blots were blocked with 5% nonfat dry milk and probed with the appropriate primary (Abs to phosphorylated proteins 1/600; actin or total Akt Ab 1/1000) and secondary HRP-conjugated secondary Abs (1/1000). The blots were developed using the ECL kit (Amersham Biosciences). The Image J program was used to quantify band intensity. Background intensity was subtracted from each sample and then fold change was determined as follows: (treated sample band intensity/untreated sample band intensity).

One × 105 day 5 MDMs were adhered to each well of a 96-well plate for 2 h at 37°C, washed to remove lymphocytes, and then incubated with RHH medium with APP-SP-A (10 μg/ml) or RHH for 2 h at 37°C. Cells were washed with and resuspended in RHH and incubated for an additional 10 min at 37°C to allow for internalization of bound SP-A. Cells were then incubated with RHH alone, LPS (1 ng/ml), or Pam3Cys (10 ng/ml) for 1 h. Cell supernatants were collected and centrifuged at 200 × g for 5 min to remove dead cells, and TNF-α was measured by ELISA. Quantitative ELISAs were performed using the Quantikine ELISA kits from R&D Systems according to the manufacturer’s instructions.

Four × 105 day 5 MDMs were adhered to each well of a 12-well tissue culture plate for 2 h at 37°C, washed to remove lymphocytes, and then incubated with APP-SP-A (10 μg/ml) or RHH medium for 10 min. The cells were then washed with RHH and incubated for another 10 min at 37°C to allow for internalization of all bound SP-A. Pam3Cys (500 ng/ml) or LPS (500 ng/ml) was added to appropriate wells for 1 h. Medium was removed and nuclear extracts from MDMs (two wells combined to make 8 × 105 MDMs/sample) were prepared using the Transfactor extraction kit (BD Clontech). The nuclear lysates were assayed for the presence of p65 in wells precoated with the DNA-binding consensus sequence by using the colorimetric TransFactor kit (BD Clontech) and samples were read at 655 nm. Untreated, SP-A plus LPS or Pam3Cys, and SP-A groups were compared with LPS or Pam3Cys. LPS or Pam3Cys were set to 100% p65. Percent inhibition was determined as follows: ((experimental cell group − LPS or Pam3Cys cell group)/(LPS or Pam3Cys cell group) × 100); 100 minus percent inhibition = percent p65 compared with LPS or Pam3Cys.

An unpaired one-tailed Student’s t test was used to analyze differences between two groups (e.g., with or without SP-A) and a one-way ANOVA with the post-Tukey test was used to analyze differences among multiple test groups.

Studies of TLR2 and TLR4 expression and function have been conducted largely in transfected murine cell lines or murine macrophages, with less data available in human monocytes and very limited data in human macrophages. There are significant differences in the structures of murine and human TLRs as well as in their responsiveness to various stimuli, such as LPS (35, 36, 37, 38, 39, 40, 41). Therefore, we began our studies by determining the basal surface protein expression of TLR2 and TLR4 by flow cytometry using an established model of human monocytes and MDMs (4). Such a model allows us to assess changes in TLR expression during differentiation of cells from monocytes to macrophages in individual donors. The results of a typical experiment are displayed as histograms (Fig. 1, A, B, C, and E–G) and cumulative data are presented as bar graphs (Fig. 1, D and H). The data show that primary human monocytes express the highest level of TLR2 and this expression decreases when monocytes differentiate into macrophages. Monocytes and macrophages express equivalent amounts of TLR4, indicating that TLR4 expression remains unchanged during differentiation. By flow cytometry, TLR2 and TLR4 were both found to be expressed on the surface of human alveolar macrophages (data not shown).

FIGURE 1.

Human monocytes and macrophages express TLR2 and TLR4. PBMC were isolated from human blood and incubated in Teflon wells with autologous serum. The monocytes differentiate into macrophages (MDM) by day 5. One (A and E)-, 3 (B and F)-, or 5 (C and G)-day-old PBMCs were harvested and incubated with either an allophycocyanin-conjugated human TLR2 mAb (or an allophycocyanin-conjugated subtype control mAb) or a PE-conjugated human TLR4 mAb (or a PE-conjugated subtype control mAb). The stained cells were analyzed using flow cytometry by gating on the monocytes or macrophages (4 ), and a representative experiment is shown. The number in the top right corner represents the specific MFI (TLR2 or TLR4 MFI − subtype control MFI). D and H, Bar graph with cumulative data (triplicate samples in each experiment; n = 5 for TLR2; n = 4 for TLR4). One-way ANOVA with post-Tukey test, ∗∗∗, p < 0.005 compared with day 1 cells (±SEM).

FIGURE 1.

