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Okuda, K.;  Shaffer, K.M.;  Ehre, C. Mucins and Cystic Fibrosis Transmembrane Conductance Regulator. Encyclopedia. Available online: (accessed on 23 June 2024).
Okuda K,  Shaffer KM,  Ehre C. Mucins and Cystic Fibrosis Transmembrane Conductance Regulator. Encyclopedia. Available at: Accessed June 23, 2024.
Okuda, Kenichi, Kendall M. Shaffer, Camille Ehre. "Mucins and Cystic Fibrosis Transmembrane Conductance Regulator" Encyclopedia, (accessed June 23, 2024).
Okuda, K.,  Shaffer, K.M., & Ehre, C. (2022, September 15). Mucins and Cystic Fibrosis Transmembrane Conductance Regulator. In Encyclopedia.
Okuda, Kenichi, et al. "Mucins and Cystic Fibrosis Transmembrane Conductance Regulator." Encyclopedia. Web. 15 September, 2022.
Mucins and Cystic Fibrosis Transmembrane Conductance Regulator

Mucociliary clearance is a critical defense mechanism for the lungs governed by regionally coordinated epithelial cellular activities, including mucin secretion, cilia beating, and transepithelial ion transport. Cystic fibrosis (CF), an autosomal genetic disorder caused by the dysfunction of the cystic fibrosis transmembrane conductance regulator (CFTR) channel, is characterized by failed mucociliary clearance due to abnormal mucus biophysical properties. 

mucus mucins cystic fibrosis (CF) CFTR single-cell transcriptomics

1. Introduction

Cystic fibrosis (CF) is an autosomal genetic disorder characterized by the dysfunction of the cystic fibrosis transmembrane conductance regulator (CFTR) protein, an ion channel transporting Cl and HCO3 that is expressed in various tissues, notably in mucus-producing organs [1]. One of the primary drivers of CF pathogenesis is the accumulation of a thick, adherent mucus, particularly in the lungs, gut, and pancreatic ducts [2]. Dysfunction of CFTR causes fluid hyperabsorption and low HCO3 concentrations, which can affect mucus network organization in numerous ways that will be discussed [3][4]. The aberrant properties of CF mucus in the lungs affect mucociliary clearance (MCC) and result in airway muco-obstruction, chronic inflammation, bacterial infection, decline in lung function, and, eventually, respiratory failure.
CF was first described in 1938 by Dr. Dorothy Andersen as “cystic fibrosis of the pancreas” after she observed histological sections of the pancreas of children who died from malnutrition [5]. In 1945, recognizing that CF affected more than the pancreas, Dr. Sydney Farber referred to the disease as “mucoviscidosis” due to the abnormally thick mucus that individuals with the disease produced [6]. Once considered a fatal disease of childhood, the advancement of therapeutics and treatment options for CF patients over recent decades increased life expectancy to nearly 50 years. In the early days, commonly used therapies focused on addressing symptoms and reversing airway obstruction via physiotherapy, bronchodilators, osmotic agents, and mucolytics/recombinant human DNase (rhDNase) [2][7][8]. While effective at slowing the progression of lung disease, these therapeutic approaches failed to address the root cause of the disease, CFTR dysfunction.
CFTR mutations are grouped in six different classes according to their impact on protein synthesis (Class I causing absence of CFTR protein), folding (Class II causing trafficking defects), or function (Class III–VI causing defects in channel gating, quantity, and/or stability). In the past decade, CFTR modulators, which consist of small molecules acting systemically to restore CFTR function, have gradually changed the way physicians care for CF patients and, more importantly, have significantly improved the quality of life of patients eligible to take these medications. In 2012, ivacaftor (VX-770), a CFTR potentiator, was the first modulator therapy to be approved to restore CFTR function with gating mutations, such as G551D [9][10][11]. In addition to improved airway clearance and pulmonary function (e.g., FEV1), patients undergoing ivacaftor treatment experienced increased BMI, decreased sweat chloride concentration, fewer hospitalizations, and reduced incidence of Pseudomonas aeruginosa infection. Although this compound was later approved for additional mild mutations (e.g., P67L, R117H), only a small fraction (<10%) of people living with CF could benefit from taking this medication [12].
In the following years, research focused on correcting the function of the most common mutation, F508Del, as roughly 85% of the CF population carries at least one copy of the class II mutation. Two additional modulator therapies were developed combining ivacaftor with a CFTR corrector compound, lumacaftor (VX-809) or tezacaftor (VX-445) [13]. While both therapies, lumacaftor/ivacaftor (LUM/IVA or Orkambi) and tezacaftor/ivacaftor (TEZ/IVA or Symdeko), were shown to improve overall lung function in patients homozygous for the F508Del mutation, efficacy remained modest compared to ivacaftor in patients with gating mutations [14][15]. In 2019, a triple-combination modulator drug, elexacaftor/tezacaftor/ivacaftor (ETI or Trikafta), combining VX-660, VX-445, and VX-770, was approved for the treatment of patients with at least one copy of F508Del mutation. The addition of a second CFTR corrector, elexacaftor, to TEZ/IVA resulted in significantly increased drug efficacy, producing a 15% increase in FEV1 and a 63% decrease in pulmonary exacerbations [16][17][18].

