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. 2022 Apr 1;8(13):eabm9718. doi: 10.1126/sciadv.abm9718

Mucus concentration–dependent biophysical abnormalities unify submucosal gland and superficial airway dysfunction in cystic fibrosis

Takafumi Kato 1,2, Giorgia Radicioni 1, Micah J Papanikolas 3, Georgi V Stoychev 1, Matthew R Markovetz 1, Kazuhiro Aoki 4, Melody Porterfield 4, Kenichi Okuda 1, Selene M Barbosa Cardenas 1, Rodney C Gilmore 1, Cameron B Morrison 1, Camille Ehre 1,5, Kimberlie A Burns 1, Kristen K White 6, Tara A Brennan 1, Henry P Goodell 1, Holly Thacker 1, Henry T Loznev 1, Lawrence J Forsberg 7, Takahide Nagase 2, Michael Rubinstein 8, Scott H Randell 1, Michael Tiemeyer 4, David B Hill 1,9, Mehmet Kesimer 1, Wanda K O’Neal 1, Stephen T Ballard 10, Ronit Freeman 3, Brian Button 1,*, Richard C Boucher 1,*
PMCID: PMC10938572  PMID: 35363522

Abstract

Cystic fibrosis (CF) is characterized by abnormal transepithelial ion transport. However, a description of CF lung disease pathophysiology unifying superficial epithelial and submucosal gland (SMG) dysfunctions has remained elusive. We hypothesized that biophysical abnormalities associated with CF mucus hyperconcentration provide a unifying mechanism. Studies of the anion secretion–inhibited pig airway model of CF revealed elevated SMG mucus concentrations, osmotic pressures, and SMG mucus accumulation. Human airway studies revealed hyperconcentrated CF SMG mucus with raised osmotic pressures and cohesive forces predicted to limit SMG mucus secretion/release. Using proline-rich protein 4 (PRR4) as a biomarker of SMG secretion, CF sputum proteomics analyses revealed markedly lower PRR4 levels compared to healthy and bronchiectasis controls, consistent with a failure of CF SMGs to secrete mucus onto airway surfaces. Raised mucus osmotic/cohesive forces, reflecting mucus hyperconcentration, provide a unifying mechanism that describes disease-initiating mucus accumulation on airway surfaces and in SMGs of the CF lung.

Hyperconcentrated mucus unifies submucosal gland and superficial airway dysfunction in cystic fibrosis.

INTRODUCTION

Cystic fibrosis (CF) is characterized by abnormal transepithelial ion transport reflecting mutations in the CF transmembrane regulator (CFTR) gene (1–3). In vitro and in vivo studies of the superficial epithelia lining CF airway surfaces suggest that dysfunctional CFTR-mediated ion transport produces hyperconcentrated mucus (4–8). Mucus hyperconcentration, defined as an increased mucus percent solid content (% solids) and/or mucin concentration, generates abnormal mucus biophysical properties, e.g., raised osmotic pressures and increased cohesive forces (7, 9). Osmotic compression of the periciliary layer (PCL) by the mucus layer in CF abolishes mucociliary transport, produces mucus accumulation on distal airway surfaces, and initiates the obstruction, infection, and inflammation that ultimately produce bronchiectasis (4, 8–11).

However, there is controversy with respect to the contribution of CF submucosal gland (SMG) dysfunction to the proximal airway features of CF lung disease. Studies in CF pig tracheas described secretion of SMG mucus strands in response to maximal cholinergic stimulation (a mimic of cough) that are postulated to adhere to proximal CF airway surfaces, accumulate, and become a nidus for persistent bacterial infection (12–17). These data are juxtaposed to other studies that suggest that CF SMG dysfunction reflects a failure to secrete SMG mucus with its robust antimicrobial components onto airway surfaces (18–22). Biophysical or clinical measurements that distinguish between these hypotheses have not been reported.

We hypothesized that normal SMG mucus has unique adaptations for host defense, but, in common with superficial airway mucus, CF SMG mucus is hyperconcentrated, which limits secretion onto airway surfaces. Established methods were used for SMG mucus collection from freshly excised porcine and human airway tissues, and multiple biochemical and biophysical properties of SMG mucus relevant to mucus transport were measured. A biomarker selective for SMG secretion was identified and used as an index of SMG secretion in vivo in CF versus normal and disease control subjects.

RESULTS

SMG mucus from CF-mimicking pig airways exhibits raised concentrations and osmotic pressures

We first studied the osmotic pressures generated by normal versus CF-like SMG mucus from the anion transport inhibitor–treated wild-type (WT) pig (23–26). Because polymer biophysical properties are higher-order powers of polymer concentration, mucus concentration was measured in parallel (5, 27). Maximal acetylcholine (Ach) stimulation, block of Cl− secretion [bumetanide (Bum)], block of bicarbonate secretion [dimethylamiloride (DMA)], combined Bum/DMA block, or vehicle in WT pig SMG mucus reproduced evidence of increased SMG mucus concentrations due to anion-transport inhibitor block of fluid secretion (% solids) (Fig. 1A). Osmotic pressures were raised under conditions of fluid secretion block, paralleling % solid measurements (Fig. 1, B and C). Note that the anion transporter inhibitor protocols have opposite effects on SMG mucus HCO3− concentrations, i.e., Bum raises HCO3− concentrations, whereas Bum/DMA lowers HCO3− concentrations, suggesting the increase in osmotic pressures generated by both inhibitor protocols reflected effects on mucus concentration and not HCO3−/pH (28, 29). The osmotic pressure values measured under fluid block conditions are predicted to compress the PCL and slow/stop mucus transport (5).

Fig. 1. Studies of anion transport–inhibited porcine airways.

Fig. 1.

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(A) Solid concentration (% solids) of porcine SMG mucus after treatment with Ach without or with DMA, Bum, or the combination of DMA and Bum. n = 6 to 8 per group. Welch’s analysis of variance (ANOVA) test followed by Dunnett’s multiple comparison test. (B) Osmotic pressure (in pascals) of porcine SMG mucus after treatment with Ach without or with DMA, Bum, or DMA/Bum. n = 6 to 8 per group. Welch’s ANOVA test followed by Dunnett’s multiple comparison test. (C) Linear correlation between % solids and osmotic pressure of pig SMG mucus. n = 29. (D) Transmission electron microscopy (TEM) images of porcine SMG ducts after Ach treatment without or with DMA and Bum. (E) Quantitation of SMG (i) mucus retention and (ii) cilial height in ducts for each treatment condition. (i) n = 4 to 10 ducts from three airway specimens for each condition from two pigs. One to eight regions of mucus retention per duct were evaluated. Each score was used for plot and statistical analyses. n = 72 regions from 25 ducts in total. Kruskal-Wallis test followed by Dunn’s multiple comparison test. (ii) n = 4 to 10 ducts from three airway specimens for each condition from two pigs. One to eight regions of cilial height per duct were measured. Mean of cilial height for each duct was used for plot and statistical analyses. Welch’s ANOVA test followed by Dunnett’s multiple comparison test.