Human monocytes and macrophages express TLR2 and TLR4. PBMC were isolated from human blood and incubated in Teflon wells with autologous serum. The monocytes differentiate into macrophages (MDM) by day 5. One (A and E)-, 3 (B and F)-, or 5 (C and G)-day-old PBMCs were harvested and incubated with either an allophycocyanin-conjugated human TLR2 mAb (or an allophycocyanin-conjugated subtype control mAb) or a PE-conjugated human TLR4 mAb (or a PE-conjugated subtype control mAb). The stained cells were analyzed using flow cytometry by gating on the monocytes or macrophages (4 ), and a representative experiment is shown. The number in the top right corner represents the specific MFI (TLR2 or TLR4 MFI − subtype control MFI). D and H, Bar graph with cumulative data (triplicate samples in each experiment; n = 5 for TLR2; n = 4 for TLR4). One-way ANOVA with post-Tukey test, ∗∗∗, p < 0.005 compared with day 1 cells (±SEM).

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Previous data on TLR2 and TLR4 mRNA expression in monocytes/macrophages are inconsistent, which likely reflects differences in cell type and activation state of the cells (38, 41, 42, 43, 44). We used real-time PCR to determine steady-state mRNA levels of TLR2 and TLR4 in human monocytes and macrophages. The results are displayed as bar graphs (Fig. 2) and cumulative data are in Table I. TLR2 mRNA expression is greatest in day 1 monocytes and steadily decreases thereafter, while TLR4 mRNA expression changes to a much lesser degree during differentiation into macrophages. Expression of MR mRNA was used as a positive control because its expression is known to increase during monocyte differentiation into macrophages (4, 45).

FIGURE 2.

TLR2 steady-state mRNA levels decrease, while TLR4 levels vary to only a small extent, as monocytes differentiate into macrophages. One-, 3-, and 5- day-old PBMC in Teflon wells were harvested and adhered to a 12-well tissue culture plate in RPMI containing 10% autologous serum. After washing away lymphocytes, the monocytes or MDMs were lysed in TRIzol and total RNA was isolated. mRNA was converted to cDNA and real-time PCR was performed. TLR2 (A), TLR4 (B), and MR (C) amplification was normalized to the β-actin housekeeping gene and the RCN number was determined. A and B, Representative experiments (mean ± SD, triplicate samples) and C is cumulative data (mean ± SEM) (n = 6 (TLR2, TLR4); n = 4 (MR)). The MR was used as a positive control as a macrophage marker. One-way ANOVA with post-Tukey test. ∗, p < 0.05 compared with day 1 monocytes. ∗∗∗, p < 0.005 compared with day 1 monocytes.

FIGURE 2.

TLR2 steady-state mRNA levels decrease, while TLR4 levels vary to only a small extent, as monocytes differentiate into macrophages. One-, 3-, and 5- day-old PBMC in Teflon wells were harvested and adhered to a 12-well tissue culture plate in RPMI containing 10% autologous serum. After washing away lymphocytes, the monocytes or MDMs were lysed in TRIzol and total RNA was isolated. mRNA was converted to cDNA and real-time PCR was performed. TLR2 (A), TLR4 (B), and MR (C) amplification was normalized to the β-actin housekeeping gene and the RCN number was determined. A and B, Representative experiments (mean ± SD, triplicate samples) and C is cumulative data (mean ± SEM) (n = 6 (TLR2, TLR4); n = 4 (MR)). The MR was used as a positive control as a macrophage marker. One-way ANOVA with post-Tukey test. ∗, p < 0.05 compared with day 1 monocytes. ∗∗∗, p < 0.005 compared with day 1 monocytes.

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Table I.

TLR2 and TLR4 steady-state mRNA levels in human monocytes and macrophagesa

Receptorb% Change RCNc Compared to Day 1 Monocytes
Day 3 MonocytesDay 5 MDMs
TLR2 −51 ± 13d −65 ± 10d 
TLR4 68 ± 40 −18 ± 11 
MR 1193 ± 415e 923 ± 474e 
Receptorb% Change RCNc Compared to Day 1 Monocytes
Day 3 MonocytesDay 5 MDMs
TLR2 −51 ± 13d −65 ± 10d 
TLR4 68 ± 40 −18 ± 11 
MR 1193 ± 415e 923 ± 474e 
a

TLR2, TLR4, and MR mRNA levels were measured by real time PCR. Mean ± SEM (n = 6 for TLR2 and TLR4 and n = 4 for MR).

b

RNA was isolated and converted to cDNA and real-time PCR was performed using TLR2, TLR4, and MR TaqMan gene expression kits.

c

RCN = E−ΔCt × 100. E is the efficiency (2 = 100% efficiency) ΔCt = Ct(target) − Ct(reference).

d

p < 0.05 relative to day 1 monocytes.

e

p < 0.005 relative to day 1 monocytes.