2. Affected Regions in CF Lung Disease

2.1. Role of CFTR in the Lungs

In light of the fact that CFTR is a transmembrane channel transporter for chloride and bicarbonate, CF lung disease pathogenesis reflects abnormal ion transport. Despite general agreement on this notion, controversy remains in identifying the specific links between abnormal ion transport and CF lung disease. In CF, defective ion transport produces: (1) reduction in airway surface liquid (ASL) volume, mucus hyperconcentration, and an early muco-inflammatory state predisposing to bacterial infection [19][20]; and (2) abnormalities in ASL pH leading to defects in airway epithelial host innate defense properties, resulting in persistent bacterial infection [21][22][23]. In addition to pathophysiological consequences related to abnormal ion transport, it is important to understand which compartments of the lung are affected in CF, as these specific regions can be the primary targets for therapeutic strategies. CFTR dysfunction typically causes no change to the alveolar region but critically affects the conducting airways, comprised of two distinct regions: (1) the proximal/large (tracheobronchial) airways that contain submucosal glands (SMGs) and cartilage; and (2) the distal/small (bronchiolar) airways (<2 mm at the diameter) that constitute the major surface area of the airways within the lung [24][25][26].

2.2. Structural and Regional Specificity of the Lungs

Compartmentalization of the lung raises two important questions: (1) how do the superficial epithelial or SMG compartments contribute to disease; and (2) which airway region, large or small, is the disease-initiating/vulnerable region or the starting point of CF pathogenesis? Studies aimed at answering these questions have shed light on the structural and regional specificity of the lungs.
The SMGs, restricted to the large airways, are a source of electrolytes, fluid, host defense proteins, and secretory mucins, predominantly MUC5B [27][28][29][30][31]. Compared to the superficial epithelium that produces mucus at baseline and maintains constant directional motion, the SMGs secrete large volumes of mucus following adrenergic or cholinergic stimulation to aid in host defense and galvanize airway clearance. In CF, dysfunctional SMG mucus secretion contributes to impaired MCC in the large airways. Small airways lack SMGs, and the superficial epithelium must clear mucus from the distal regions, a process that relies heavily on the coordinated beating of the cilia. However, mucus hyperconcentration can cause compression of the cilia and subsequent mucostasis [32]. The limited backup mechanism, i.e., lack of SMGs, to clear mucus in small airways predisposes this region to impaired MCC and disease. Studies have postulated that the superficial epithelium of small bronchiolar airways is one of the most severely affected regions in CF [19][20].

2.3. Clinical Observations Related to Small Airway Diseases in CF

Evidence for small airways as the central site of CF pathogenesis has been based on pathology [33][34][35][36][37][38][39], pulmonary function [38][39], imaging [40][41][42][43][44], and lower airway sampling studies [19][36][37]. Pathologic studies of CF lungs, including CF children who died early of the disease, have identified small airway mucus plugs as a routine feature of this syndrome [33][34]. Micro-CT studies of CF lungs harvested at the time of transplant revealed that mucus plugs develop after roughly the 6th generation of airways and progressively increase in frequency in the distal bronchiolar regions [45]. Moreover, pulmonary function tests designed to detect regions of airflow obstruction confirmed the presence of small airway obstruction as the first detectable spirometry measure associated with CFTR dysfunction [38][39]. Consistent with this finding, radionuclide MCC studies have demonstrated the predominance of peripheral (small airways) over central (large airways) clearance defects [40][46]. Further supporting these observations, studies of bronchoalveolar lavage fluids performed on CF preschoolers have shown that the incipient mucus material harvested from CF lungs consisted of abundant and irregular/rough MUC5B/MUC5AC mucus flakes, indicating disturbance of mucus homeostasis in the distal airways in CF lung disease (see mucin distribution and effects on mucin network below) [19].