CF-mimicking maneuvers produce mucus retention and compressed cilia in SMG ducts

The presence of mucus and cilial height in SMG ducts of the control (Ach) and CF-mimicking (Ach + Bum + DMA) maneuvers were investigated using transmission electron microscopy (TEM). SMG mucus retention was observed under Bum/DMA (low gland fluid flow/anion secretion inhibition) conditions (Fig. 1, D and E). Cilia in glands treated with Bum/DMA were also compressed, suggesting that ductal mucus retention reflected osmotic compression of cilia (Fig. 1, D and E) (5). Collectively, these data suggest that anion/fluid secretion inhibitors produced a hyperconcentrated pig SMG mucus that, via the osmotic compression of the ductal surface PCL, was retained in SMGs.

Human CF SMG mucus exhibits increased concentrations, osmotic pressures, and cohesive strength

Consistent with the absence of CFTR anion/fluid secretory function in CF, CF SMG mucus was hyperconcentrated with increased osmotic pressures compared to non-CF SMG mucus (Fig. 2, A and B). Mucus concentrations correlated with osmotic pressure over ranges bracketed by CF and non-CF SMG mucus (Fig. 2C). Studies designed to measure the forces required to “tear” mucus from gland orifices for transport up airway surfaces, i.e., cohesive forces (30, 31), identified elevated cohesive forces in CF compared to non-CF SMG mucus (Fig. 2D).

Fig. 2. Studies of non-CF and CF human SMGs.

Fig. 2.

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(A) Solid concentration (% solids) of human SMG mucus from non-CF (n = 35) and CF (n = 11) subjects. Welch’s t test. (B) Osmotic pressure (in pascals) of human SMG mucus from non-CF (n = 11) and CF (n = 8) subjects. Welch’s t test. (C) Linear correlation between % solids and osmotic pressure of human SMG mucus. (D) Cohesive strength (in millinewtons per centimeter) of human SMG mucus from non-CF (n = 8) and CF (n = 5) subjects. Welch’s t test. (E) SMG ducts of healthy control versus CF airways after Ach treatment captured by TEM. Scale bars, 2 μm. (F) Quantitation of human SMG (i) mucus ductal retention and (ii) cilial height in healthy control versus CF groups. (i) n = 37 regions from 15 healthy control ducts and n = 40 regions from 12 CF ducts. One to eight regions of mucus retention per duct were evaluated depending on TEM image availability. Each score was used for plot and statistical analyses. Wilcoxon’s signed-rank test. (ii) n = 15 ducts from three healthy control donors and n = 12 ducts from three CF donors. One to seven regions of cilial height per duct were measured. Mean of cilial height for each duct was used for plot and statistical analyses. Welch’s t test.

CF SMGs exhibit mucus retention in ducts and compressed cilia

Consistent with the findings in CF-mimicking pigs (Fig. 1, D and E), human CF SMG ducts exhibited TEM mucus retention and cilial compression (Fig. 2, E and F). These findings are consistent with previous reports of increased CF SMG mucus viscosity (29), and they predict that CF SMG mucus secretion is hindered by both osmotic compression of SMG mucus onto ductal surfaces and increased cohesive forces that limit mucus separation from ductal orifices.

Mucus strand formation as a unique feature of SMG mucus

We initiated studies of the properties of SMG mucus by comparing its composition with superficial airway epithelial mucus. Because SMG mucus in naïve porcine models has been reported to contain “strands” that maintain their structure after secretion onto airway surfaces, rather than dissolve into the ambient surface mucus layer (12–15), we investigated whether strand formation was unique to SMG mucus, i.e., not a feature of superficial epithelial mucus, and whether CF SMGs produced abnormal strands. Superficial human bronchial epithelial (HBE) mucus dissolved fully into phosphate-buffered saline (PBS), leaving behind no strand-like structures. In contrast, ~50% of Ach-stimulated non-CF human and pig SMG mucus failed to dissolve (Fig. 3A). Addition of a nonionic, nondenaturing detergent (IGEPAL) promoted dissolution of the insoluble SMG mucus component (table S1). Light scattering of the PBS-soluble SMG mucins demonstrated that SMG mucins were of similar size (Rg) as HBE mucins (Fig. 3B). Light scattering of the insoluble SMG mucins was not possible. Instead, we immunostained the insoluble components of SMG mucus and identified SMG strands in mucus as composed of MUC5B but not MUC5AC mucin (Fig. 3C).

Fig. 3. Characterizations of SMG mucus.

Fig. 3.

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(A) Solubility of HBE cell culture mucus and human and porcine SMG mucus in PBS. (B) Molecular size, i.e., radius of gyration (Rg), of the soluble fraction of SMG mucus and HBE mucus as determined by light scattering. n = 15 for HBE, n = 4 for human and n = 3 pig SMG mucus. Welch’s ANOVA test (P = 0.1003). (C) (i) Immunofluorescent staining of MUC5B and MUC5AC in non-CF SMG mucus strands. Scale bars, 10 μm. (ii) Intensity of MUC5B and MUC5AC signals on the yellow line in the left panel. (D) Glycan abundance of less complex (cores 1 and 3) glycans in HBE versus SMG mucus. Unpaired t test. n = 3 for each. (E) Abundance of glycans with shorter (≤3) monosaccharides in HBE versus SMG mucus. Unpaired t test. n = 3 for each.

Glycomics reveals that SMG mucins contain shorter, less-branched glycans

Glycomics mass spectroscopy of soluble MUC5B from SMG versus superficial HBE specimens from three matched donors was performed. SMG mucin–associated glycans contained a larger proportion of shorter, less-branched (cores 1 and 3) carbohydrate structures than HBE mucins (Fig. 3, D and E, fig. S1, and data file S1). Analyses of anionic O-glycans, in which the negative charge is imparted either as sulfate or as sialic acid, were also performed. Sulfation of extended O-glycans, but not sialic acid, was reduced in SMG relative to HBE MUC5B (figs. S1 and S2 and data files S1 and S2).

CF SMG mucus strands exhibit distinct features

We visualized strands in the insoluble SMG component from non-CF and CF specimens using scanning electron microscopy (SEM). Consistent with CF SMG mucus hyperconcentration (Fig. 2A), strand density was higher in CF SMG mucus (Fig. 4A). The width and length of SMG strands were similar in non-CF and CF (Fig. 4, B and C). Alignment analyses demonstrated that non-CF SMG mucus strands contained extraordinarily long arrays of aligned bundles (Fig. 4D), whereas CF SMG mucus strands exhibited randomly entangled bundles (Fig. 4E). Quantitation of orientational order revealed less order, i.e., more heterogeneity, in CF strands (Fig. 4, F and G).

Fig. 4. Characteristics of SMG mucus strands.

Fig. 4.