SP-A binds to one or more receptors on macrophages (22) which triggers signaling events that alter the biology of these cells (28, 46, 47). Previous work from our laboratory has shown that SP-A increases MR expression on human MDMs, while another laboratory found that SP-A increases SR-A expression on rat macrophages (4, 5); both receptors are known PRRs. The effect(s) of SP-A on TLR expression and function in human macrophages has not been explored. We used flow cytometry to determine whether SP-A regulated the basal levels of TLR2 and TLR4 surface expression on MDMs. A typical experiment is displayed as histograms in Fig. 3, A (TLR2) and B (TLR4), with SP-A incubation for 2 h. Incubation with SP-A for 1, 6, and 20 h showed equivalent results (data not shown). Our results show that SP-A increases TLR2, but not TLR4, expression on human macrophages. We next sought to determine whether the increase in TLR2 expression is due to a unique function of SP-A and/or SP-A-specific receptor engagement or whether the function could be shared with other structurally related proteins such as the complement protein C1q which also shares some functional similarities with SP-A (28). We determined that C1q does not regulate TLR2 expression (−8 ± 9%, mean ± SD, n = 2).

FIGURE 3.

SP-A increases TLR2, but not TLR4, surface expression on MDMs. Five-day old PBMC in Teflon wells were incubated with or without SP-A (10 μg/ml) for 2 h (A and B). Following the same protocol as in Fig. 1, SP-A-treated and untreated MDMs were incubated with either a human allophycocyanin-conjugated TLR2 mAb or allophycocyanin-conjugated subtype control mAb (A) or human PE-conjugated TLR4 mAb or PE-conjugated subtype control mAb (B). The stained cells were analyzed using flow cytometry by gating on the MDMs and the average of triplicate samples is shown in this experiment which is representative of n = 4 (A) and n = 5 (B). Five-day-old MDMs in monolayer culture on glass coverslips were incubated with or without SP-A (10 μg/ml) for 2 h. After washing, cells were fixed in paraformaldehyde (no permeabilization) and stained with mouse anti-human TLR2 mAb, mouse anti-human TLR4 mAb, mouse anti-human MR mAb, or subtype control mAb followed by a secondary Alexa Fluor 488-conjugated anti-mouse Ab. Glass coverslips were mounted and visualized by confocal microscopy. A representative experiment is shown in C (n = 3 (TLR2); n = 2 (TLR4, MR)). Cumulative data are shown in D. A Student t test was performed. ∗, p < 0.05 relative to TLR2 expression on untreated MDMs. The MR was used as a positive control.

FIGURE 3.

SP-A increases TLR2, but not TLR4, surface expression on MDMs. Five-day old PBMC in Teflon wells were incubated with or without SP-A (10 μg/ml) for 2 h (A and B). Following the same protocol as in Fig. 1, SP-A-treated and untreated MDMs were incubated with either a human allophycocyanin-conjugated TLR2 mAb or allophycocyanin-conjugated subtype control mAb (A) or human PE-conjugated TLR4 mAb or PE-conjugated subtype control mAb (B). The stained cells were analyzed using flow cytometry by gating on the MDMs and the average of triplicate samples is shown in this experiment which is representative of n = 4 (A) and n = 5 (B). Five-day-old MDMs in monolayer culture on glass coverslips were incubated with or without SP-A (10 μg/ml) for 2 h. After washing, cells were fixed in paraformaldehyde (no permeabilization) and stained with mouse anti-human TLR2 mAb, mouse anti-human TLR4 mAb, mouse anti-human MR mAb, or subtype control mAb followed by a secondary Alexa Fluor 488-conjugated anti-mouse Ab. Glass coverslips were mounted and visualized by confocal microscopy. A representative experiment is shown in C (n = 3 (TLR2); n = 2 (TLR4, MR)). Cumulative data are shown in D. A Student t test was performed. ∗, p < 0.05 relative to TLR2 expression on untreated MDMs. The MR was used as a positive control.

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To complement our data with SP-A, we used confocal microscopy to visualize TLR surface expression. Consistent with the flow cytometry data, SP-A increased TLR2, but not TLR4, surface expression on MDMs (Fig. 3, C and D). MR staining was used as a positive control because SP-A increases MR surface expression on MDMs (4).

After determining that SP-A regulates TLR2, but not TLR4 surface protein expression, we next examined whether SP-A regulates TLR2 through a transcriptional mechanism. We used real-time PCR to examine whether SP-A regulates TLR2 mRNA expression in monocytes and macrophages. We examined SP-A regulation of TLR4 mRNA expression for comparison. The results are displayed as bar graphs (Fig. 4). Although SP-A decreased TLR2 mRNA expression in day 1 monocytes and increased its expression to a small extent in day 3 monocytes; importantly, it did not regulate TLR2 mRNA expression in day 5 MDMs. In contrast to its effects on TLR2 mRNA levels, SP-A had very little effect on TLR4 mRNA expression in monocytes and macrophages. Thus, the effect of SP-A on regulating TLR2 protein expression on macrophages (as opposed to monocytes), appears to be via a posttranslational mechanism.

FIGURE 4.

