Systematic Analysis of Cell-Type Differences in the Epithelial Secretome Reveals Insights into the Pathogenesis of Respiratory Syncytial Virus–Induced Lower Respiratory Tract Infections

Human respiratory syncytial virus (RSV) is a ubiquitous paramyxovirus pathogen that infects virtually all children by the age of 3, causing significant morbidity (1, 2). The presence of RSV infection produces three times the risk for subsequent hospitalization over that seen in infections with other common cold viruses, and hospitalization rates as high as 17 per 1000 cases are seen in prospective epidemiological studies on young children (3).

RSV infections produce a variety of clinical syndromes, including upper respiratory tract infections (with or without recurrent otitis media) and lower respiratory tract infections (LRTIs) (1, 4). Although the majority of infections produce an uncomplicated upper respiratory infection, in ∼2% of predisposed children RSV can spread into the lower airways, producing an LRTI (1, 5, 6). Consequently, RSV is the most common cause of childhood LRTIs (7) and is responsible for 120,000 hospitalizations for LRTI (bronchiolitis) in the United States annually (8), and it represents the leading cause of infant viral death worldwide (9). In addition to its acute morbidity, severe LRTIs are associated with reshaping the pulmonary immune response, producing Th2 polarization and enhancing susceptibility to recurrent virus–induced wheezing through the next several decades of life (10). The mechanisms by which LRTIs reprogram the pulmonary immune system are not fully understood.

As a mucosa-restricted virus, in natural infections RSV initially replicates in the epithelium of the nasopharynx. In immunologically naive infants, RSV spreads by cell–cell transfer and extracellular binding, producing discontinuous foci of infection in the tracheal epithelium (11). Further spread of RSV infection to distal bronchioles and the alveolar epithelium induces necrosis and sloughing. Sloughed epithelial cells are encased in fibrin and mucin, producing a ball-valve obstruction and ventilation–perfusion mismatching, impairing gas exchange and causing hypoxia, one of the most significant clinical features of LRTIs. The mechanism by which RSV produces epithelial sloughing was recently elucidated in part: the intracellular expression of RSV nonstructural protein 2 disrupts ciliary tubulin IV, obliterating epithelial ciliary function and inducing detachment from the basal lamina (12).

Because RSV LRTIs occur in immunologically naive infants, the innate immune response plays an initiating role in LRTI pathogenesis. It was extensively demonstrated that RSV replication in epithelial cells triggers a coordinated global genomic response that results in epithelial secretion of mucins, defensins, antiviral cytokines, and proinflammatory chemokines that coordinate inflammation and adaptive immunity (13–15). Enhanced activation of innate inflammation was demonstrated in children with bronchiolitis, in whom enhanced expression of IL-6, IL-8, and macrophage inhibitory protein (MIP)-1β were observed (16). Important to the pathogenesis of LRTIs, the epithelium in the proximal conductive airways and the respiratory surface play functionally distinct roles in innate defenses (17, 18). Under normal conditions, inhaled pathogens are cleared via the mucociliary escalator from ciliated epithelial cells. This defense is coordinated with the actions of the airway lining fluid, which is rich in antioxidants, defensins, and lysozyme secreted by Clara cells and submucosal glands, and that of mucous glycoprotein secreted by goblet cells (19). When these conductive epithelial cells are infected with RSV, they secrete CXCL chemokines MIP-1 and type I/III IFNs. These latter cytokines activate resident plasmacytoid dendritic cells (DCs) to upregulate CD40, CD80, and CD86 costimulatory molecules, inducing their migration to draining lymph nodes to stimulate cytotoxic T lymphocytes, which include NK and CD4/8⁺ memory T cells (17).

In contrast, when nonciliated epithelial cells located in the conductive airways (terminal bronchioles and alveoli) are infected with RSV, they express the CCL-type cytokines (TARC, MCP-1, and MDC), type I/III IFNs, and surfactants (15). These factors activate alveolar macrophages and pulmonary DCs (pDCs) to induce neutrophil recruitment and promote macrophage survival. Gene-profiling experiments suggested that these CCL-type chemokines are produced to a greater degree than by epithelial cells of the conducting airways (20). These findings suggest the presence of distinct regional, cell type–specific differences between activated conductive versus respiratory epithelial cells that may be immunologically important (15, 20). Despite this knowledge, there has been no systematic analysis of secreted proteins that would provide an understanding of the distinct mechanisms of innate immunity in different regions of the pulmonary tree.

To address this deficiency, we undertook a systematic analysis of RSV-induced secreted proteins in two principal cell types of the airways: human bronchial epithelial cells (hBECs), derived from the trachea and representing the conducting portion of the airway and nonciliated human small airway epithelial cells (hSAECs), representing nonciliated cells from terminal bronchioles that play a role in lower airway obstruction in RSV LRTIs (21). We first standardized a workflow by analysis of control and RSV-induced conditioned medium (CM) in human telomerase (Tert)-immortalized hSAECs and hBECs using quantitative label-free mass spectrometry (MS). Our analysis was highly reproducible and identified distinct patterns of induced and inhibited proteins. Interestingly, exosomes constituted a significant fraction of the secretome; their protein contents also differed by cell type and were affected by RSV infection. We extended this workflow to analyze multiple nonimmortalized primary cells from independent donors. Strikingly, hSAECs showed enhanced expression of immunologically important chemokines: CCL20/MIP3α, thymic stromal lymphopoietin (TSLP), and CCL3-like 1 (CCL3-L1). We demonstrate that CCL20/MIP3α is the most active mucin-inducing factor in RSV CM from hSAECs and discuss the implications of regional differences in the epithelial secretome for the pathogenesis of LRTIs.

Immortalized hBECS (tert-hBECs) and hSAECs (tert-hSAECs) were established by transducing primary cells with human telomerase and cyclin-dependent kinase-4 retrovirus constructs (22, 23). hBECs and hSAECs were grown in basal medium supplemented with growth factors (Lonza, Walkersville, MD) in 10-cm petri dishes in a humidified incubator with 95% air/5% CO₂ at 37°C. At 80–90% confluence, the medium was changed, fresh basal medium without growth supplements was added to the plates, and the cells were infected with sucrose cushion–purified RSV (pRSV) (multiplicity of infection [MOI] = 1.0) for 24 h. CM was collected and centrifuged at 2000 × g at 4°C for 20 min to remove any dead cells. The supernatant was centrifuged at 10,000 × g at 4°C for 10 min to remove any cell debris. The supernatant was used immediately for secretome analysis. Cells from the same plates were lysed in TRIzol Reagent for whole-cell protein preparation. Experiments were performed in biological triplicates.