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(A) Concentration of strands within SMG mucus samples. Values were normalized by sample volume, imaged, and divided by the number of mean healthy strands. (B) Mean strand width of non-CF versus CF SMG mucus. (C) Mean strand length of non-CF versus CF SMG mucus. (D) SEM images of SMG strands from non-CF subjects. Inset was falsely colored according to angular alignment of pixels quantified with OrientationJ. (E) SEM images of CF SMG strands. Inset was falsely colored according to angular alignment of pixels. Samples from n = 9 (non-CF) and n = 4 (CF) donors. One to five strands per donor were measured. Mean of each donor was plotted. (F and G) Alignment of structures within non-CF versus CF SMG mucus. (F) Histograms of the relative alignment angles of SEM images in (D) and (E). Non-CF sample exhibits increased alignment of surface structures, whereas CF strand does not. (G) Quantification of alignment fraction for each clinical sample based on decreased numbers of bins of (D) and (E) to reflect slight deviations in angle from strand directionality. n = 3 each for non-CF and CF. (A) Mann-Whitney test. (B, C, and G) Unpaired t test.

Proline-rich protein 4 is an SMG-selective biomarker

Biomarkers selective for SMG secretions were investigated in human airway tissues. Four candidate glandular proteins, lactoferrin (LTF), zinc-α2-glycoprotein (AZGP1/ZAG), lysozyme (LYZ), and proline-rich protein 4 (PRR4), were screened on the basis of previous reports of selective SMG expression (32–34). RNA in situ hybridization (RNA-ISH) and immunohistochemistry (IHC) demonstrated that PRR4 was selectively expressed in SMG serous cells and not in superficial epithelia lining large or small airways in healthy or diseased subjects (Fig. 5, A and B, and fig. S3). In contrast, LTF, AZGP1, and LYZ were variably detected in surface epithelia or infiltrating cells (fig. S4, A to F). Quantitative polymerase chain reaction (qPCR) data from freshly dissected specimens from different pulmonary regions confirmed the specificity of PRR4 for SMGs (fig. S4G). Costaining for PRR4 and MUC5B distinguished SMG serous and mucous acini, respectively (Fig. 5C). SMG mucus passing through gland ducts exhibited copositivity for PRR4 and MUC5B (Fig. 5C, white arrow). In addition, SMG mucus strands exhibited colocalization of PRR4 and MUC5B (Fig. 5D). Accordingly, PRR4 was selected as a biomarker to identify the contribution of SMGs to human airway secretions.

Fig. 5. PRR4 in airways and clinical samples.

Fig. 5.

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(A and B) RNA-ISH and IHC for PRR4 in (A) healthy control and (B) CF human airways. Scale bars, 500 μm (low power images) and 200 μm (magnified images). (C) Immunofluorescent costaining of PRR4 and MUC5B in human proximal airways. SMG ductal mucus shows copositivity for PRR4 and MUC5B (white arrow). Scale bars, 200 μm. (D) Immunofluorescent costaining of PRR4 and MUC5B in human SMG mucus strands. (E and F) Label-free quantitative mass spectrometry (MS) analysis of (E) PRR4 and (F) human neutrophil elastase (HNE) peptides after tryptic digestion of induced sputum samples from healthy donors (n = 18), CF (n = 20), non-CF bronchiectasis (NCFB) (n = 10), and primary ciliary dyskinesia (PCD) (n = 9). Kruskal-Wallis test followed by Dunn’s multiple comparison test. (G) PRR4 content in bronchoalveolar lavage (BAL)–obtained flakes. (i) Immunofluorescent costaining of WGA and PRR4 for healthy control and CF mucus flakes (scale bars, 100 μm) and (ii) ratio of staining intensity of PRR4 to WGA in mucus flakes. n = 3 for healthy control and n = 6 for CF. Mann-Whitney test. (H) (i) Immunofluorescent staining of PRR4 and MUC5B for SMGs in healthy control versus CF tissues. Scale bars, 100 μm. (ii) Quantification of serous cell (PRR4) versus mucous cell (MUC5B) signals in SMG in healthy control versus CF. n = 9 for healthy control and n = 6 for CF. Welch’s t test.

Sputum PRR4 level is reduced in CF but not in other diseases

Mass spectrometry (MS)–based proteomics analyses were performed on induced sputum samples collected from healthy controls and subjects with chronic airway diseases to investigate whether disease status affected the SMG contribution to airway secretions. CF sputum exhibited markedly decreased PRR4 concentrations compared to both healthy (~30-fold reduction) and bronchiectatic disease controls (~300-fold reduction) (Fig. 5E). This result did not reflect PRR4 degradation by human neutrophil elastase (HNE), as sputum HNE levels were similar in CF and non-CF bronchiectasis (NCFB)/primary ciliary dyskinesia (PCD) subjects (Fig. 5F) and PRR4 was not degraded by coincubation with HNE in vitro (fig. S5).

Mucus flakes from CF subjects show reduced PRR4

Mucous flakes have been described in the insoluble material harvested by bronchoalveolar lavage fluid (BALF) from children with CF (10). Aliquots of these CF and control BALF samples were obtained and costained with a PRR4 antibody and wheat germ agglutinin (WGA) to localize mucin-associated glycans. The ratio of PRR4 to lectin staining was reduced in CF compared to healthy controls (Fig. 5G).

Healthy and CF airways have similar serous versus mucous cell proportion in SMG

To test whether reduced PRR4 levels in CF reflected reduced synthetic capacity, we quantified the proportion of serous versus mucous cells of SMGs in lungs resected from people with CF at time of transplant using PRR4 and MUC5B immunostaining. Tissues from healthy and CF subjects exhibited similar ratios of PRR4-positive serous to MUC5B-positive mucous cells (Fig. 5H).

DISCUSSION

CF airway surface epithelia exhibit hyperconcentrated mucus with raised osmotic pressures and cohesive strengths that are central to disease pathogenesis (4, 5, 9). However, less is known about native SMG mucus function and how it may be disturbed in CF. In this study, we tested the hypothesis that mucus hyperconcentration is a common defect driving CF pathophysiology in both airway surface epithelia and SMGs.

SMG mucus from pigs treated with anion/fluid secretion blockers was hyperconcentrated, consistent with previous reports by Ballard et al. (26, 35) and Joo et al. (18, 19) (Fig. 1A). Human CF SMG mucus was also hyperconcentrated compared to non-CF SMG mucus (Fig. 2A), consistent with defective fluid secretion (16, 36). Elevated mucus osmotic pressures were measured in mucus from CF-mimicking pig and CF SMGs that paralleled increased mucus concentrations (Figs. 1, B and C, and 2, B and C). Cohesive forces of human CF SMG mucus, also a function of increased mucus concentration, were also significantly elevated compared to control (Fig. 2D). TEM investigations revealed that SMG ducts of CF-mimicking pig and human CF tissues exhibited both mucus retention and, as predicted from osmotic pressure measurements, cilial compression (Figs. 1, D and E, and 2, E and F). These observations are consistent with previous reports of increased CF SMG mucus viscosity (29) and predict that CF SMG mucus secretion is hindered by both SMG mucus osmotic compression of ductal surfaces and increased cohesive forces that limit mucus separation from ductal orifices.