SP-A regulates steady-state mRNA expression of TLR2, but not TLR4, during monocyte differentiation into macrophages. SP-A (10 μg/ml) was added to 1-, 3-, and 5-day-old PBMC in selected Teflon wells for 1, 2, 6, or 20 h. PBMC were harvested and monocytes/macrophages adhered to a 12-well tissue culture plate in autologous serum. After washing away lymphocytes, the monocytes or MDMs were lysed in TRIzol and total RNA was isolated. mRNA was converted to cDNA and real-time PCR was performed. TLR2 and TLR4 mRNA amplification was normalized to the β-actin housekeeping gene. The fold change was determined by comparing SP-A-treated samples to samples with no SP-A. Cumulative data are shown (±SEM) (TLR2: n = 6 (day 5, 20 h); n = 5 (day 5, 2 h); n = 4 (day 1, 1, 2, 6, and 20 h; day 3, 1 and 2 h; day 5 1 and 6 h); n = 2 (day 3, 6 and 20 h)) (TLR4: n = 6 (day 1, 2 h); n = 5 (day 1, 1 and 20 h); n = 4 (day 1, 6 h); n = 3 (day 3, 1 and 2 h; day 5, 2 and 20 h); n = 2 (day 3, 6 h; day 5, 6 h); n = 1 (day 3, 20 h; day 5, 20 h)). Student t test was performed. ∗, p < 0.05 compared with day 1 monocytes.

FIGURE 4.

SP-A regulates steady-state mRNA expression of TLR2, but not TLR4, during monocyte differentiation into macrophages. SP-A (10 μg/ml) was added to 1-, 3-, and 5-day-old PBMC in selected Teflon wells for 1, 2, 6, or 20 h. PBMC were harvested and monocytes/macrophages adhered to a 12-well tissue culture plate in autologous serum. After washing away lymphocytes, the monocytes or MDMs were lysed in TRIzol and total RNA was isolated. mRNA was converted to cDNA and real-time PCR was performed. TLR2 and TLR4 mRNA amplification was normalized to the β-actin housekeeping gene. The fold change was determined by comparing SP-A-treated samples to samples with no SP-A. Cumulative data are shown (±SEM) (TLR2: n = 6 (day 5, 20 h); n = 5 (day 5, 2 h); n = 4 (day 1, 1, 2, 6, and 20 h; day 3, 1 and 2 h; day 5 1 and 6 h); n = 2 (day 3, 6 and 20 h)) (TLR4: n = 6 (day 1, 2 h); n = 5 (day 1, 1 and 20 h); n = 4 (day 1, 6 h); n = 3 (day 3, 1 and 2 h; day 5, 2 and 20 h); n = 2 (day 3, 6 h; day 5, 6 h); n = 1 (day 3, 20 h; day 5, 20 h)). Student t test was performed. ∗, p < 0.05 compared with day 1 monocytes.

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Once we established that SP-A differentially regulates TLR2 and TLR4 expression on macrophages, we studied whether SP-A regulates TLR function, because changes in expression do not always correlate with changes in function. When TLR2 or TLR4 is activated by an appropriate ligand, a signaling cascade is initiated that can result in the production of proinflammatory cytokines. We determined whether SP-A regulates the secretion of one of the major proinflammatory cytokines, TNF-α, in response to LPS (TLR4 ligand) or Pam3Cys (TLR2 ligand) by macrophages. We found that SP-A markedly decreased TNF-α secretion by MDMs in the presence of either ligand (Table II). Thus, although SP-A regulates the expression of only TLR2, these data suggest that SP-A can alter the biological activity of both TLR2 and TLR4.

Table II.

Effect of SP-A on TNF-α secretion from human macrophagesa

TreatmentLigand Addedb% Decrease in TNF-αcn
SP-A None 
SP-A LPS 82 ± 12d 
SP-A Pam3Cys 88 ± 13d 
TreatmentLigand Addedb% Decrease in TNF-αcn
SP-A None 
SP-A LPS 82 ± 12d 
SP-A Pam3Cys 88 ± 13d 
a

TNF-α secretion by MDMs was measured by ELISA.

b

Day 5 MDMs adhered to a 96-well tissue culture plate were incubated with or without SP-A (10 μg/ml) for 2 h and then with or without the indicated TLR ligand.

c

Values shown represent percent change in TNF-α secretion = (treatment group TNF-α secretion − control group TNF-α secretion)/control group secretion × 100). Control group is MDMs with no treatment or ligand added; mean TNF-α values (pg/ml) for LPS, Pam3Cys, SP-A plus LPS and SP-A plus Pam3Cys were 108, 108, 19 and 13, respectively.

d

p < 0.05 relative to TNF-α secretion by untreated MDMs.

Because SP-A regulates the biological outcome of TLR signaling pathways, we chose to study the effect of SP-A on the phosphorylation of specific protein kinases of these pathways, a prerequisite for kinase activity. We first examined whether SP-A regulates the phosphorylation of the inhibitory protein IκBα. IκBα is a key regulator of NF-κB activation, because it binds to the NF-κB complex in the cytosol to prevent translocation of NF-κB to the nucleus and subsequent transcription of NF-κB-regulated genes. After IκBα is phosphorylated, it is eventually degraded, which allows the NF-κB complex to translocate to the nucleus (48, 49). As shown in Fig. 5, we found that SP-A decreases the phosphorylation of IκBα in MDMs in response to TLR ligands. SP-A does not regulate the level of total IκBα protein in these cells (data not shown).