For studies with primary hBECs (phBECs) and primary hSAECs (phSEACs), cells from three donors were obtained from Lonza (Supplemental Table I). CM was prepared from hBECs or hSAECs 24 h postinfection (MOI = 1.0). When indicated, CM for UV-inactivated RSV-infected cells was used to stimulate hBECs at a 1–25% (v/v) concentration for the indicated times. UV inactivation was as previously described (24). For Ab neutralization, 20 μl of RSV-CM was mixed with anti-CCL20 Ab (R&D Systems, Minneapolis, MN).

Exosome isolation was performed by differential centrifugation at +4°C to minimize protein degradation. Cells were removed by low-speed centrifugation at 400 × g for 10 min. The cleared supernatant was sequentially centrifuged at 2000 × g for 15 min and 10,000 × g for 30 min to remove any remaining cell debris/microvesicles. Exosomes were finally pelleted by ultracentrifugation at 100,000 × g for 2 h and washed in PBS (without Ca²⁺ or Mg²⁺) at 100,000 × g for 60 min. After washing, the pellet was resuspended in a total of 200 μl of PBS. Exosome size was estimated by dynamic light scattering using a Malvern High-Performance Particle Sizer (Malvern Instruments, Westborough, MA). Data acquisition and analysis were performed using Dispersion Technology Software (V4.1.26.0) configured for HPPS analysis. Each experimental group had three independent replicates.

The human RSV A2 strain was grown in Hep-2 cells and prepared by sucrose cushion centrifugation, as described (25). The viral titer was determined by a methylcellulose plaque assay. pRSV aliquots were quick-frozen in dry ice–ethanol and stored at −70°C until use.

Airway cells were plated on cover glasses pretreated with rat tail collagen (Roche Applied Sciences). The cells were fixed with 4% paraformaldehyde in PBS, permeabilized with 0.1% Triton X-100, blocked in 10% goat serum, and incubated with primary rabbit polyclonal Ab to cytokeratin-7 or -19, as indicated (26, 27). After incubation with Alexa Fluor–goat anti-Rb Ab, cells were washed and counterstained with DAPI. The cells were visualized with a Zeiss fluorescence LSM510 confocal microscope using a 63× objective.

Immunofluorescence (IF) of paraffin-embedded mouse lung was performed following standard protocols. Briefly, lung sections (5 μm) were deparaffinized in xylene and hydrated in ethanol (100–70%). Sections were washed with deionized water. Ag unmasking was done with 1.0 mM EDTA (pH 8). Sections were washed in H₂O and blocked with 10% goat serum for 1 h at room temperature in the dark, followed by primary Ab against CCL20 (1:200 dilution; Abcam) overnight at 4°C in the dark. Sections were washed with TBS-Tween (0.1%) buffer and incubated with goat secondary Ab conjugated with Alexa Fluor 568 (DAKO) for 1 h at room temperature in the dark. Sections were washed in TBS-Tween (0.1%), incubated with DAPI for 1 min, mounted on coverslips, visualized by confocal microscopy (Zeiss), and photographed at ×63 magnification.

Apoptosis was measured using a commercial Annexin V–FITC Apoptosis Kit, following the manufacturer’s protocol (BioVision, Milpitas, CA). Briefly, hBECs and hSAECs (1 × 10⁶) were infected or not with RSV (MOI = 1.0) for 24 h. Cells were dislodged with Accutase (Millipore), washed once with PBS, and incubated with 5 μl of Annexin V–FITC and 5 μl of propidium iodide (PI) in 500 μl of binding buffer for 5 min at room temperature in the dark. Annexin V–FITC and PI labeling was measured by flow cytometry.

A 10-μl aliquot from the exosome suspension was diluted in deionized water, applied to 200 mesh Formvar/carbon-coated copper grids (Electron Microscopy Sciences) for 10 min at room temperature (24°C), and negatively stained with 2% uranyl acetate. The grids were examined with a Philips CM-100 transmission electron microscope at 60 kV FEI (Thermo Fisher Scientific, San Jose, CA). Exosome images were acquired with a Gatan Orius 2001 charge-coupled device camera.

About 10 ml of the CM supernatant was added into a 3K filter unit (Millipore, Billerica, MA) and centrifuged at 14,000 × g for 15 min. Next, 400 μl of 8 M urea was added to the filter unit and centrifuged at 14,000 × g for 15 min. The filter was washed again using the same procedure. The solution remaining in the filter device was collected for protein digestion. Proteins were reduced with 10 mM DTT for 30 min, followed by alkylation with 30 mM iodoacetamide for 60 min in the dark. The sample was diluted 1:1 with 50 mM ammonium bicarbonate. Proteins were digested with 1.0 μg of Trypsin/Lys-C (Promega) for 12 h at 37°C and then diluted 4:1 with 50 mM ammonium bicarbonate. The proteins were further digested with 1.0 μg trypsin (Promega) for 16 h at 37°C. The digestion was stopped with 0.5% trifluoracetic acid (TFA), and the peptides were desalted on a reversed-phase Sep-Pak C18 cartridge (Waters); peptides were eluted using 80% acetonitrile (ACN). The eluate was dried in a SpeedVac, and the peptides were acidified with 2% ACN–0.1% TFA.

About 50 μg of proteins in 8 M guanidine was reduced with 10 mM DTT, alkylated with 30 mM iodoacetamide, and sequentially digested with 1.0 μg of Trypsin/Lys-C and 1.0 μg of trypsin, as described above, for secretome proteins.

The proteins present in the exosomes were separated from the lipid components by chloroform/methanol precipitation (28). After resuspension of the chloroform/methanol precipitation pellet in 45 μl of 8 M guanidine, proteins were reduced with DTT, alkylated with iodoacetamide, and sequentially digested with Trypsin/Lys-C, as described above.