SMG mucus flows as a liquid but has been reported to contain strands that maintain their structure after secretion onto airway surfaces rather than dissolve into the ambient surface mucus layer (14, 15). We investigated the characteristics of SMG versus superficial epithelial soluble mucins and whether strand formation was unique to SMG mucus. Notably, superficial HBE mucus dissolved fully into PBS, leaving behind no strand-like structures, whereas ~50% of non-CF human and pig SMG mucus failed to dissolve into excess PBS (Fig. 3A).

Glycomics analyses for the HBE versus SMG mucus from matched donors revealed differences in MUC5B glycosylation with potential implications for strand formation. First, SMG mucin glycans consisted of larger proportion of shorter, less-branched (cores 1 and 3) carbohydrate structures than HBE mucins (Fig. 3, D and E, and data file S1). Mucin glycosylation with shorter glycans presents fewer hydroxyls for capture of water molecules through hydrogen bonding, a feature predicted to promote hydrophobic interactions between SMG MUC5B mucins and strand formation (37). Second, anionic O-glycans, in which the negative charge is imparted either as sulfate or sialic acid, were also measured. Sulfation of extended O-glycans, but not sialic acid, was reduced in SMG relative to HBE MUC5B, also consistent with a more hydrophobic, strand-prone SMG MUC5B mucin (figs. S1 and S2 and data file S1). The observation that a nonionic, nondenaturing detergent (IGEPAL) promoted dissolution of the insoluble SMG strands is consistent with glycomics predictions (table S1). The colocalization of a unique SMG-secreted protein with SMG strands, i.e., PRR4 (discussed below), also suggests a role for protein cross-linkers in strand formation (Fig. 5D). We also speculate that the additional glycosylation structures characteristic of SMG mucus broadens the spectrum of mucin-binding sites for bacteria and/or viruses, increasing the likelihood of pathogen clearance by mucociliary transport. Further functional investigations of SMG mucus are necessary to fully characterize the roles of SMG secretion in host defense.

Collectively, these data suggest specialized functions for surface versus SMG mucus in normal lung physiology. Superficial epithelial mucins likely dissolve rapidly upon secretion to replenish mucus cleared by transport. In contrast, strands secreted by SMGs maintain their structure on airway surfaces, i.e., “permanent” gels (10). We speculate that strands aggregate with inhaled particles after cough-stimulated secretion to form collections (masses) of sufficient size to promote cough clearance in times of stress (30).

Distinct features of CF SMG mucus strands were identified (Fig. 4). Consistent with CF SMG mucus hyperconcentration, strand density was higher in CF SMG mucus (Fig. 4A). The width and length of SMG strands were similar in non-CF and CF mucus and strand width approximated the inner diameter of mucus tubules (~15 μm in diameter), suggesting that strand width is defined by distal SMG duct structures (Fig. 4, B and C) (38). Notably, SEM demonstrated that non-CF human SMG mucus strands contained extraordinarily long arrays of aligned bundles, whereas CF SMG mucus strands exhibited randomly entangled bundles (Fig. 4, D and E). Quantitation of orientational order revealed less order, i.e., more heterogeneity, in CF strands (Fig. 4, F and G). We speculate that decreased alignment of CF SMG strands reflects abnormal strand formation in a flow-limited, hyperconcentrated environment.

Last, clinical investigations were performed to distinguish between the hypotheses that CF SMGs fail to secrete mucus versus to secrete a strand-laden mucus that accumulates on airway surfaces (12, 17–19). PRR4 was identified as a biomarker selective for SMG secretion to quantitate by mass spectroscopy the contribution of SMGs to airway secretions (Fig. 5, A to D, and fig. S4). NCFB and PCD subjects exhibited raised sputum PRR4 levels compared to healthy control subjects, consistent with SMG hypersecretion due to hypertrophied SMGs and/or increased stimulation of SMG secretion (39). In contrast, CF sputum exhibited markedly decreased PRR4 concentrations compared to either normal subjects or bronchiectatic disease controls. Consistent with these results were data demonstrating that mucus flakes in CF BALF exhibited reduced PRR4 relative to mucin staining than control subjects. The reduced PRR4 levels in CF did not reflect PRR4 degradation by HNE (fig. S5) or diminished PRR4 secretory capacity (Fig. 5H). Collectively, these data suggest that the SMG contribution to CF pathogenesis reflects reduced CF SMG secretion onto, not accumulation of SMG mucus strands on, proximal airway surfaces.

Our study has several limitations. Because of the limited availability of fresh human airway tissues, specimens from non-CF diseased (e.g., smokers, chronic bronchitis) subjects were included in some studies (labeled as “non-CF”). More detailed studies are necessary to fully characterize the SMG mucus properties in non-CF disease states to estimate the effect of tissue selection in this study. Another limitation is that maximal cholinergic stimulation of SMGs for 2 hours was required to generate sufficient SMG mucus volumes for analyses. Prolonged stimulation likely does not mimic the rapid (<1 min) stimulation of SMGs during cough. In addition, it is not known that HBE cultures recapitulate the mucin glycosylation manifest patterns in vivo. Although our recent data indicate that HBE mucus exhibits mucin glycosylation patterns comparable to endotracheal tube (ETT) mucus, our in vivo source of mucus (40), direct comparisons of HBE mucus, and in vivo superficial airway mucus for glycomics analyses are needed. A final limitation is that we have not characterized the function of PRR4. Investigation of function of PRR4 and other proline-rich protein families (fig. S4G) may reveal additional aspects of CF SMG pathogenesis.

In summary, SMG mucus exhibits unique characteristics configured for host defense, e.g., strand formation suitable for cough clearance. SMG mucus from human CF and CF-mimicking porcine airways is hyperconcentrated compared to non-CF SMG mucus. Hyperconcentrated SMG mucus exhibits increased osmotic pressures and cohesive forces predicted to retain mucus in SMG ducts and reduce secretion. The concentration of an SMG-selective secretory protein (PRR4) was lower in CF sputum than healthy controls and markedly lower than in subjects with non-CF bronchiectatic diseases, suggesting that CF SMG mucus secretion onto proximal CF airway surfaces is impaired. Accordingly, restoration of SMG host defense proteins/peptides on CF airway surfaces early in life may be therapeutic (41). Last, these data suggest that increased mucus osmotic/cohesive forces, a product of abnormal CFTR-dependent mucus hydration (hyperconcentration), provide a unifying mechanism to describe the muco-obstructive distal and proximal airway components of CF lung disease.

MATERIALS AND METHODS

Human airway dissection

Human tissues were provided by the University of North Carolina (UNC) Tissue Procurement and Cell Culture Core under protocols approved by UNC Office of Research Ethics Biomedical Institutional Review Board (IRB; numbers 03-1396 and 17-3281). Non-CF airway tissues were obtained from lungs of donors that could not be used for transplant. CF airways were obtained from lungs resected from patients with CF undergoing transplantation at UNC Hospitals. Excised tissues were transported in saline on ice, and all studies were conducted within 12 to 36 hours of lung removal. Proximal airways (second to fourth generation bronchi) were manually dissected. Intraluminal mucus was removed by washing with Ringer’s bicarbonate solution until the flushed solution appeared clear, and residual fluid was collected as completely as possible with positive-pressure pipettes.