2.4. Airflow, Shear Forces, and Mucus Clearance in the Small Airways

Regional observations may explain why CF small airways are so vulnerable to mucus plugging. First, studies describing the relationship between pulmonary airflow and mucus adhesion to cell surfaces suggest that airflow-induced shear forces required to dis-adhere and clear mucus are restricted to the most proximal region (trachea) [47]. In the small airways, airflow rates are ~2 logs lower than in the proximal airways, which correlates with proportionate decrements in shear forces and provides physical reasons for small airways vulnerability in CF [48]. Second, the absence of submucosal gland secretions is likely to contribute to the failure of mucus clearance in the small airway regions. Third, ciliated cells in the small airways are sparse and possess shorter cilia [49][50], limiting the mechanical forces for mucus transport and peripheral MCC [51]. These findings highlight the importance of understanding how each MCC component contributes to increased CF mucus viscoelasticity and/or is associated with the failure of MCC in the small airway regions. In the next chapters, researchers focus on site-specific expression and function of secretory mucins and CFTR, both of which are major components that regulate airway mucus biophysical properties.

3. Regional Distribution of Secretory Mucins in the Conducting Airways

3.1. Mucin Concentrations in Health and Disease

MUC5AC and MUC5B, the dominant gel-forming mucins present in the mucus layer lining the airways, play distinct pathophysiological roles in the lungs pertaining to their individual expression patterns and biochemical/biophysical properties [52][53][54][55]. In normal airway secretions, the concentration of MUC5B is reported to be 10 times higher than MUC5AC, with the latter detected in trace amounts in healthy subjects [56][57]. While total mucin concentration increases in a variety of muco-obstructive lung diseases, including CF [20][58], non-CF bronchiectasis [59], COPD [57][60], and asthma [61], the ratio of MUC5AC/MUC5B differs depending on disease phenotype. Compared to healthy subjects, both MUC5AC and MUC5B proteins are elevated in adult CF airway secretions [20][58] and in COPD sputa [57][60], while elevated MUC5AC but not MUC5B is a hallmark of asthmatic sputum [61][62].

3.2. Airway Mucins and Their Functions

Different organs produce MUC5AC and MUC5B in the human body; for instance, the stomach secretes MUC5AC, and the female reproductive tract secretes MUC5B, suggesting distinct functional properties. Studies with Muc5b-deficient mice demonstrated that Muc5b was required for airway defense and MCC [63], despite the fact that the large Muc5b SMG reservoir is constrained to the proximal trachea in mice [64]. In contrast, Muc5ac-deficient mice showed no defect in airway defense and/or clearance [63]. Studies performed on human bronchial epithelial (HBE) cell cultures showed that a pathological MUC5AC-rich mucus induced by IL-13 stimulation is more adherent to airway epithelial surfaces compared to the typical MUC5B-rich mucus, which correlates with reduced mucociliary transport (MCT) [62][65]. A recent study analyzed macromolecular mucin assembly using Calu3 cells genetically manipulated to produce either MUC5AC or MUC5B [66]. Electron and atomic force microscopy revealed that MUC5B adopted a linear pattern, while MUC5AC exhibited a high degree of branching. Quartz crystal microbalance (QCM) dissipation analyses indicated that MUC5AC formed a denser, stiffer, and more viscoelastic mucus with a higher order of oligomerization, providing an explanation for the more adhesive properties of MUC5AC as compared to MUC5B. A mouse model overexpressing Muc5ac in the lungs demonstrated protection against influenza infection via the trapping of viruses through the terminal sialic acids cloaking the mucin protein core [67]. In contrast, mice lacking Muc5ac were unable to expulse enteric parasites swiftly [68]. These in vivo and in vitro models demonstrated that MUC5AC and MUC5B have distinct functions and both mucins are required to protect the lungs against inhaled particles and/or pathogens.