A nanoflow ultra-HPLC instrument (EASY-nLC) was coupled on-line to a Q Exactive mass spectrometer with a nanoelectrospray ion source (all from Thermo Fisher Scientific). Peptides were loaded onto a C18 reversed-phase column (25 cm long, 75 μm inner diameter) and separated with a linear gradient of 5–35% buffer B (100% ACN in 0.1% formic acid) at a flow rate of 300 nl/min over 240 min. MS data were acquired using a data-dependent Top15 method dynamically choosing the most abundant precursor ions from the survey scan (400–1400 m/z) using higher-energy collisional dissociation fragmentation. Survey scans were acquired at a resolution of 70,000 at m/z 400. Unassigned precursor ion charge states, as well as singly charged species, were excluded from fragmentation. The isolation window was set to 3 kDa and fragmented with a normalized collision energy of 27. The maximum ion injection times for the survey scan and the tandem MS (MS/MS) scans were 20 and 60 ms, respectively, and the ion target values were set to 1e6 and 1e5, respectively. Selected sequenced ions were dynamically excluded for 30 s. Data were acquired using Xcalibur software (Thermo Fisher Scientific).

Mass spectra were analyzed with MaxQuant software version 1.5.2.8 using the Andromeda search engine (29, 30). The initial maximum allowed mass deviation was set to 10 ppm for monoisotopic precursor ions and to 0.5 kDa for MS/MS peaks. Enzyme specificity was set to trypsin, defined as C-terminal to arginine and lysine excluding proline, and a maximum of two missed cleavages was allowed. Carbamidomethyl cysteine was set as a fixed modification, and N-terminal acetylation and methionine oxidation were set as variable modifications. The spectra were searched with the Andromeda search engine against the Human and RSV Swiss-Prot sequence database (containing 20,193 human protein entries and 11 RSV protein entries) combined with 248 common contaminants and concatenated with the reversed versions of all sequences. Protein identification required at least one unique or razor peptide per protein group. Quantification in MaxQuant was performed using the built-in XIC-based label-free quantification (LFQ) algorithm (30). The required false discovery rate (FDR) for identification was set to 1% at the peptide and protein levels, and the minimum required peptide length was set to 6 aa. Contaminants, reverse identification, and proteins only identified by site were excluded from further data analysis. The raw data and database search results were deposited in ProteomeXchange (http://www.proteomexchange.org/) under Project Accession Number PXD005814. For comparative analysis, the LFQ values were log2 transformed. After filtering (at least two valid LFQ values in at least one group), the remaining missing LFQ values were inputted from a normal distribution (width: 0.3; down shift: 1.8). Significance analysis of microarrays (SAM) was used to assess the statistical significance of protein abundances using 1% FDR adjustment and a 2-fold cutoff (31).

The normalized spectral abundance factor (NSAF) value for each protein was calculated as

where the total MS intensity (I) of the matching peptides from protein k was divided by the protein length (L) and then by the sum of I/L for all uniquely identified proteins in the dataset (32).

For pairwise comparisons, missing NSAF values for proteins that were only present in either CM or the whole-cell lysate (WCL) were inputted from a normal distribution (width: 0.3; down shift: 1.8). We used the Perseus bioinformatics platform (http://141.61.102.17/perseus_doku/doku.php?id=start) for principal component analysis (PCA), unsupervised hierarchical clustering, gene ontology (GO) annotation enrichment, and the Fisher exact test. We used Ingenuity Pathway Analysis for upstream regulator analysis. Gene set enrichment analysis was performed by quantifying canonical pathway enrichment (33). Exosome analyses were performed by searching the ExoCarta exosome database (http://www.exocarta.org/) (34).

The stable isotope dilution (SID)-selected reaction monitoring (SRM)-MS assays were developed as described previously (35). The peptides were chemically synthesized to incorporate isotopically labeled [¹³C₆¹⁵N₄] arginine or [¹³C₆¹⁵N₂] lysine to a 99% isotopic enrichment (Thermo Fisher Scientific). The secretome and cellular proteome were digested as described above. The tryptic digests were reconstituted in 30 μl of 5% formic acid/0.01% TFA. An aliquot of 10 μl of diluted stable isotope–labeled standard (SIS) peptides was added to each tryptic digest. These samples were desalted with a ZipTip C18 cartridge; the peptides were eluted with 80% ACN, dried, reconstituted in 30 μl of 5% formic acid/0.01% TFA, and analyzed directly by liquid chromatography (LC)-SRM-MS using a TSQ Vantage triple quadrupole mass spectrometer equipped with a nanospray source (Thermo Fisher Scientific). Online chromatography was performed using an Eksigent NanoLC-2D HPLC system (AB SCIEX, Dublin, CA). An aliquot of 10 μl of each tryptic digest was injected onto a C18 reversed-phase nano-HPLC column (PicoFrit, 75 μm × 10 cm; tip inner diameter 15 μm) at a flow rate of 500 nl/min. Column was eluted by 20 min of 98% mobile phase A (0.1% formic acid), followed by a 15-min linear gradient from 2 to 30% mobile phase B (0.1% formic acid/90% ACN). The TSQ Vantage was operated in high-resolution SRM mode with Q1 and Q3 set to 0.2- and 0.7-kDa full width half maximum. All acquisition methods used the following parameters: 1800 V ion spray voltage, a 275°C ion transferring tube temperature, a collision-activated dissociation pressure of 1.5 mTorr, and the S-lens voltage used the values in the S-lens table generated during MS calibration.

All SRM data were manually inspected to ensure peak detection and accurate integration. The chromatographic retention time and the relative product ion intensities of the analyte peptides were compared with those of the SIS peptides. The variation in the retention time between the analyte peptides and their SIS counterparts should be within 0.05 min, and no significant differences in the relative product ion intensities of the analyte peptides and SIS peptides were observed. The peak areas in the extract ion chromatography of the native and SIS version of each signature peptide were integrated using Xcalibur 2.1. The default values for noise percentage and baseline subtraction window were used. The ratios between the peak area of the native and SIS versions of each peptide were calculated.

For gene-expression analyses, 1 μg of RNA was reverse transcribed using Super Script III in a 20-μl reaction mixture (35). One microliter of cDNA product was diluted 1:2, and 2 μl of diluted product was amplified in a 20-μl reaction mixture containing 10 μl of SYBR Green Supermix (Bio-Rad) and 0.4 μM each of forward and reverse gene-specific primers (Table I). The reaction mixtures were aliquoted into a Bio-Rad 96-well clear PCR plate, and the plate was sealed with Bio-Rad Microseal B film before insertion into the PCR machine. The plates were denatured for 90 s at 95°C and then subjected to 40 cycles of 15 s at 94°C, 60 s at 60°C, and 1 min at 72°C in an iCycler (Bio-Rad). PCR products were subjected to melting curve analysis to assure that a single amplification product was produced. Quantification of relative changes in gene expression was calculated using the ΔΔCt method (27). Data were expressed as fold change mRNA normalized to cyclophilin or PolB mRNA abundance, as indicated as an internal control.