Pig airway dissection

All procedures with swine performed at the University of South Alabama were conducted under protocols approved by the University of South Alabama Institutional Animal Care and Use Committee in compliance to the U.S. Public Health Service Policy on Humane Care and Use of Laboratory Animals. Pig bronchial dissection was performed as previously described (26). For the collection of SMG mucus for solubility and light scattering assays performed at UNC, freshly excised porcine airway tissues were obtained from a local slaughterhouse. Main stem or segmental bronchi were dissected in the laboratory.

Gland mucus collection

Pig and human airway SMG mucus collection was performed using established protocols with minor modifications (26, 36). The proximal end of an isolated airway was cannulated with polyethylene tubing and ligated with sutures. All distal branches of the excised preparation were tightly ligated with suture material to physically separate the airway lumen from the incubation buffer, enabling SMG mucus to be secreted into the lumen without contamination by incubation buffer [Krebs-Ringer bicarbonate (pH 7.4) under continuous gassing with 95% O2 and 5% CO2 at 37°C]. For the porcine main stem to segmental bronchi, both proximal and distal edges were sealed with silicon plugs. Ach chloride (A6625, Sigma-Aldrich) was added to the incubation solution at a concentration of 10 μM to stimulate SMG secretion. For porcine experiments, DMA (100 μM; A4562, Sigma-Aldrich) and/or Bum (10 μM; B3023, Sigma-Aldrich) were used together to block bicarbonate and/or chloride secretion, respectively. After 2-hour incubation interval with Ach with or without anion transport inhibitors, the bronchial tissues were removed from the buffer solution and decannulated. After wiping the outer surface of the preparation to remove buffer solution, the airway was opened longitudinally. Mucus secreted into the lumen was collected by aspiration with a 10-μl positive pressure pipette. In each experiment, % solid concentration was measured immediately (described below), and the remaining sample was kept frozen at −80°C until further investigation.

% Solid measurement

Percent solid concentrations (% solids) of human and porcine SMG mucus (3 to 5 μl) were measured gravimetrically as described previously (42). The mucus aliquots were weighed on a preweighed piece of aluminum foil. The samples were heated at 80°C in an oven overnight to allow the liquid content to evaporate completely. The final weight of the dried foil and mucus sample was determined and solid weight % (% solids) was calculated.

Osmotic pressure measurement

Osmotic pressures of human and pig SMG mucus (20 to 30 μl) were measured with a custom-designed osmometer incorporating a 10-kDa membrane as previously reported (5). Twenty to 30 μl of SMG mucus was placed into the fluid chamber, and the steady-state osmotic pressure of mucus was acquired by the pressure transducer placed on the other side of the osmotic membrane.

Cohesion measurements

The cohesive strength of human SMG mucus samples (20 to 40 μl) was measured using a custom-designed peel test device (30, 31). Twenty to 40 μl of an SMG mucus sample was placed on a mucus-binding mesh, and the peeling force to tear the mucus apart was measured.

TEM for porcine and human airway tissues

Airway tissues were immersed in 2% paraformaldehyde/2.5% glutaraldehyde/0.15 M sodium phosphate (pH 7.4) and stored for several days at 4°C. After postfixation for 1 hour with buffered potassium ferrocyanide–reduced osmium (1% osmium tetroxide in 0.1 M Sorenson’s buffer for porcine tissues or 1% osmium tetroxide/1.25% potassium ferrocyanide/0.15 M sodium phosphate for human tissues), the samples were washed in deionized water and then dehydrated through a graded ethanol and propylene oxide (43). The samples were infiltrated with a 1:1 and then a 1:2 mixture of propylene oxide and Poly/Bed 812 epoxy resin for 4 hours and infiltrated overnight in 100% resin (08792-1, Polysciences). Samples were transferred to embedding molds and polymerized at 60°C overnight. Using a diamond knife, 1-μm-semithin sections were cut, mounted on slides, and stained with 1% toluidine blue to examine by light microscopy and isolate regions of interest. Ultrathin sections (70 to 80 nm) were cut with a diamond knife and mounted on 200-mesh copper grids, followed by staining with 4% aqueous uranyl acetate for 12 min and Reynold’s lead citrate for 8 min (44). The samples were observed using a JEOL JEM-1230 TEM operating at 80 kV (JEOL USA), and images were acquired with the Gatan Orius SC1000 CCD Digital Camera and Gatan Microscopy Suite 3.0 software (Gatan). Quantitative measurement of cilial height and semiquantitative scoring of mucus retention in SMG ducts were performed by a blinded observer. Cilial height from the ductal cell surface to the tip of the cilia was quantified. Mucus retention was scored from 0 to 3 with 0.5 intervals.

SEM of SMG mucus

SMG mucus was diluted 10-fold in PBS and mixed for 2 hours at room temperature (RT). Samples were allowed to sediment onto a 3-aminopropyltriethoxysilane (APTES)–coated surface for 15 min before being fixed with 2% paraformaldehyde and 2.5% glutaraldehyde in PBS for another 15 min. The samples were washed three times with sterile water and dehydrated into ethanol using increments of 10% in 10-min increments. The samples were then critically point-dried, mounted onto an SEM specimen holder using copper tape, coated with 5-nm AuPd (108 Auto, Cressington), and imaged on SEM (FEI Helios Nanolab 600) using 5-kV accelerating voltage. Analysis of orientation was performed using OrientationJ (http://bigwww.epfl.ch/demo/orientation/) using a method adapted from Nerger et al. (45).

Human airway surface mucus production

HBE cells were isolated from nonsmoker donors without respiratory diseases by the UNC Tissue Procurement and Cell Culture Core as described (46). For glycomics comparison of SMG versus HBE mucus, isolated HBE primary (passage 0) cells, donor-matched to tissue sections from which SMG mucus was collected, were seeded on collagen (5005, Advanced BioMatrix)–coated Transwell inserts (3460, Corning) at a density of 0.3 million cells per insert. For solubility and light scattering studies, HBE primary cells were expanded as previously described (47). Passage 1 or 2 cells were seeded on collagen-coated Transwell inserts at a density of 0.3 million cells per insert. The cells were cultured under submerged condition until confluence, followed by air-liquid interface (ALI) culture with UNC ALI media (48). After a full differentiation confirmed by ciliation under light microscopic observation, secreted mucus was allowed to accumulate on apical surface for up to 3 weeks and collected by washing with PBS. The HBE mucus was pooled from multiple codes and then concentrated with ultrafiltration using 50-kDa molecular weight cutoff (MWCO) filter (UFC905096, Millipore) to the % solids of 4%.