3.3. MUC5AC and MUC5B Distribution in the Respiratory Tree

In addition to distinct biophysical properties, the unique distribution of MUC5B and MUC5AC in the lungs contributes to their specific roles (i.e., gliding vs trapping). The classic paradigm for the human respiratory tree described MUC5B as a predominant feature of SMG secretion and MUC5AC as a marker of goblet cells that originated from the superficial epithelium after resident cells adopted a mucin-producing phenotype [27][69]. However, data generated from mice indicated that the superficial epithelium secretes Muc5ac along with Muc5b, which was mediated by non-goblet, secretory club cells [70][71]. Recent human studies conducted on healthy subjects confirmed extensive MUC5B mRNA and protein expression from both SMGs and superficial epithelial cells [72][73]. Transcript and protein mapping for MUC5AC and MUC5B from the trachea to the terminal bronchioles in healthy subjects revealed different expression patterns for these secretory mucins, with the most striking difference located in the small conducting airways, while the terminal and respiratory bronchioles remain mucin-free areas [73]. In healthy individuals, cells positive for the club cell secretory protein (CCSP) produce both MUC5AC and MUC5B throughout the large conducting airways. Hence, both mucins are expressed in the proximal airways. At approximately the 10th generation of bronchiolar airways (diameter <2 mm), MUC5AC expression ceases while MUC5B expression remains, leaving MUC5B as the sole secretory mucin produced in the distal airways. Morphometric calculation determined that MUC5B expressed in the surface area of the small airways is slightly greater than MUC5B expressed in the SMGs in the lung, designating the distal airway superficial epithelium as the major source of MUC5B in the lungs. While both MUC5B and MUC5AC protein levels were elevated in CF airway secretions, the expression of MUC5B transcripts predominantly increased in the small airway epithelia, suggesting the importance of MUC5B overproduction in this region in CF small airway pathogenesis [74].

4. Cell Types Expressing CFTR in Human Conducting Airways

4.1. Early Findings on Cell Types Expressing CFTR

Determining the cell types responsible for CFTR expression and function is fundamentally important for the development of effective therapies based on CFTR gene transfer/repair. Targeting the appropriate cells that normally express CFTR in the lungs and understanding how these cells use innate defense mechanisms are critical steps for the development of novel molecular approaches aimed at restoring CFTR gene expression. Over the last decades, significant controversy has surrounded the cell types expressing CFTR in human airways. Early studies based on immunohistochemistry indicated the localization of CFTR within non-ciliated cells in the superficial epithelium, including CK14-positive basal cells in the large airways [75] and CCSP-positive cells in the small airways [76]. Conversely, a separate study showed CFTR protein at the apical plasma membrane of ciliated cells in the superficial epithelium [77]. Several possible limitations to these morphometric studies may explain the discrepancy, including sensitivity and specificity of antibodies targeting CFTR [78], differences in airway regions examined, and lack of systematic quantitation for CFTR signals. However, multiple original studies identified a rare cell type called “hot cells” with intense CFTR signals in the submucosal gland ducts and the superficial epithelium of human large airways [75][77][79]. Although reported over a decade ago, these cells had not been studied until very recently (see next paragraph).

4.2. New Insights with Single-Cell Transcriptional Profiling

The latest technological advancements have allowed for transcriptome analyses at the single-cell level, resulting in the identification of new cell types in multiple organs within a wide range of species [80][81][82]. Single-cell transcriptional profiling studies, focusing on human and mice large airways, identified CFTR-rich epithelial cells, called pulmonary ionocytes [83][84], as the potential “hot cells” identified in the early morphometric studies. A compelling study showed a positive association between the number of pulmonary ionocytes and the activity of CFTR in HBE cell cultures, despite ionocyte numbers accounting for a small fraction (<1–2%) of the total cells [84]. In contrast, a different study demonstrated no loss of CFTR-mediated Cl secretory function in HBE cells in which ionocytes were genetically depleted by CRISPR/Cas9 technology targeting FOXI1, a key transcription factor specific to ionocyte lineage [72]. Thus far, it remains unclear whether pulmonary ionocytes contribute significantly to Cl and HCO3 secretion in the lungs. Data from mouse trachea revealed that 54.4% of Cftr transcripts were concentrated in ionocytes, despite this cell type comprising <1% of the total cells, suggesting that ionocytes and at least another cell type play a role in CFTR expression and function in the murine airways [83]. More recently, comprehensive single-cell transcriptome analyses performed on human lungs identified secretory club cells as the most common cell type expressing CFTR transcripts in the large and small airway regions [85]. Intraregional characterization of CFTR and FOXI1 transcripts along the proximal–distal axis of the lungs revealed discrepancies between robust CFTR expression and trivial numbers of ionocytes in the distal airway regions, suggesting that secretory club cells may convert to the dominant cell type expressing functional CFTR in the small airways. Another study using single-cell transcriptional profiling revealed that the secretory cell is the most common cell type expressing CFTR in the non-CF superficial epithelium, which remains true in the CF trachea, despite significant dynamic transcriptome alterations in CF [86]. In CF tracheas, specific changes in transcriptional profiles of secretory cells included overactivated secretory functions, exhausted metabolic profiles, and elevated endoplasmic reticulum stress pathways. The persistence of CFTR transcripts in CF secretory cells indicates that this cell type could be an attractive therapeutic target for CFTR gene editing and/or transfer in the more accessible airway surface epithelium of the trachea. Further efforts are required to fully understand CFTR function in specific cell types (i.e., ionocytes vs secretory cells) in the different regions of the lung.