BALB/c mice (Harlan) were inoculated intranasally with 50 μl of pRSV (final inoculum, 10⁷ PFU) diluted in PBS under light anesthesia, as previously described (36). Twenty-four hours later, the animals were euthanized, and their lungs were fixed in paraformaldehyde for immunohistochemical analysis. Sections were prepared and stained with H&E or periodic acid–Schiff (PAS; Abcam) using standard techniques.

In the first experiment, we examined the effects of RSV on secreted proteins in hBECs derived from the trachea compared with hSAECs derived from the terminal bronchioles. Because tert-immortalized cells maintain a highly differentiated epithelial phenotype over continuous passages, we used these cells for establishing and validating a secretome analysis pipeline. tert-hBECs form pseudostratified ciliated monolayers on air–liquid interfaces (22). In contrast, tert-hSAECs express high amounts of p63 aldehyde dehydrogenase and differ from tert-hBECs by their resistance to naphthalene injury, a characteristic of bronchiolar cells (37).

Our previous work showed that RSV effectively replicates in both cell types, expressing cytokines, producing infectious virions, and inducing syncytia formation (38, 39). To directly compare the levels of RSV replication, tert-hBECs and tert-hSAECs were infected with pRSV (MOI = 1, 24 h). Cells were lysed, and the expression of RSV nucleoprotein, matrix protein, phosphoprotein, and matrix M2-1 was determined by LC-MS/MS (Fig. 1A). The levels of RSV expression for all proteins measured were dramatically elevated in tert-hBECs and tert-hSAECs relative to uninfected cells. Interestingly, RSV protein replication was 4-fold higher in tert-hBECs than in tert-hSAECs (Fig. 1A). Similar results were observed in the CM from each cell type (although the protein abundance was lower as a result of medium dilution), indicating viral secretion (Fig. 1B).

To determine the effects of RSV infection on cell viability, apoptosis and necrosis rates for control and RSV-infected tert-hBECs and tert-hSAECs (MOI = 1, 24 h) were measured by flow cytometry. Cellular necrosis was minimal in both cell types in the absence or presence of RSV infection. However, in the absence of infection, tert-hBECs had a higher basal apoptotic rate than did tert-hSAECs (18.8 versus 10.8%, Fig. 1C). In both cell types, RSV infection reduced the apoptosis rate. The apoptotic rate of tert-hBECs fell from 18.8 to 12.5% and that of tert-hSAECs decreased from 10.8 to 7.9% (Fig. 1C). These data are consistent with earlier studies demonstrating that cell death is not a major initial effect of RSV infection (40) and that the apoptosis rate is reduced upon RSV infection as a result of NF-κB activation by the innate pathway, inducing antiapoptotic genes (41). Together, these data indicate that, under these conditions, the cells are viable and are actively replicating and secreting RSV.

We next sought to quantify the reproducibility of our label-free proteomics workflow to detect changes in secreted proteins by cell type and in response to RSV infection. Cell culture supernatants from four independent biological replicates from tert-hBECs and tert-hSAECs were analyzed in mock-infected cells and at 24 h after RSV infection (Fig. 2A). A total of 1559 proteins was identified in the supernatants with an FDR < 1%, as determined by target-decoy database searching. To determine biological reproducibility, pairwise analysis of the log2-transformed protein abundance was performed. Pearson correlations (r²) were >0.85, indicating a high degree of concordance (Supplemental Fig. 1, Table I). We noted that, across samples, mean r² was greater for RSV-induced CM versus control (for tert-hBECs: +0.957 ± 0.005 versus +0.863 ± 0.008, respectively, for tert-hSAECs: +0.936 ± 0.011 versus +0.863 ± 0.01, respectively; p < 0.05, Student t test for both). These data indicate that the method was reproducible and that the highly abundant proteins in the RSV-induced CM were more accurately measured.

To further extend our findings that the proteins in the secretome are not derived from cellular lysis, cellular lysates were prepared from the same experiment and analyzed in parallel (Fig. 2A). A total of 1929 unique proteins was quantified from the tert-hBEC and tert-hSAEC WCLs. The global protein expression patterns in the CM and WCLs were examined first using PCA. In this analysis, 91.6% of the variability was accounted for in the first two dimensions, indicating a robust analysis (Fig. 2B). The WCLs from control or RSV-infected cells clustered by cell type and presence of RSV infection, indicating that biological replicates were consistent. We noted that the control tert-hBEC WCLs formed a distinct cluster from that of the tert-hSAEC WCLs, separated by the second principal component dimension (Fig. 2B). The RSV-infected tert-hBEC WCLs moved in the second dimension to form a cluster that overlapped with control and RSV-infected tert-hSAEC WCLs. The control secretomes of tert-hBECs and tert-hSAECs clustered together, widely separated in the first dimension from the WCL clusters. Upon RSV infection, the tert-hBEC and tert-hSAEC secretomes migrated up in the second dimension and down in the first dimension (Fig. 2B). Together, these analyses indicate that the secretome represented a distinct protein set from the cellular lysate and that RSV induced significant changes in its composition.

A Venn diagram comparison of WCLs versus the secretome is shown in Fig. 2C; 460 proteins were present only in the secretome, 885 proteins were unique to the cellular proteome, and 1044 proteins were present in both datasets. To further confirm that these protein sets were distinct, we conducted unbiased gene ontology cellular component (GOCC) enrichment analysis of the proteins present only in the CM dataset. This analysis indicated that the secretome was enriched with proteins derived from the extracellular region (205 of 460 proteins), whereas the cytoplasmic and mitochondrial proteins were depleted (Fisher exact test, Benjamini–Hochberg FDR 0.1%; data not shown). In contrast, proteins unique to the WCL were enriched with regard to mitochondrial, ribosomal, and nuclear proteins, and proteins in the extracellular region were depleted.

To confirm that the secreted proteins were independent of cellular lysis in a more quantitative manner, we used the NSAF method to confirm enrichment of the proteins in CM and WCLs. NSAF is a widely used spectral counting method for label-free proteomic quantitation (42–44). In spectral counting, larger proteins usually generate more peptides and, thus, more spectral counts than smaller proteins. Therefore, the number of spectral counts for each protein is first divided by the protein length, which defines the spectral abundance factor (SAF). Furthermore, to accurately account for sample-to-sample variation, individual SAF values are normalized by dividing by the sum of all SAFs for proteins identified in the sample, resulting in the NSAF value (32). In this manner, NSAF values are standardized across distinct samples, allowing direct comparisons of the relative protein abundance across samples.