Solubility measurements

Collected SMG and HBE mucus was diluted 40-fold with PBS and kept at 4°C for 4 days to allow the mucus to solubilize. After the 4 days, HBE mucus was completely dissolved, whereas the SMG mucus dissolved only partly in the PBS. Undissolved portions were diluted in PBS, and % solids of the insoluble part was calculated. The SMG supernatant, containing the soluble component of SMG mucus, was collected for light scattering. The PBS-insoluble SMG mucus was exposed to surfactant (1 mM in PBS; IGEPAL CA-630, Sigma-Aldrich), and solubility was reexamined 48 hours later.

Light scattering

Static light scattering for soluble components of SMG and HBE mucus were performed on a research goniometer (BI-200SM, Brookhaven Instruments), equipped with a 640-nm laser. A series of solutions with different concentrations were measured at scattering angles between 40° and 150°. The data were plotted as a Zimm plot, from which the radius of gyration (Rg) of the solutes was extracted. The log-normal size distribution of the solutes was obtained by cumulant analysis of the correlation function between 100 ns and 10 ms. Before measurements, all solutions were filtered through a 0.45-μm hydrophilic polyvinylidene difluoride syringe filter (Titan 3, Thermo Fisher Scientific) to remove dust particles and large cell debris.

Analysis of mucin O-glycosylation

O-glycans from HBE and SMG mucus were released by reductive β-elimination and analyzed by nanospray ionization multidimensional MS (NSI-MSn) as described (49, 50). Briefly, mucus samples were precipitated from cold 80% aqueous acetone, dried, and resuspended for reductive β-elimination. Following release and clean-up, O-glycans were permethylated and recovered by water–dichloromethane (DCM) extraction. Nonsulfated and sulfated O-glycans were recovered in the DCM and aqueous phases, respectively. The two phases were separated and dried, and resulting glycans were analyzed by NSI-MSn. MS data were acquired in both negative and positive ion modes to detect sulfated and nonsulfated glycans, respectively. For quantification of permethylated, nonsulfated glycans, a known amount of maltotetraose permethylated with isotopically heavy methyliodide was added to each O-glycan sample, and mucin glycan amounts were measured in reference to the standard (51). For sulfated glycans, the same quantification was not possible because of ionization suppression caused by the sulfate moiety. Therefore, sulfated glycans were quantified on the basis of the signal intensity measured in the mass spectrometer. Graphical representations of monosaccharide residues are presented in accordance with the broadly accepted Symbolic Nomenclature For Glycans, and O-glycan analysis was performed according to the MIRAGE (minimum information required for a glycomics experiment) guidelines (52). The raw data were deposited to an online repository GlycoPOST (https://glycopost.glycosmos.org/) with the accession number of GPST000241.

Tissue slide preparation for RNA-ISH and IHC

Human airway and lung tissues were provided by the UNC Tissue Procurement and Cell Culture Core. The donors included healthy controls (nonsmokers without any history of respiratory diseases), CF, asthma, NCFB, and PCD. Tissues were collected from multiple generations, i.e., trachea, main bronchi, and distal lungs with bronchioles. The dissected tissues were fixed with 10% neutral buffered formalin for 24 to 36 hours and embedded in paraffin. The formalin-fixed paraffin-embedded (FFPE) blocks were cut to 5 μm.

RNA in situ hybridization

RNA-ISH was performed using the RNAscope 2.5 HD Reagent Kit-RED (322350, Advanced Cell Diagnostics) according to the manufacturer’s instructions and published protocols (47). Probes targeting PRR4 (522791), LTF (425101), AZGP1 (401861), and LYZ (421441), in addition to positive and negative controls (310041 and 310043, respectively), were hybridized at 40°C for 2 hours in the HybEZ oven (241000), followed by signal amplification and washing. Signals were visualized by Fast Red, followed by counterstaining with hematoxylin. The probe targeting human housekeeping gene Ubiquitin C (UBC, 310041) served as a positive control to evaluate RNA quality. A bacterial gene, Bacillus subtilis dihydrodipicolinate reductase (DapB, 310043) served as negative control. The images were acquired using an Olympus VS200 slide scanner light microscope with a 60× 1.42 numerical aperture objective.

Immunostaining

FFPE tissue sections were baked at 60°C for 2 hours, deparaffinized three times with xylene for 10 min, and rehydrated with graded ethanol. Antigen retrieval was performed by boiling slides in 0.1 M sodium citrate (pH 6.0; 1200-W microwave settings: 100% power for 6.5 min, followed by 60% power for 6 min twice). After cooling, quenching of endogenous peroxidase was performed with 0.5% hydrogen peroxide in methanol for 15 min. Then, slides were blocked with Blocking One (03953-95, Nacalai Tesque) at RT for 30 min. The primary antibodies rabbit anti-PRR4 (PA5-59883, Invitrogen; 1:2500), anti-LTF (10933-1-AP, Proteintech; 1:1000), anti-ZAG (13399-1-AP, Proteintech; 1:1000), and anti-LYZ (MA5-32154, Invitrogen; 1:1000) were applied to slides and incubated at 4°C overnight. Sections were washed with PBS and incubated with biotinylated donkey anti-rabbit immunoglobulin G (IgG) secondary antibodies (711-065-152, Jackson ImmunoResearch; 1:400) at RT for 1 hour. Slides were treated with VECTASTAIN Kit (PK-4000, Vector Laboratories) according to the manufacturer’s instructions. Sections were stained with 3,3′-diamino benzidine and counterstained with Fast Red. Images were acquired as described above. For immunofluorescent costaining, tissue sections were treated with rabbit anti-PRR4 antibody and mouse anti-human MUC5B antiserum (raised in-house, M-25; 1:1000) at 4°C overnight, followed by fluorescent staining with Alexa Fluor Plus 647–conjugated donkey anti-rabbit IgG (A32795, Invitrogen; 1:1000) and Alexa Fluor Plus 555–conjugated donkey anti-mouse antibody (A32773, Invitrogen; 1:1000) at RT for 30 min. After autofluorescence quenching (SP-8400-15, Vector Laboratories), tissue was mounted with 4′,6-diamidino-2-phenylindole (DAPI)–containing medium (P36931, Invitrogen). Fluorescent images were acquired using Olympus VS200 scanner with DAPI (488 nm), Cy3 (555 nm), and Cy5 (647 nm) filters. Isotype controls were prepared using rabbit and/or mouse IgG (011-000-003 and 015-000-003, Jackson ImmunoResearch). Volume ratio of serous (PRR4 positive) cells to mucous (MUC5B positive) cells volume in SMGs was quantified with Olyvia (Olympus) and Visiopharm (Visiopharm) software.