FIGURE 5.

SP-A regulates the phosphorylation of IκBα in macrophages following the addition of TLR ligands. Five-day-old MDMs were adhered to a 12-well tissue culture plate, washed, and incubated overnight in autologous serum at 37°C. Cells were incubated with or without SP-A (10 μg/ml) for 10 min. After washing, cells were incubated for an additional 10 min at 37°C to internalize any bound SP-A. LPS (10 min) or Pam3Cys (5 min) was added to appropriate wells. MDMs were lysed and SDS-PAGE and Western blots were performed using pIκBα or actin (control) Abs. Representative Western blots are shown (A and D) and bar graphs (B and E), generated by densitometric analysis, represent cumulative data (duplicate samples in each experiment; mean ± SEM; n = 4: LPS, 10-min incubation, or n = 4: Pam3Cys, 5-min incubation). C and F, Five-day-old MDMs were adhered to a 12-well tissue culture plate, washed, and incubated overnight in autologous serum at 37°C. Cells were incubated with or without SP-A (10 μg/ml) for 10 min. After washing, cells were incubated for an additional 10 min at 37°C to internalize any bound SP-A. LPS (1 h) or Pam3Cys (1 h) was added to appropriate wells, medium was removed, and nuclear extracts were prepared. p65 nuclear translocation was measured according to the manufacturer of the colorimetric TransFactor kit (mean ± SEM; n = 3). A Student t test was performed. ∗, p < 0.05 compared with LPS or Pam3Cys.

FIGURE 5.

SP-A regulates the phosphorylation of IκBα in macrophages following the addition of TLR ligands. Five-day-old MDMs were adhered to a 12-well tissue culture plate, washed, and incubated overnight in autologous serum at 37°C. Cells were incubated with or without SP-A (10 μg/ml) for 10 min. After washing, cells were incubated for an additional 10 min at 37°C to internalize any bound SP-A. LPS (10 min) or Pam3Cys (5 min) was added to appropriate wells. MDMs were lysed and SDS-PAGE and Western blots were performed using pIκBα or actin (control) Abs. Representative Western blots are shown (A and D) and bar graphs (B and E), generated by densitometric analysis, represent cumulative data (duplicate samples in each experiment; mean ± SEM; n = 4: LPS, 10-min incubation, or n = 4: Pam3Cys, 5-min incubation). C and F, Five-day-old MDMs were adhered to a 12-well tissue culture plate, washed, and incubated overnight in autologous serum at 37°C. Cells were incubated with or without SP-A (10 μg/ml) for 10 min. After washing, cells were incubated for an additional 10 min at 37°C to internalize any bound SP-A. LPS (1 h) or Pam3Cys (1 h) was added to appropriate wells, medium was removed, and nuclear extracts were prepared. p65 nuclear translocation was measured according to the manufacturer of the colorimetric TransFactor kit (mean ± SEM; n = 3). A Student t test was performed. ∗, p < 0.05 compared with LPS or Pam3Cys.

Close modal

After determining that SP-A decreases the phosphorylation of IκBα, we next examined whether SP-A directly regulates the nuclear translocation of the NF-κB complex. We observed that SP-A significantly decreases the nuclear translocation of p65, a major component of the NF-κB complex, in the presence of TLR ligands (Fig. 5).

A recent report has shown the ability of SP-A to activate SHP-1, a tyrosine phosphatase, and thereby decrease the phosphorylation of p38, one member of the MAPK family, in response to LPS (28). Thus, we sought to determine whether SP-A regulates the phosphorylation of each of the MAPKs in the presence of TLR ligands in human macrophages. These results would provide further insight as to whether SP-A directly regulates NF-κB activity or whether the effect is upstream of this transcriptional complex. We found that while SP-A decreases the phosphorylation of p38 and ERK, it does not affect the phosphorylation of JNK in MDMs, as assayed by Western blot analysis (Fig. 6).

FIGURE 6.

SP-A selectively regulates the phosphorylation of MAPKs in macrophages following the addition of TLR ligands. Following the same protocol as in Fig. 5, treated MDMs were lysed and SDS-PAGE and Western blots were performed using pERK (A and D), pp38 (B and E), pJNK (C and F), or actin (control) Abs. Representative Western blots are shown and bar graphs, generated by densitometric analysis, represent cumulative data (duplicate samples in each experiment; mean ± SEM; n = 4 for pERK, LPS and Pam3Cys 15-min incubation; n = 4 for pp38, LPS and Pam3Cys 15-min incubation; and n = 3 for pJNK, LPS and Pam3Cys 10-min incubation). A Student t test was performed. ∗, p < 0.05 compared with LPS and Pam3Cys.

FIGURE 6.