We then conducted a pairwise comparison of NSAF values for proteins in the RSV CM and the WCLs in RSV-infected tert-hBECs (Fig. 2D). A two-sample t test was used to assess the statistical significance between protein enrichment in the RSV CM and the WCLs. For tert-hBECs, proteins enriched in the CM (indicated in red, Benjamini-Hochberg FDR 1%) included macrophage inhibitory factor-1, macrophage migration CXCL-10, IFN-stimulated gene-15, high mobility group box 1/2, IFN-induced protein with tetratricopeptide repeats-3, IFN-λ2, and others. All of these proteins are well-characterized, secreted proteins with defined roles in innate immunity. The abundance of these proteins was depleted in WCLs. Conversely, we identified high-abundance intracellular proteins in the cell extract (indicated in blue, Benjamini–Hochberg FDR 1%) that were depleted in the CM, including histones H2HAC/1H2BM, mitochondrial ssDNA-binding protein-1, heat shock 10 kDa protein 1, and actin γ-1 (Fig. 2D).

Similar observations were made with regard to the comparison of high-abundance proteins in tert-hSAEC CM and WCLs (Fig. 2E). Although the proteins making up the tert-hBEC and tert-hSAEC CM were similar, we noted that CCL5 was in high abundance in tert-hBEC CM and was much lower in tert-hSAEC CM. Conversely, CCL20 and IL-6 were much more abundant in the tert-hSAEC secretome than in the tert-hBEC secretome (Fig. 2D, 2E). Together, these data indicate that CM samples represent a distinct proteome profile versus that in the WCL. We will refer to this population of proteins as the “secretome” in the remainder of this article.

To further support the conclusion that the secretome and cell lysates represent distinct protein pools, we conducted unbiased GOCC enrichment. The top-ranked cellular components for the RSV-induced secretome of tert-hBECs (indicated in red) were extracellular matrix, extracellular space, and extracellular organelle, indicating that this sample was enriched in extracellular proteins relative to the reference human proteome (Fig. 2F). The cellular components corresponding to cell part, macromolecular complexes, and ribonucleoprotein complexes were depleted in the secretome (indicated in blue, Fig. 2F). In contrast, in tert-hBEC WCLs, the GOCCs nucleolar ribonucleoprotein complex, NADH dehydrogenase complex, and ribosomal complex were the top-ranked components (red bars, Fig. 2F). Similarly, the proteinaceous extracellular matrix and extracellular matrix were the two most significantly depleted cellular components (blue bars, Fig. 2F). These data further support that the proteins identified in the secretome represent a distinct population from the intracellular proteome in WCLs. There were similar findings for tert-hSAECs, with extracellular proteins being the most highly enriched proteins in the hSAEC secretome GOCC analysis (Fig. 2G). We noted that the cellular component terms for the hBEC and hSAEC secretomes were almost identical (Fig. 2F, 2G).

We noted that the majority of the 1044 proteins that were present in both the tert-hBEC and tert-hSAEC secretomes were cytosolic proteins. These cytosolic proteins may be secreted via unconventional protein-secretory pathways, perhaps mediated by Golgi or endosomal export mechanisms (45). To this point, we found that 65 of these common proteins, including heat shock cognate 71 kDa protein (HSPA8, GAPDH) and annexin-A2 (34) are prominent exosomal proteins. To provide some insight into whether endosomal transport was contributing to the RSV-induced secretome, we isolated and quantified exosomal proteins from control and RSV-infected tert-hBECs and tert-hSAECs.

Ultracentrifuge-purified exosomes were 112.8 ± 2.0 nm in size by dynamic light scattering and exhibited a characteristic membrane composition under transmission electron microscopy (Fig. 3A). From this fraction, 937 exosomal proteins were identified; of these, 564 proteins were identified in the secretome (Fig. 3B; the 373 proteins unique to the exosomal fraction were below the limit of detection and were not observed in the secretome analysis). The 937 proteins were subjected to GOCC. Significantly enriched cellular components included lysosomal, vacuolar, and endoplasmic reticular (Fig. 3C). Of the 937 identified exosome proteins, 853 were quantified across the experimental groups. Pairwise comparison by cell type and presence of RSV infection was accomplished using the Student t test, as shown in a volcano plot, where the log2 fold change is plotted versus the log10-transformed p value (Fig. 3D). RSV infection caused upregulation of 220 tert-hSAEC and 241 tert-hBEC exosomal proteins and downregulation of 146 tert-hSAEC and 223 tert-hBEC exosomal proteins (FDR < 0.05). The protein contents in the exosome also display cell type differences. In the basal state, 31 proteins were more abundant in the tert-hSAEC exosome fraction, and 60 proteins were more abundant in the tert-hBEC exosome fraction (FDR < 0.05). After RSV infection, 273 exosome proteins were different by cell type, with 134 proteins more abundant in RSV-infected tert-hSAEC exosomes and 139 proteins more abundant in RSV-infected tert-hBEC exosomes (FDR < 0.05).

We applied a statistical filter using a p value (Student t test with Benjamini–Hochberg FDR correction < 0.05) and an expression filter of ±5-fold change between control and RSV-infected NSAF to identify the most highly differentially expressed exosomal proteins. Next, the expression patterns were compared by hierarchical clustering (Fig. 3E). These data clearly indicate that the exosomal proteins are different by cell type and are modified by the presence of RSV infection.

Of the 559 proteins in the secretome not directly found in the exosomal fraction, 80 proteins have identifiable signal peptides, with the remainder being enriched in lysosomal and vacuolar proteins. These findings suggest that the RSV-induced epithelial secretome is mediated primarily by exosomal protein release, with a smaller fraction due to lysosome- or vacuole-mediated export and a small fraction by classical protein secretion.

We next examined the differential expression patterns of secretomes by cell type. SAM was applied to identify differentially expressed proteins using 1% FDR. SAM analysis identified 71 proteins that were distinct in the control secretomes from tert-hBECs versus tert-hSAECs (Fig. 4A). Of these, 61 were upregulated in the tert-hSAEC secretome. Similarly, 131 proteins showed differential expression in RSV-induced secretomes from tert-hBECs and tert-hSAECs (Fig. 4B). Of these, 65 were upregulated in hSAECs.