4.3. The Role of Secretory Cells in the Small Airways

Airway secretory club cells have been thoroughly studied for their capacity to secrete host innate defense proteins (e.g., CCSP, LTF, and SLPI) and metabolize inhaled xenobiotics (e.g., p-xylene, benzo(a)pyrene, and ethylnitrosourea) that can be cytotoxic and/or carcinogenic [87][88][89][90]. One of the more prominent characteristics of this cell type is the plasticity of these cells that act as progenitor cells for specialized airway epithelial cells, e.g., ciliated and goblet cells. As suggested by recent transcriptomic data, secretory club cells express ion channel genes, including CFTR and SCNN1, and, therefore, maintain the balance of salt and water in the small airways [85]. However, only a few studies have characterized ion transport activity in the small airways and the potential roles for secretory club cells in controlling ion and water movements across the epithelium in the distal lung [91][92][93][94]. Blouquit et al. demonstrated that disease-controlled small airways exhibit active ENaC-mediated Na+ absorption and CFTR-mediated Cl secretion, while CF small airways failed to modulate ion fluxes and maintain ASL homeostasis [93]. Van Scott et al. purified rabbit club cells to confirm the presence of basal Na+ absorption and inducible Cl secretion in this cell type [95]. More recently, Kulaksiz et al. confirmed expression, localization, and function of CFTR in club cells of rat and human small airways [96]. Since both MUC5B and CFTR localize to the secretory club cells, and since secretory club cells provide host defense mechanisms, this cell type should be considered as a multi-dimensional mucin/ion/host defense regulatory cell that plays a crucial role in regulating mucosal defense and airway clearance, particularly in the small airways. Cell type-specific mechanistic studies will be important to determine how a single cell type regulates multiple secretory functions.