Immunocytochemistry of SMG mucus

SMG mucus was diluted 10-fold in PBS and mixed for 2 hours at RT. Samples were allowed to sediment onto an APTES-coated surface for 15 min before being fixed with 2% paraformaldehyde for another 15 min. Samples were washed three times with PBS. Samples were then incubated with PRR4 antibody and MUC5B antisera or MUC5B antibody (H-300, Santa Cruz Biotechnology; 1:1000) and MUC5AC antibody (45 M1, Invitrogen; 1:1000) at 4°C overnight, followed by washing three times with PBS and incubated with secondary antibodies overnight at 4°C. Samples were washed again, stained with DAPI (D9542-5MG, Sigma-Aldrich; 1:10,000) for 15 min, then washed, and mounted onto slides (9990402, Thermo Fisher Scientific). Images were acquired with a Zeiss 880 confocal microscope. To view colocalization of MUC5B and MUC5AC, Airyscan acquisition mode was used for imaging (Fig. 3C).

RNA isolation from multiple regions of human airways

Freshly excised airway tissues from healthy nonsmoker donors were dissected under light microscopy. Small (~1 cm2) sections that contained surface and submucosal tissue were dissected from trachea and main bronchus. Small bronchioles were carefully dissected from surrounding alveolar tissue. Distal lung parenchyma sections of 0.5 to 1cm2 in size were cut as alveolar tissues. Dissected specimens were minced with a homogenizer, and total RNA was extracted with TRI reagent (T9424, Sigma-Aldrich) and an RNA extraction kit (R2052, Zymo Research).

cDNA synthesis and reverse transcription qPCR

Two hundred nanograms of RNA was used for synthesizing cDNA (1708840, Bio-Rad) according to the manufacturer’s instructions. qPCR was performed with QuantStadio 6 (Thermo Fisher Scientific). TaqMan PCR probes (Life Technologies) used for the investigation of SMG markers included PRR4 (Hs00200615_m1), PRB1 (Hs00818764_m1), PRB3 (Hs00818925_m1), PRB4 (Hs00864002_m1), PRH2 (Hs00818765-mH), LTF (Hs00914334_m1), AZGP1 (Hs00426651_m1), and LYZ (Hs00426232_m1). A marker for distal bronchioles and alveoli, SFTPB (Hs00167036_m1), was measured to assure where airway regions were obtained from. TBP (Hs00427620_m1) was chosen for the internal control. All probes were designed to span exons.

Label-free quantification MS for induced sputum

Induced sputum samples were obtained from the UNC sample repositories [IRB numbers 16-3142 (NCFB), 15-2431 (healthy controls and CF), and 02-0948 (PCD)]. One hundred microliters of samples were prepared with the Filter-Aided Sample Preparation method. Briefly, samples were denatured with 800 μl of 6 M GuHCl (pH 8.0) and reduced by adding dithiothreitol (DTT) to a final concentration of 20 mM for 1 hour at 65°C. After reduction, samples were alkylated with iodoacetamide (I1149, Sigma-Aldrich) at a final concentration of 50 mM for 1 hour at 25°C in the dark. Samples were centrifuged at 14,000g for 10 min, and the filter was washed twice with 50 mM ammonium hydrogen carbonate (NH4HCO3). A total of 0.5 μg modified trypsin (proteomics grade, T6567, Sigma-Aldrich) was added, and samples were incubated for 18 hours at 37°C. The peptides solutions were concentrated by vacuum centrifugation and dissolved in 30 μl of 0.1% formic acid. MS was performed with a Dionex ultimate 3000 RSLCnano system coupled to a hybrid quadrupole orbitrap mass spectrometer with a Nano spray source (Thermo Fisher Scientific Q-Exactive). Samples (3 μl) were loaded into a trap column (Acclaim PepMap 100, 100 μm × 2 cm, nanoViper C18, 5 μm, 100 Å), at 5 μl/min with aqueous solution containing 0.1% (v/v) trifluoroacetic acid and 2% acetonitrile. The column used for peptides separation was an Acclaim PepMap RSLC, 75 μm × 15 cm, nanoViper C18, 2 μm, 100 Å. Liquid chromatography–MS runs were 120 min long, with two liquid phases: 97% water, 3% acetonitrile, and 0.1% formic acid (buffer A) and acetonitrile and 0.1% formic acid (buffer B). The gradients (percentage of buffer B) of the run (120 min) applied were the following: 0 to 7 min, 4%; 7 to 90 min, 30%; 90 to 105 min, 80%; 105 to 120 min, 4%. Data were acquired at a resolution of 70,000 at mass/charge ratio (m/z) of 200, target automatic gain control value of 5 × 105, maximum fill times of 200 ms, and a multiplex degree of 6 with an isolation width of 3 m/z.

Proteins were identified by searching against the most current human database with Proteome Discover 1.4, setting a maximum of two missed cleavage sites for trypsin, oxidation of methionine (+15.995 Da) as a dynamic modification, and carbamidomethylation of cysteine (+57.0210 Da) as static a modification. Proteins were quantified using Scaffold (Proteome Software), using the normalized total precursor intensity, with 95% probability by the Scaffold Local false discovery rate algorithm.

PRR4 and WGA staining for airway mucus flakes

Mucus flakes were obtained from BALF from healthy nonsmoker adult subjects and school-age CF subjects as previously reported under IRB-approved protocol at UNC Chapel Hill (IRB numbers 91-0679 and 11-1445) (53). All studies were approved by the UNC Ethics Committee and informed consents from the subjects or parents obtained before any study procedures. BALF (10 μl) was applied to glass microscopy slides using a cytocentrifuge (StatSpin CytoFuge 2, Beckman Coulter). Slides were fixed with 10% neutral buffered formalin, washed with PBS, and blocked with 3% bovine serum albumin at RT for 1 hour. PRR4 and mucus were immunohistochemically stained by PRR4 antibody (1:1000) and biotinylated WGA (BK-1000, Vector Laboratories; 1:1000), respectively, at 4°C overnight. Slides were washed three times with PBS, followed by incubation with secondary antibodies [Alexa Fluor Plus 488–conjugated donkey anti-rabbit IgG (A32795, Invitrogen; 1:1000), Abnova Fluorescent Dye 594-I–conjugated streptavidin (U0291, Abnova; 1:1000), and DAPI (D3571, Invitrogen; 1:1000)] at RT for 1 hour. The slides were then washed with PBS and mounted with FluorSave-mounting media (345789, Millipore). The staining images were obtained with an Olympus FV1000 confocal microscope using a 20× objective and uniform settings. Average PRR4 intensity was normalized to overall mucus signal by dividing by the average WGA intensity.