The differentially expressed proteins were subjected to two-dimensional hierarchical clustering. Hierarchical clustering groups proteins (rows) whose expression patterns are most similar across cell type and treatment condition. The samples (columns) are also clustered by the patterns of proteins. In this analysis, the treatment groups coclustered in the vertical dimension, consistent with the findings in PCA that the replicates from each cell type are highly similar (Fig. 4C). By inspection of the proteins in the rows, five distinct patterns of protein expression emerged. Cluster 1 represents proteins abundant in control tert-hBECs whose expression is inhibited in response to RSV (not expressed by tert-hSAECs). Cluster 2 represents proteins expressed by tert-hSAECs whose abundance is decreased in response to RSV. Cluster 3 contains proteins only expressed by RSV-infected tert-hSAECs. Cluster 4 contains proteins induced by RSV that are common to both cell types. Cluster 5 contains proteins induced by tert-hBECs but not tert-hSAECs. We note that Cluster 3 contains a number of immunologically significant proteins, including CXCL1, IL-6, and CCL20. These data suggest that RSV induces cell type differences in the secretion of immunologically significant cytokines. Collectively, our analysis pipeline developed using tert-immortalized human epithelial cells enables the reliable analysis of epithelial secretomes to understand differences by cell type and RSV-induced expression patterns.

We next applied our analysis pipeline to phBECs and phSAECs. To control for donor effects, the analysis was conducted on three independent donors (donor demographic data are shown in Supplemental Table I); two biological replicates were analyzed for each. As validation of the distinct phenotypes, IF microscopy was used to examine differences in epithelial cytokeratin expression. phBECs express cytokeratins 7 and 19; in contrast, hSAECs have low (or undetectable) cytokeratin 7 expression and strong cytokeratin 19 expression (Fig. 5A). Both cell types support active RSV replication and secretion of virus (data not shown).

Secretome fractions were prepared from control and RSV-infected primary cells. We identified 2376 proteins with FDR 1%. SAM analysis identified 577 proteins in the secretome from control (uninfected) cells whose expression varied by cell type and 966 proteins in the secretome from RSV-infected cells whose expression varied by cell type (FDR of 1%). To focus on the most robust differentially expressed proteins, we filtered proteins that showed a ≥5-fold expression change; this resulted in 492 of the most highly significant and induced proteins. This filtered dataset was subjected to two-dimensional hierarchical clustering, where each column represents a sample, and each row represents an individual protein’s abundance. We observed that each sample coclustered with its replicate, as well as by being grouped by cell type and RSV treatment (Fig. 5B). The row-wise clustering of proteins produced a pattern highly similar to that observed in the Tert-immortalized hBECs and hSAECs (Figs. 4C, 5B); in this example, Cluster 2 represents 11 proteins upregulated by RSV infection in phBECs, Cluster 3 represents 116 proteins upregulated by RSV infection in phSAECs, and Cluster 4 contains 203 proteins induced by both cell types (Table II).

To validate the differentially expressed protein patterns, we independently quantified their expression using SID-SRM-MS. This method is a “targeted” MS approach for the detection and accurate quantification of proteins in a complex background where signature proteotypic peptides unique to the protein of interest are monitored in a high mass accuracy mass spectrometer (35, 46–48). Because the peptide fragments are then subjected to fragmentation, the assay provides structural specificity and, therefore, is the most accurate approach available for direct quantification of target proteins in a complex mixture (48). SRM assays were developed for the measurement of 15 proteins. These assays confirmed the constitutive cell type–specific expression of fibronectin, as well as the RSV-induced expression of guanine nucleotide-binding protein, annexin-X2, ras oncogene-7A, aldolase-A, heat shock protein-90, vimentin, IL-6, integrin-α3, caveolin-1, and IFN-induced protein with tetratricopeptide repeats-1/2/3 by phSAECs (Fig. 6). These data indicated that RSV induces differential expression of proteins by cell type.

To gain further understanding of RSV-inducible proteins, we analyzed the clusters by GO biological function and gene set enrichment analysis (GSEA), primarily focusing on Clusters 2, 3, and 4 (Fig. 5B). The 11 proteins in Cluster 2 were too limited to identify extensive enriched GO categories. Relative to the human proteome, Cluster 2 was depleted in the GO category for metabolic process (Fig. 7A). GSEA showed enrichment for the extrinsic prothrombin pathway and fibrinolysis pathway, predominantly determined by the presence of fibrinogen (Fig. 7A). The 116 proteins in Cluster 3, uniquely expressed by RSV-infected hSAECs, showed enrichment for metabolic processes (organic acid, cellular ketone, and small molecule) relative to the human proteome, as well as depletion in RNA processing (Fig. 7B). GSEA identified enrichment of canonical pathways for glycoxylate, carbonyl, porphyrin, and amino acid metabolism, as well as oxidative reduction (Fig. 7B). This analysis suggests that phSAECs secrete proteins controlling nucleotide–sugar bioenergetic processes in response to RSV.

The 203 proteins secreted by both epithelial cell types (Cluster 4, Fig. 5B) were analyzed in the same manner. This analysis identified the most GO functions numerically, many of which could be collapsed into mRNA processing (splicing, catabolism, RNase complex assembly), DNA cell cycle regulation, and others (Fig. 7C). GSEA indicated enrichment in mRNA destabilization, Wnt signaling, HIV factor interactions, and mRNA interaction/metabolism (Fig. 7C).

To obtain further insights into the differentially regulated proteins, we subjected all proteins showing differential expression to Ingenuity Pathways Analysis upstream regulator analysis (49). Upstream regulator analysis compares the known effect (transcriptional activation or repression) of a transcriptional regulator on its target genes with the observed changes in protein abundance. In phBECs, the epithelium-specific ets homologous factor was predicted to be more upregulated and responsible for regulating MUC1, serum amyloid A2, and kallikrein-related peptidase-6/7 (Fig. 8A). In phSAECs, the NF-κB transcription factor was predicted to be activated to a greater degree in response to RSV infection than in hBECs. The NF-κB network is responsible for regulating TSLP, CCL20, BMP2, MMP3, and SOD2 (Fig. 8B).