SP-A selectively regulates the phosphorylation of MAPKs in macrophages following the addition of TLR ligands. Following the same protocol as in Fig. 5, treated MDMs were lysed and SDS-PAGE and Western blots were performed using pERK (A and D), pp38 (B and E), pJNK (C and F), or actin (control) Abs. Representative Western blots are shown and bar graphs, generated by densitometric analysis, represent cumulative data (duplicate samples in each experiment; mean ± SEM; n = 4 for pERK, LPS and Pam3Cys 15-min incubation; n = 4 for pp38, LPS and Pam3Cys 15-min incubation; and n = 3 for pJNK, LPS and Pam3Cys 10-min incubation). A Student t test was performed. ∗, p < 0.05 compared with LPS and Pam3Cys.

Close modal

Having determined that SP-A regulates the phosphorylation of IκBα and members of the MAPK family, we sought to identify a novel upstream target of SP-A’s activity that is known to regulate both NF-κB and MAPKs. The protein kinase Akt has been shown to both positively and negatively regulate NF-κB and cytokine production, depending on the stimulus, cell type, and activation state of the cell (34, 50, 51, 52, 53, 54, 55). Akt can also regulate the MAPK pathway and thereby affect the activation of AP-1 and NF-κB (51). In support of this possibility, we had previously determined that PI3K activity is involved in the up-regulation of MR expression by SP-A (33). PI3Ks are activated either directly through TLR2 or through MyD88 for the TLR4 pathway (55, 56, 57). PI3K activation can lead to the phosphorylation and activation of Akt (58, 59, 60). Therefore, we chose to study whether SP-A regulates the phosphorylation of Akt. Using Western blots, we found that SP-A decreased the phosphorylation of Akt in the presence of LPS or Pam3Cys in MDMs (Fig. 7). Total Akt expression was not affected by the presence of SP-A (data not shown).

FIGURE 7.

SP-A regulates the phosphorylation of Akt in human macrophages following the addition of TLR ligands. Five-day-old MDMs were adhered to a 12-well tissue culture plate, washed, and incubated overnight in autologous serum at 37°C. Cells were incubated with or without SP-A (10 μg/ml) for 10 min. After washing, cells were incubated for an additional 10 min at 37°C to internalize any bound SP-A. LPS (30 min) or Pam3Cys (15 min) was added to appropriate wells. MDMs were lysed and SDS-PAGE and Western blots were performed with lysates using pAkt Ab or actin Ab (control). Representative Western blots are shown (A and C) and bar graphs (B and D), generated by densitometric analysis, represent cumulative data (duplicate samples in each experiment; mean ± SEM; n = 3, LPS; n = 4, Pam3Cys). A Student t test was performed. ∗, p < 0.05 compared with LPS or Pam3Cys.

FIGURE 7.

SP-A regulates the phosphorylation of Akt in human macrophages following the addition of TLR ligands. Five-day-old MDMs were adhered to a 12-well tissue culture plate, washed, and incubated overnight in autologous serum at 37°C. Cells were incubated with or without SP-A (10 μg/ml) for 10 min. After washing, cells were incubated for an additional 10 min at 37°C to internalize any bound SP-A. LPS (30 min) or Pam3Cys (15 min) was added to appropriate wells. MDMs were lysed and SDS-PAGE and Western blots were performed with lysates using pAkt Ab or actin Ab (control). Representative Western blots are shown (A and C) and bar graphs (B and D), generated by densitometric analysis, represent cumulative data (duplicate samples in each experiment; mean ± SEM; n = 3, LPS; n = 4, Pam3Cys). A Student t test was performed. ∗, p < 0.05 compared with LPS or Pam3Cys.

Close modal

AM recognize and remove microbes through increased activity of a subset of PRRs. Two major PRRs are TLR2 and TLR4 which, depending on the stimulus, initiate proinflammatory responses or these inflammatory responses can be inhibited by the activation of TLR negative regulators. TLR2 and TLR4 initiate an inflammatory immune response by binding to a wide variety of pathogen-associated molecular patterns (61, 62) through the extracellular leucine-rich repeat domains of these receptors which lead to the activation of the TLR cytoplasmic signaling domains. TLR2 and TLR4 signaling can indirectly activate the adaptive immune response through the production of inflammatory cytokines which, in turn, can lead to dendritic cell maturation and migration to lymph nodes where Ag-derived peptide presentation to T cells occurs (63).

We used our primary human monocyte/macrophage differentiation cell model to address several unanswered questions with regard to TLR2 and TLR4 expression and activity on these cells. We were able to discern important differences in the transcriptional and posttranslational mechanisms for TLR2 and TLR4 expression between monocytes and macrophages, both with respect to basal levels and following SP-A stimulation. Furthermore, our studies have identified a new signaling pathway targeted by SP-A, i.e., involvement of the major cell regulator Akt, members of the MAPK family, and a key downstream regulator of NF-κB, IKBα, ultimately leading to a marked reduction in the production of TNF-α.