Production of recombinant PRR4 protein

PRR4 cDNA was obtained from open reading frame clone plasmid (OHu15631, GenScript). PRR4 cDNA without a signal peptide sequence (corresponding to amino acids 17 to 134) was cloned into the pM-secSUMOstar Vector (7121, LifeSensors). SUMOstar-PRR4 vectors were transfected into Expi293 cells (1 mg of DNA per liter of transfection) per the manufacturer’s instructions, with culture supernatants harvested and sterile-filtered on day 3 after transfection. Supernatants were then concentrated, buffer-exchanged by tangential-flow filtration into TALON-binding buffer [50 mM sodium phosphate (pH 7.4), 500 mM NaCl, 5 mM imidazole, 0.05% sodium azide, and 10% glycerol], and then loaded onto hand-packed 1-ml TALON (635507, Takara Bio) columns recirculating overnight at 1.5 ml/min. Twenty-four hours later, columns were changed to flow-through and run until all supernatants were loaded. Columns were then washed with 20-column volume (CV)–binding buffer, followed by binding buffer with 10 mM imidazole. Proteins were then eluted with 20 CV-binding buffer supplemented with 150 mM imidazole. Fractions were assessed by SDS–polyacrylamide gel electrophoresis (PAGE), and those containing bands corresponding to His-SUMO-tagged PRR4 (10 mM wash and elution) were pooled. Pooled fractions were then dialyzed against PBS with 2 mM DTT (3.5-kDa MWCO). SUMOstar protease (SP4110, LifeSensors; 10 U/mg of substrate) was added to dialyzed protein, and the reaction was incubated overnight at 4°C to cleave the His-SUMO tags. Cleaved proteins were then passed back over the 1-ml TALON columns to separate the His-SUMO tags from the cleaved PRR4 proteins. Subtractive columns were washed and eluted as described above. Fractions were again assessed by SDS-PAGE to confirm separation, with flow-through and 5 mM imidazole fractions pooled, concentrated, and dialyzed into PBS (3.5-kDa MWCO). The final purified protein was confirmed to be PRR4 by both Western blotting and MS.

Confirmation of no degradative effect for PRR4 by neutrophil elastase

To quantitate the potential effects of proteolysis on the proteomics results, ETT mucus samples were treated with designated quantities of HNE (10 to 50 μg/ml; 324681, Sigma-Aldrich). ETT mucus was obtained as previously reported from ETT tips following postsurgical extubation at UNC General Hospital under UNC IRB–approved protocols (IRB number 17-2211) (40). Aliquots of ETT mucus (500 μl) were treated with 10, 20, and 50 μg of HNE and rotated/mixed for 30 min and then incubated at 37°C for overnight. PBS was added to the control. PRR4, MUC5B, and MUC5AC peptide intensities were measured by MS as described above (fig. S5).

Validation of mouse anti-MUC5B antiserum for immunostaining

The mouse anti-MUC5B antiserum raised in-house was not previously published. As a validation, we performed IHC of MUC5B for human tracheal serial sections with the antiserum (M-25; 1:1000) and previously established commercial rabbit anti-MUC5B antibody (H-300, Santa Cruz Biotechnology; 1:1000). Staining with these antibody and antiserum appeared identical, confirming the sensitivity and specificity of the novel antiserum (fig. S6).

Statistical analyses

Statistical analyses and figure creation were performed with GraphPad Prism 9.0.2. Each single measured/calculated value and means ± SEM were plotted in the graphs unless otherwise specified. Statistical methods used were described in the figure captions. Statistical tests were two-tailed whenever appropriate.

Acknowledgments

We thank the UNC CF Center Tissue Procurement and Cell Culture Core for providing human airway tissues and primary cells and the UNC Animal Histology Core for paraffin embedding and sectioning the tissues. We also thank H. Dang for statistical analyses and E. Roe for editing the manuscript.

Funding: This research was supported by grants from National Institutes of Health (NIDDK UH3 HL123645, P01 HL110873, R01 HL136961, P30 DK 065988, P01 HL108808, and R01HL125280), and Cystic Fibrosis Foundation (BOUCHE15R0 to R.C.B., BUTTON19G0 to B.B., FREEMA19G0 to R.F., OKUDA19I0 and OKUDA20G0 to K.O., KATO20I0 to T.K., HILL20Y2-OUT and HILL19G0 to D.B.H., EHRE20XX0 to C.E., and KESIME17G0 to M.K.), and a research grant from Cystic Fibrosis Research Incorporation to K.O. T.K. was supported by the Senior Research Training Fellowship by the American Lung Association (RT-575362). M.R.M. was supported by the Postdoctoral Research Fellowship by the Cystic Fibrosis Foundation (MARKOV18F0). Glycomic analysis of airway mucins was supported by grants from the National Institutes of Health (NIGMS P41GM103490 and Common Fund U01GM125267 to M.T.). SEM of SMG mucus strands was performed at the Chapel Hill Analytical and Nanofabrication Laboratory (CHANL), a member of the North Carolina Research Triangle Nanotechnology Network (RTNN), which was supported by the National Science Foundation, grant ECCS-1542015, as part of the National Nanotechnology Coordinated Infrastructure (NNCI). The UNC Hooker Imaging Core Facility, the Microscopy Services Laboratory, and the Protein Expression & Purification Core Facility were all supported in part by P30 CA016086 Cancer Center Core Support Grant to the UNC Lineberger Comprehensive Cancer Center.

Author contributions: T.K., S.T.B., B.B., and R.C.B. conceived the project and designed experiments. S.H.R. provided human airway tissues and primary cells. T.K., G.V.S., and S.T.B. collected pig SMG mucus. T.K. collected human SMG mucus. T.K., H.P.G, H.T., H.T.L., and B.B. performed % solids, osmotic pressures, and cohesion measurements. G.V.S. performed solubility and light scattering assays. K.A., M.P., and M.T. performed glycomics analyses. M.J.P. and R.F. acquired SEM and immunofluorescent images of SMG mucus strands and characterized their structural properties. K.A.B. and K.K.W. acquired electron microscopy images of porcine and human airway tissues. K.O. prepared human histology sections and RNA samples from human airway tissues. R.C.G. performed RNA-ISH. T.K., S.M.B.C., and T.A.B. performed immunostaining and reverse transcription qPCR. G.R. and M.K. performed proteomic analysis. M.R.M, C.B.M., and D.B.H performed analyses of mucus flakes. C.B.M. and C.E. provided mouse anti-MUC5B antiserum. L.J.F. performed PRR4 overexpression and purification. T.N. and M.R. provided feedback to overall work. T.K., B.B., W.K.O., and R.C.B. analyzed and interpreted the data. B.B. and R.C.B. supervised the project. T.K., R.F., W.K.O., B.B., and R.C.B drafted and finalized the manuscript.

Competing interests: The authors declare that they have no competing interests.

Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. The human lung tissues, cells, and specialized culture media can be provided by S.H.R. pending scientific review and a completed material transfer agreement. Requests for the materials should be submitted to S.H.R. (scott_randell@med.unc.edu).

Supplementary Materials

This PDF file includes:

Figs. S1 to S6

Table S1

sciadv.abm9718_sm.pdf (2.4MB, pdf)

Other Supplementary Material for this manuscript includes the following:

Data S1 and S2

sciadv.abm9718_data_s1_and_s2.zip (249.6KB, zip)

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Associated Data

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

Supplementary Materials

Figs. S1 to S6

Table S1

sciadv.abm9718_sm.pdf (2.4MB, pdf)

Data S1 and S2

sciadv.abm9718_data_s1_and_s2.zip (249.6KB, zip)

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