Our protein-level analysis of the proteins unique to hSAECs (Cluster 3, Fig. 5B) identified three proteins highly relevant to the pathogenesis of RSV LRTI: TSLP, CCL20, and CCL3-L1. These proteins contain signal peptide sequences and are found in the free (nonexosomal) fraction. TSLP is produced by RSV-infected airway epithelium and promotes Th2 differentiation by inducing the maturation of Ag-presenting DCs (50, 25). CCL20 is a potent inducer of epithelial mucin production (51), Th17 lymphocyte expansion and DC chemotaxis. These activities promote formation of mucosal lymphoid tissue playing an important role in the pathology of RSV-induced lung inflammation (52). CCL3-L1 is chemotactic for monocytes and lymphocytes and interacts with CCR5, a receptor linked to RSV LRTIs (53).

Therefore, we selected these three immunologically important proteins that were previously implicated in the pathogenesis of RSV LRTIs for validation experiments. To independently measure their differential expression by RSV infection and cell type, we developed and applied highly specific SID-SRM-MS assays to measure target protein abundance in the phBEC and phSAEC secretomes in the presence or absence of RSV infection. We were able to detect a 10-fold increase in CCL20, TSLP, and CCL3-L1 expression in RSV-infected phSAECs relative to uninfected controls (Fig. 9A). In contrast, RSV-infected phBECs showed only a 2-fold induction of each chemokine and at much lower amounts than that in phSAECs (Fig. 9A). As shown in Fig. 9B, the presence or absence of BSA/growth factor supplements in the cell culture medium did not affect the RSV-induced cell type differences in the secretion of CCL20, TSLP, and IL-6.

Although mucins constitute an important arm of the innate immune response via their ability to trap microorganisms (54), they also play an important role in the pathogenesis of airway obstruction in RSV LRTIs. As described earlier, mucous plugging of the small airways is an important mechanism for atelectasis in bronchiolitis, producing ventilation-perfusion mismatching and hypoxia (55, 56). To determine whether the RSV-induced CCL20 production was at levels sufficient for biological activity, we first evaluated if phBECs express the CCR6 receptor mRNA using quantitative real time PCR (Q-RT-PCR). Both cell types express CCR6 in uninfected and infected conditions (Fig. 9C). We next stimulated naive phBECs with rCCL20 or CM from RSV-infected phSAECs and assayed for MUC5A mRNA expression by Q-RT-PCR. rCCL20 induced a reproducible 8-fold induction of MUC5AC mRNA expression relative to control, which was lost at higher concentrations (Fig. 9D). RSV CM produced a similar 8-fold induction of MUC5A (Fig. 9E). Both of these activities were inhibited by the addition of neutralizing CCL20 Ab; preimmune IgG had no effect (Fig. 9F, 9G). Together, these data indicate that RSV-infected phSAECs produce biologically active CCL20 that stimulates mucin production.

Our quantitative in vitro proteomic studies suggest that lower airway epithelial cells exhibit enhanced CCL20 secretion upon RSV infection. To confirm this, we examined CCL20 expression in a BALB/c mouse model of acute RSV infection established in our laboratories (57). IF assays were performed on proximal and distal airways in control and RSV-infected mice. In the absence of primary Ab, no fluorescence was observed (Fig. 10A). In proximal airways, IF was faintly distributed within the epithelial layer and was weakly induced upon RSV infection (data not shown). In contrast, CCL20 IF was strongly induced in the smaller airways, upon RSV infection (Fig. 10A). These (<1-mm diameter) airways were lined with a single layer of cuboidal epithelium and lacked cartilage representing bronchiolar and terminal bronchiolar structures (adjacent to alveoli).

To examine whether enhanced mucin production was seen in this model, tissue sections were stained with PAS to assess mucin production. We observed enhanced PAS staining in these smaller, distal airways (Fig. 10B). Together, these studies confirm that lower airway epithelial cells in the distal airways produce enhanced CCL20 expression, with increased mucin production in this mouse model of RSV infection.

RSV is an important human pathogen representing the most common cause of childhood LRTIs (7) and is the leading cause of infant viral death (9). RSV directly replicating in the airway epithelium triggers pulmonary innate immune responses. RSV replication produces the viral molecular patterns 5′-phosphorylated RNA and dsRNA, which are recognized by the cytoplasmic RIG-I cytoplasmic pattern recognition receptor that induces NF-κB and IRF3 translocation and activation; these inducible transcription factors coordinate the expression of inflammatory chemokines via extensive cross-talk pathways (58). Consequently, infected epithelial cells rapidly secrete type I and type III IFNs (24, 37, 59), and 17 C-, CXC-, and CC-type chemokines (15, 60, 61). How RSV infection of the lower airway contributes to the observed pathophysiology is largely unknown. Previous limited work comparing ciliated and transformed type II–like alveolar cells found that these different cell types exhibit qualitatively distinct patterns of RSV-induced CC chemokine production (20). In this study, we apply unbiased proteomics to identify 577 high-confidence proteins whose RSV-induced expression patterns differ between primary human epithelial cells derived from the conductive airway (trachea) and those of the small airways (bronchioles). A surprising finding was that about one third of the proteins identified in the secretome are exosomal. Although a number of RSV-inducible proteins are common, RSV induces a group of proteins unique to phSAECs that is immunologically significant: TSLP, CCL20, CCL3-L1, and IL-6. Differential expression of these proteins was independently validated by specific SID-SRM-MS assays. We demonstrate that CCL20 is the major mucin-promoting cytokine in the hSAEC secretome and validate its preferential expression in the lower airways in a BALB/c mouse model of RSV infection. These data advance our understanding of the epithelial innate response and provide insight into how RSV LRTI is associated with enhanced mucin production and lower airway obstruction.

The epithelial surface of the pulmonary mucosa consists of highly differentiated, regionally distinct epithelial cell types, each playing a specialized role in normal pulmonary function and host defense. Upper airway ciliated epithelial cells produce protective epithelial lining fluid and regulate water and ion transport, whereas pseudostratified tracheal airway cells produce mucins and coordinated ciliary beating to facilitate mucociliary particulate clearance (62). Despite a significant amount of cellular proteomic profiling of RSV–epithelial interactions (63–65), a systematic understanding of the spectrum of secreted proteins as a function of epithelial phenotype has not been obtained. In this study, we selected a model of primary human airway epithelial cells cultured in submerged monolayers. This cellular model has been used extensively for the study of innate responses to RSV and provided important insights into the mechanisms of activation of the innate response and its subversion by RSV nonstructural proteins. Although growth under these conditions confers basal cell–like characteristics, recent studies showed that basal cell infection is important for amplifying RSV infection in ciliated cells (66). Our validation of differential cytokine expression in the mouse model provides further relevance of our findings to pulmonary host defense. An important extension of our work will be to conduct these studies with polarized cells in air–liquid interface.