There are important differences between human and murine TLRs with regard to steady-state mRNA levels, surface protein expression, protein structure, and regulation. Some of the TLR2 differences include: little homology between human and murine promoter regions, expression on murine but not human T cells, and different regulatory elements controlling the expression of the murine and human TLR2 genes (41). Similarly, the TLR4 human and murine genes are regulated differently and murine TLR4 mRNA is more broadly expressed than human TLR4 mRNA (35, 64). Also, although there is strong sequence homology between the human and murine TLR4 promoters, there are differences in the structures of these promoters, which could account for the differences in expression (35). Since the majority of TLR studies involve murine cell lines and primary murine cells, we chose to focus our attention on whether human monocytes and MDMs expressed TLR2 and TLR4.

We observed the highest TLR2 surface protein and mRNA expression in day 1 monocytes and this expression decreased as monocytes differentiated into macrophages. Unlike TLR2, TLR4 surface protein expression stayed constant and TLR4 mRNA expression varied only slightly as monocytes differentiated into macrophages. The differences in basal surface protein expression and mRNA steady-state levels between TLR2 and TLR4 suggested that they may also be differentially regulated by SP-A. Our TLR2 mRNA data are consistent with a previous finding by Haehnel et al. (41) where TLR2 mRNA expression (Northern blot analysis) in monocytes decreased after 24 h of adherence to a tissue culture plate. Although others have shown TLR2 and TLR4 mRNA expression in human monocytes or MDMs (38, 44, 65), these studies did not measure mRNA levels during monocyte differentiation into macrophages. Our cell surface protein expression data are consistent with a previous report that showed more TLR2 and TLR4 surface protein expression on human monocytes compared with human AM (66).

We found that while SP-A up-regulated TLR2 surface protein expression, it did not affect TLR4 surface expression on macrophages. Interestingly, SP-A decreased TLR2 mRNA expression in day 1 monocytes, but increased to a small extent (not significant) mRNA expression in day 3 cells. Importantly, SP-A did not regulate TLR2 mRNA expression in day 5 macrophages. These data show that SP-A differentially regulates TLR2 mRNA expression during monocyte differentiation into macrophages. Due to the lack of TLR2 mRNA regulation by SP-A in day 5 MDMs, our data support that SP-A up-regulation of TLR2 surface expression on MDMs is most likely the result of a posttranslational mechanism, possibly by translocating intracellular pools of preformed TLR2 to the cell surface. Confocal microscopy experiments using fixed and permeabilized MDMs suggest that intracellular pools of TLR2 exist (our unpublished observation). Our previous work indicated a similar mechanism for SP-A’s up-regulation of the MR on macrophages (4) and there are reports that TLR2 and TLR4 can be found within intracellular vesicles (67, 68).

We examined the relationship between SP-A’s effects on TLR expression and its effects on TLR function because effects on expression and function do not always correlate. There are contradictory data regarding the role of SP-A and TLRs in the inflammatory response. By using CHO cells and TLR4-deficient/wild-type mice, one group observed that SP-A can activate the NF-κB signaling pathway and up-regulate the synthesis of cytokines, such as TNF-α, by relying predominantly on a functional TLR4 complex (69), while another group did not observe SP-A activation of NF-κB (70). Differences in these results may depend on the SP-A purification process used by the researchers. Purified SP-A used by the first group contained higher concentrations of LPS compared with that of the second group. This difference in LPS contamination could result in the proinflammatory effects by SP-A which have been observed (71).

We observed a significant reduction, but not a complete elimination, of TNF-α secretion when macrophages were pretreated with SP-A and then treated with either the TLR4 ligand, LPS, or the TLR2 ligand Pam3Cys. Our TLR4 data are consistent with previous publications that showed that SP-A decreases TNF-α secretion in the presence of LPS (28, 72, 73, 74). Overall, these data provide further evidence for the role of SP-A in dampening the proinflammatory response in the lung environment.

Recently, SP-A was shown to directly associate with TLR2 and TLR4 proteins and thereby attenuate a proinflammatory response in rat macrophages and murine cell lines (46, 75). In our studies, we used a protocol which allows for complete internalization of SP-A in MDMs before adding the TLR ligands (33). Following incubation with SP-A, we used specific mAbs to readily detect the TLRs (increased in the case of TLR2), indicating that surface TLR binding sites were not occupied by SP-A and were free to bind other ligands. Thus, our studies point to a different mechanism independent of TLR blockade by SP-A, whereby SP-A binds to one or more of its receptors, initiating a signaling cascade that results in altered expression and activity of the TLRs. Several potential SP-A receptors have been identified, including signal regulatory protein α, calreticulin/CD91 (28) and SP-R210 (76, 77).

We found that SP-A significantly decreased the phosphorylation of IκBα in macrophages following the addition of the TLR2 and TLR4 ligands. Our data contradict previous findings by Wu et al. (70), which did not find an effect of SP-A on the phosphorylation of IκBα in the presence of LPS. The differences could be due to the low concentration of LPS that was used to stimulate the cells and the difference in cell type. Additionally, our data show that SP-A directly regulates the NF-κB complex by decreasing the nuclear translocation of p65.