  1. Ehre, C.; Ridley, C.; Thornton, D.J. Cystic fibrosis: An inherited disease affecting mucin-producing organs. Int. J. Biochem. Cell Biol. 2014, 52, 136–145.
  2. Morrison, C.B.; Markovetz, M.R.; Ehre, C. Mucus, mucins, and cystic fibrosis. Pediatric Pulmonol. 2019, 54 (Suppl. 3), S84–S96.
  3. Boucher, R.C. Evidence for airway surface dehydration as the initiating event in CF airway disease. J. Intern. Med. 2007, 261, 5–16.
  4. Quinton, P.M. Cystic fibrosis: Impaired bicarbonate secretion and mucoviscidosis. Lancet 2008, 372, 415–417.
  5. Andersen, D.H. Cystic Fibrosis of the Pancreas and Its Relation to Celiac Disease: A Clinical and Pathologic Study. Am. J. Dis. Child. 1938, 56, 344–399.
  6. Navarro, S. Historical compilation of cystic fibrosis. Gastroenterol. Hepatol. 2016, 39, 36–42.
  7. Pinto, M.C.; Silva, I.A.L.; Figueira, M.F.; Amaral, M.D.; Lopes-Pacheco, M. Pharmacological Modulation of Ion Channels for the Treatment of Cystic Fibrosis. J. Exp. Pharmacol. 2021, 13, 693–723.
  8. Davis, P.B. Cystic fibrosis since 1938. Am. J. Respir. Crit. Care Med. 2006, 173, 475–482.
  9. Van Goor, F.; Hadida, S.; Grootenhuis, P.D.J.; Burton, B.; Cao, D.; Neuberger, T.; Turnbull, A.; Singh, A.; Joubran, J.; Hazlewood, A.; et al. Rescue of CF airway epithelial cell function in vitro by a CFTR potentiator, VX-770. Proc. Natl. Acad. Sci. USA 2009, 106, 18825–18830.
  10. Accurso, F.J.; Rowe, S.M.; Clancy, J.P.; Boyle, M.P.; Dunitz, J.M.; Durie, P.R.; Sagel, S.D.; Hornick, D.B.; Konstan, M.W.; Donaldson, S.H.; et al. Effect of VX-770 in persons with cystic fibrosis and the G551D-CFTR mutation. N. Engl. J. Med. 2010, 363, 1991–2003.
  11. Ramsey, B.W.; Davies, J.; McElvaney, N.G.; Tullis, E.; Bell, S.C.; Dřevínek, P.; Griese, M.; McKone, E.F.; Wainwright, C.E.; Konstan, M.W.; et al. A CFTR potentiator in patients with cystic fibrosis and the G551D mutation. N. Engl. J. Med. 2011, 365, 1663–1672.
  12. Skilton, M.; Krishan, A.; Patel, S.; Sinha, I.P.; Southern, K.W.; Cochrane Cystic Fibrosis and Genetic Disorders Group. Potentiators (specific therapies for class III and IV mutations) for cystic fibrosis. Cochrane Database Syst. Rev. 2019, 2019, CD009841.
  13. Clancy, J.P.; Rowe, S.M.; Accurso, F.J.; Aitken, M.L.; Amin, R.S.; Ashlock, M.A.; Ballmann, M.; Boyle, M.P.; Bronsveld, I.; Campbell, P.W.; et al. Results of a phase IIa study of VX-809, an investigational CFTR corrector compound, in subjects with cystic fibrosis homozygous for the F508del-CFTR mutation. Thorax 2012, 67, 12–18.
  14. Gramegna, A.; Contarini, M.; Aliberti, S.; Casciaro, R.; Blasi, F.; Castellani, C. From Ivacaftor to Triple Combination: A Systematic Review of Efficacy and Safety of CFTR Modulators in People with Cystic Fibrosis. Int. J. Mol. Sci. 2020, 21, 5882.
  15. Donaldson, S.H.; Pilewski, J.M.; Griese, M.; Cooke, J.; Viswanathan, L.; Tullis, E.; Davies, J.C.; Lekstrom-Himes, J.A.; Wang, L.T.; VX11-661-101 Study Group. Tezacaftor/Ivacaftor in Subjects with Cystic Fibrosis and F508del/F508del-CFTR or F508del/G551D-CFTR. Am. J. Respir. Crit. Care Med. 2018, 197, 214–224.
  16. Keating, D.; Marigowda, G.; Burr, L.D.; Daines, C.; Mall, M.A.; McKone, E.F.; Ramsey, B.W.; Rowe, S.M.; Sass, L.A.; Tullis, E.; et al. VX-445–Tezacaftor–Ivacaftor in Patients with Cystic Fibrosis and One or Two Phe508del Alleles. N. Engl. J. Med. 2018, 379, 1612–1620.