Recent advances in secretome profiling provided interesting insights into inducible protein secretion. Secretome profiling of macrophages using a proteomic pipeline similar to ours showed that TLR4 activation induced the production of >775 proteins present in concentrations > 1 pg/ml (67). Like our study, the vast majority of secreted proteins that were identified included lysosomal, cytoplasmic, and nuclear proteins, proteins that typically lack signal peptides. To minimize the possibility that these unconventionally secreted proteins represented cellular necrosis or apoptosis, we used a lower MOI and sampled early time points, before significant apoptosis was produced in our experimental design. In fact, direct measurements of the apoptotic rate showed less apoptosis in RSV-infected cells than in control cells as a result of the antiapoptotic effects of NF-κB demonstrated by earlier studies (40). Moreover, direct comparison of secreted versus WCL proteomic profiles, functional classification by GO, and unbiased PCA clustering all consistently indicate that the CM and WCLs are drawn from distinct protein pools. Instead, we demonstrate in this study that a major fraction of secreted proteins from epithelial cells is contained within membrane-bound exosomes.

Exosomes are membrane-bound nanoparticles derived from intracellular multivesicular bodies that contain proteins and microRNAs important in intercellular communication (68). Studies on exosomes isolated from bronchoalveolar lavage samples of atopic asthmatics showed that the exosomes may be involved in the regulation of bronchial hyperresponsiveness and inflammation (69, 70). Other studies showed that exosomes isolated from highly differentiated ciliated tracheal epithelial cells mediate a protective antiviral response via the expression of sialoproteins that blocks influenza infection (71). Others found that IL-13, a mediator of allergy and Th2 lymphocyte recruitment in asthma, induces exosome production in the airway that promotes the chemotaxis of macrophages (72). The role of exosomes in anti-RSV innate immunity and inflammation will require further exploration.

Additional proteins identified in the RSV secretome appear to be derived from unconventional protein-secretory pathways. Our identification of lysosomal, vacuolar, and nuclear proteins expressed in response to RSV replication suggests that unconventional protein secretion, perhaps through Golgi compartments, is also a component of the epithelial innate response.

Bioinformatic comparisons of the RSV-infected hBEC secretome and the hSAEC secretome show that both cell types inducibly express >200 common proteins. GO analysis indicates that these proteins play diverse functional roles. Although our focus is on proteins of potential immunological importance to LRTIs, these data may help to identify other potential mechanisms of pulmonary innate defense. We note, for example, the presence of multiple types of RNA-binding proteins of distinct functional classes; these may serve to protect the airway by binding or metabolizing infectious nucleic acids. Similarly, the identification of Wnt signaling, an important epithelial morphogenic pathway (73), may provide clues to the mechanisms of airway remodeling induced by RSV infection.

RSV LRTIs were associated with Th2 polarization and enhanced aeroallergen sensitivity (74, 75). The epithelium plays an important role in shaping the T cell lymphocytic response through the patterns of chemokines and cytokines produced during infection. TSLP is an RSV-inducible, epithelium-derived chemokine that is important in allergic and RSV-induced airway inflammation (76, 77) and is secreted by airway cells in an NF-κB–dependent mechanism (49), consistent with our upstream factor analysis. Although TSLP has a broad variety of target cell responses, its ability to activate pDC populations and promote Th2 lymphocyte–predominant inflammation is especially relevant to the pathophysiology of severe LRTI infection. Similarly, CCL20 is important in pDC activation and the recruitment of Th17 lymphocytes to sites of inflammation (78). The actions of Th17 cells also were implicated in the pathogenesis of RSV LRTIs (79); clinical studies of children with severe RSV infection showed increased IL-17 levels in their airway fluids. Interestingly, IL-17 appeared to be proinflammatory and promucinogenic yet, paradoxically, was protective against hypoxia (79). Our study surprisingly shows the novel observation that TSLP and CCL20 are preferentially secreted by RSV-infected lower airway epithelial cells, perhaps providing information on how LRTI is associated with Th2 lymphocyte skewing, DC recruitment, and Th17 activation.

Severe RSV infections produce a characteristic pathological pattern that includes epithelial necrosis, sloughing, and mucous plugging of the small bronchioles, resulting clinically in hypoxia and hyperaeration (12, 55, 56). Mucus plugging is a complex phenomenon determined by viral and host factors. Although the RSV nonstructural protein NS2 was demonstrated to induce shedding of bronchiolar epithelial cells (12), the mechanisms for enhanced mucin production in RSV infection are not well understood. Rodent models of RSV disease did not consistently shown high levels of mucin production, unless the animals had prior allergic sensitization (80) or strains of RSV were used that selectively induce IL-13 production (81).

In this study, we observe that RSV-induced CCL20 production is restricted to the epithelium and that it is preferentially secreted by lower airway cells. Other elegant studies showed that CCL20 plays a pathogenic role in the mouse model of infection. Knockouts of the CCL20 receptor (CCR6) or CCL20 neutralization showed that inhibition of the CCL20–CCR6 axis decreases RSV-induced MUC5A and Gob5 expression and Th2 responses (52). Our study has demonstrated that CCL20 is the major biological protein inducing mucin production secreted from RSV-infected hSAECs. Interestingly, the molecular pathway producing MUC5A expression is mediated by cleavage of the EGFR ligand by the cell surface ADAM17/TACE distintegrin (51). This previous study was unable to induce MUC production in primary airway epithelial cells; in this study, we demonstrate that rCCL20 and CCL20 in hSAEC CM induces MUC5A expression in phBECs and that phBECs express CCR6R. These different results may be due to cell-type differences in the expression of the ADAM17/TACE distintegrin or variability in CCR6 expression in cultured cells.

In summary, we apply quantitative, label-free MS profiling to identify cell type-specific patterns in RSV-induced protein secretion that arises from the exosomal compartment. We validate the differential secretion patterns across multiple donors and confirm our findings in a mouse model in vivo. We demonstrate that lower airway secretion of CCL20 is a mucin-promoting factor that may account for enhanced mucous plugging in distal airways in RSV LRTIs. These data provide insights into the pathogenesis of Th2 polarization and mucous plugging in this disease.

We thank Dr. David Konkel for critically editing the manuscript. Core laboratory support was provided by the Sealy Center for Molecular Medicine Selected Reaction Monitoring Facility and the University of Texas Medical Branch Optical imaging and Histochemistry facilities.