We found that SP-A decreases the phosphorylation of p38 and ERK, but not JNK in the presence of TLR ligands. SP-A regulation of the phosphorylation of p38 in the presence of LPS are consistent with previous findings by Gardai et al. (28) using murine RAW cells and human macrophages. However, they reported that SP-A also regulated the phosphorylation of JNK, although the data were not shown. If RAW cells were used for their pJNK experiments, the differences in our findings could be explained by their use of a murine macrophage-like cell line vs our human MDMs. Our data suggest that JNK is regulated by distinct mechanisms compared with p38 and ERK. Although the p38, ERK, and JNK pathways can be activated by unique upstream signaling molecules, there are overlapping upstream and downstream kinases, which could explain why we observed that SP-A regulated only two of the three MAPKs (78).

We previously found that PI3K was involved in SP-A’s increase in MR expression on macrophages (33). Also, previous studies have shown that when TLR2 or TLR4 is activated by appropriate ligands, PI3Ks are activated (79). The activation of PI3K initiates a signaling cascade that results in the phosphorylation and activation of Akt. Akt is a kinase that is involved in numerous cell functions, one of which is regulation of NF-κB (80, 81, 82). The literature provides evidence for a potential link among Akt, MAPKs, and the activation of AP-1 and NF-κB (51). We found that SP-A decreased the phosphorylation of Akt in the presence of Pam3Cys and LPS, which is associated with the decrease in phosphorylation of MAPKs and IκBα, finally leading to a diminished proinflammatory response.

Based on our previous work and current work and the literature, we propose a model of TLR modulation by SP-A (Fig. 8), where SP-A binds to its macrophage receptor(s) and signals TLR2 intracellular pools to travel to the cell surface, which increases TLR2 surface expression. Simultaneously, SP-A signals also regulate TLR activity by diminishing proinflammatory cytokine production as the result of a decrease in the phosphorylation of a key regulator of NF-κB, IκBα, and a decrease in the nuclear translocation of p65. SP-A down-regulates kinases upstream of IκBα by decreasing the phosphorylation of Akt and MAPKs in response to TLR ligands. Therefore, while SP-A increases TLR2 expression, which can enhance pathogen recognition, it limits TLR signaling so that the lung is not damaged by an overreactive inflammatory response.

FIGURE 8.

SP-A is a key regulator of TLR expression and signaling in macrophages. SP-A binds to its receptor(s) leading to increased expression of TLR2 but not TLR4 (this report), the MR (∗) (4 ) and SR-A (∗∗) (5 ). Simultaneously, SP-A regulates TLR activity in response to agonists by decreasing the phosphorylation of key TLR signaling proteins, including Akt and MAPKs. Finally, SP-A decreases the phosphorylation of IκBα and nuclear translocation of p65 which results in diminished proinflammatory cytokine production.

FIGURE 8.

SP-A is a key regulator of TLR expression and signaling in macrophages. SP-A binds to its receptor(s) leading to increased expression of TLR2 but not TLR4 (this report), the MR (∗) (4 ) and SR-A (∗∗) (5 ). Simultaneously, SP-A regulates TLR activity in response to agonists by decreasing the phosphorylation of key TLR signaling proteins, including Akt and MAPKs. Finally, SP-A decreases the phosphorylation of IκBα and nuclear translocation of p65 which results in diminished proinflammatory cytokine production.

Close modal

There is accumulating evidence that SP-A regulates the macrophage surface expression and/or function of a subset of PRRs including the MR, SR-A, and, based on this report, TLR2 and TLR4, and controls the host inflammatory response and microbicidal activity against various pathogens and particulates routinely encountered in the lung (Fig. 8). This is a critical function of SP-A that preserves the normal architecture of the alveolar environment and gas exchange and provides for effective microbial clearance and sterility of the alveolar environment.

We thank Dr. Mark Wewers for performing bronchoscopies to obtain BAL for the human alveolar macrophage experiments. Also, we thank Dr. Francis McCormack and Dr. Bruce Trapnell for providing BAL from alveolar proteinosis patients for SP-A isolation and purification. We acknowledge the support of two core facilities at the Ohio State University, the Dorothy M. Davis Heart and Lung Research Institute Flow Cytometry Core Laboratory, and the Campus Microscopy and Imaging Facility.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by National Institutes of Health Grant AI059639 (to L.S.S.).

3

Abbreviations used in this paper: AM, alveolar macrophage; MDM, monocyte-derived macrophage; MR, mannose receptor; SP-A, surfactant protein A; Pam3Cys, Pam3Cys-Ser-(Lys)4 hydrochloride; PRR, pattern recognition receptor; BAL, bronchoalveolar lavage; APP, alveolar proteinosis patient; MFI, mean fluorescence intensity; RCN, relative copy number; Ct, cycle threshold; SR-A, scavenger receptor A.

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