  17. Middleton, P.G.; Mall, M.A.; Dřevínek, P.; Lands, L.C.; McKone, E.F.; Polineni, D.; Ramsey, B.W.; Taylor-Cousar, J.L.; Tullis, E.; Vermeulen, F.; et al. Elexacaftor-Tezacaftor-Ivacaftor for Cystic Fibrosis with a Single Phe508del Allele. N. Engl. J. Med. 2019, 381, 1809–1819.
  18. Heijerman, H.G.M.; McKone, E.F.; Downey, D.G.; Van Braeckel, E.; Rowe, S.M.; Tullis, E.; Mall, M.A.; Welter, J.J.; Ramsey, B.W.; McKee, C.M.; et al. Efficacy and safety of the elexacaftor plus tezacaftor plus ivacaftor combination regimen in people with cystic fibrosis homozygous for the F508del mutation: A double-blind, randomised, phase 3 trial. Lancet 2019, 394, 1940–1948.
  19. Esther, C.R., Jr.; Muhlebach, M.S.; Ehre, C.; Hill, D.B.; Wolfgang, M.C.; Kesimer, M.; Ramsey, K.A.; Markovetz, M.R.; Garbarine, I.C.; Forest, M.G.; et al. Mucus accumulation in the lungs precedes structural changes and infection in children with cystic fibrosis. Sci. Transl. Med. 2019, 11, eaav3488.
  20. Henderson, A.G.; Ehre, C.; Button, B.; Abdullah, L.H.; Cai, L.-H.; Leigh, M.W.; DeMaria, G.C.; Matsui, H.; Donaldson, S.H.; Davis, C.W.; et al. Cystic fibrosis airway secretions exhibit mucin hyperconcentration and increased osmotic pressure. J. Clin. Investig. 2014, 124, 3047–3060.
  21. Tang, X.X.; Ostedgaard, L.S.; Hoegger, M.J.; Moninger, T.O.; Karp, P.H.; McMenimen, J.D.; Choudhury, B.; Varki, A.; Stoltz, D.A.; Welsh, M.J. Acidic pH increases airway surface liquid viscosity in cystic fibrosis. J. Clin. Investig. 2016, 126, 879–891.
  22. Shah, V.S.; Meyerholz, D.K.; Tang, X.X.; Reznikov, L.; Abou Alaiwa, M.; Ernst, S.E.; Karp, P.H.; Wohlford-Lenane, C.L.; Heilmann, K.P.; Leidinger, M.R.; et al. Airway acidification initiates host defense abnormalities in cystic fibrosis mice. Science 2016, 351, 503–507.
  23. Schultz, A.; Puvvadi, R.; Borisov, S.M.; Shaw, N.C.; Klimant, I.; Berry, L.J.; Montgomery, S.T.; Nguyen, T.; Kreda, S.M.; Kicic, A.; et al. Airway surface liquid pH is not acidic in children with cystic fibrosis. Nat. Commun. 2017, 8, 1409.
  24. Weibel, E.R. Chapter II–Organization of the Human Lung. In Morphometry of the Human Lung; Academic Press: New York, NY, USA, 1963; pp. 4–9.
  25. Weibel, E.R. Morphometry of the human lung: The state of the art after two decades. Bull. Eur. Physiopathol. Respir. 1979, 15, 999–1013.
  26. Spencer, H. Morphometry of the Human Lung. J. Anat. 1964, 98, 457.
  27. Widdicombe, J.H.; Wine, J.J. Airway Gland Structure and Function. Physiol. Rev. 2015, 95, 1241–1319.
  28. Wickstrom, C.; Davies, J.; Eriksen, G.V.; Veerman, E.C.I.; Carlstedt, I. MUC5B is a major gel-forming, oligomeric mucin from human salivary gland, respiratory tract and endocervix: Identification of glycoforms and C-terminal cleavage. Biochem. J. 1998, 334, 685–693.
  29. Jeong, J.H.; Joo, N.S.; Hwang, P.H.; Wine, J.J. Mucociliary clearance and submucosal gland secretion in the ex vivo ferret trachea. Am. J. Physiol. Lung Cell. Mol. Physiol. 2014, 307, L83–L93.
  30. Ermund, A.; Meiss, L.N.; Rodriguez-Pineiro, A.M.; Bähr, A.; Nilsson, H.E.; Trillo-Muyo, S.; Ridley, C.; Thornton, D.J.; Wine, J.J.; Hebert, H.; et al. The normal trachea is cleaned by MUC5B mucin bundles from the submucosal glands coated with the MUC5AC mucin. Biochem. Biophys. Res. Commun. 2017, 492, 331–337.
  31. Kato, T.; Radicioni, G.; Papanikolas, M.J.; Stoychev, G.V.; Markovetz, M.R.; Aoki, K.; Porterfield, M.; Okuda, K.; Cardenas, S.M.B.; Gilmore, R.C.; et al. Mucus concentration–Dependent biophysical abnormalities unify submucosal gland and superficial airway dysfunction in cystic fibrosis. Sci. Adv. 2022, 8, eabm9718.
  32. Button, B.; Cai, L.-H.; Ehre, C.; Kesimer, M.; Hill, D.B.; Sheehan, J.K.; Boucher, R.C.; Rubinstein, M. A Periciliary Brush Promotes the Lung Health by Separating the Mucus Layer from Airway Epithelia. Science 2012, 337, 937–941.
  33. Burgel, P.-R.; Montani, D.; Danel, C.; Dusser, D.J.; Nadel, J.A. A morphometric study of mucins and small airway plugging in cystic fibrosis. Thorax 2007, 62, 153–161.
  34. Zuelzer, W.W.; Newton, W.A., Jr. The pathogenesis of fibrocystic disease of the pancreas; A study of 36 cases with special reference to the pulmonary lesions. Pediatrics 1949, 4, 53–69.
  35. Sheppard, M.N. The pathology of cystic fibrosis. In Cystic Fibrosis; Hodson, M.E., Geddes, D.M., Eds.; Chapman and Hall: London, UK, 1995; pp. 131–149.
  36. Khan, T.Z.; Wagener, J.S.; Bost, T.; Martinez, J.; Accurso, F.J.; Riches, D.W. Early pulmonary inflammation in infants with cystic fibrosis. Am. J. Respir. Crit. Care Med. 1995, 151, 1075–1082.
  37. Noah, T.L.; Murphy, P.C.; Alink, J.J.; Leigh, M.W.; Hull, W.M.; Stahlman, M.T.; Whitsett, J.A. Bronchoalveolar Lavage Fluid Surfactant Protein-A and Surfactant Protein-D Are Inversely Related to Inflammation in Early Cystic Fibrosis. Am. J. Respir. Crit. Care Med. 2003, 168, 685–691.
  38. Mellins, R.B. The Site of Airway Obstruction in Cystic Fibrosis. Pediatrics 1969, 44, 315–318.
  39. Ranganathan, S.C.; Stocks, J.; Dezateux, C.; Bush, A.; Wade, A.; Carr, S.; Castle, R.; Dinwiddie, R.; Hoo, A.-F.; Lum, S.; et al. The Evolution of Airway Function in Early Childhood Following Clinical Diagnosis of Cystic Fibrosis. Am. J. Respir. Crit. Care Med. 2004, 169, 928–933.
  40. Donaldson, S.H.; Bennett, W.D.; Zeman, K.L.; Knowles, M.R.; Tarran, R.; Boucher, R.C. Mucus Clearance and Lung Function in Cystic Fibrosis with Hypertonic Saline. N. Engl. J. Med. 2006, 354, 241–250.
  41. Tiddens, H.A.; Donaldson, S.H.; Rosenfeld, M.; Paré, P.D. Cystic fibrosis lung disease starts in the small airways: Can we treat it more effectively? Pediatric Pulmonol. 2010, 45, 107–117.
  42. Bennett, W.D.; Olivier, K.N.; Zeman, K.L.; Hohneker, K.W.; Boucher, R.C.; Knowles, M.R. Effect of uridine 5′-triphosphate plus amiloride on mucociliary clearance in adult cystic fibrosis. Am. J. Respir. Crit. Care Med. 1996, 153, 1796–1801.
  43. Schwiebert, E.M.; Benos, D.J.; Egan, M.E.; Stutts, M.J.; Guggino, W.B. CFTR Is a Conductance Regulator as well as a Chloride Channel. Physiol. Rev. 1999, 79, S145–S166.
  44. Kunzelmann, K. The cystic fibrosis transmembrane conductance regulator and its function in epithelial transport. Rev. Physiol. Biochem. Pharmacol. 1999, 137, 1–70.
  45. Boon, M.; Verleden, S.E.; Bosch, B.; Lammertyn, E.J.; McDonough, J.E.; Mai, C.; Verschakelen, J.; Kemner-van de Corput, M.; Tiddens, H.A.; Proesmans, M.; et al. Morphometric Analysis of Explant Lungs in Cystic Fibrosis. Am. J. Respir. Crit. Care Med. 2016, 193, 516–526.
  46. Regnis, J.A.; Zeman, K.L.; Noone, P.G.; Knowles, M.R.; Bennett, W.D. Prolonged airway retention of insoluble particles in cystic fibrosis versus primary ciliary dyskinesia. Exp. Lung Res. 2000, 26, 149–162.
  47. Button, B.; Goodell, H.P.; Atieh, E.; Chen, Y.-C.; Williams, R.; Shenoy, S.; Lackey, E.; Shenkute, N.T.; Cai, L.-H.; Dennis, R.G.; et al. Roles of mucus adhesion and cohesion in cough clearance. Proc. Natl. Acad. Sci. USA 2018, 115, 12501–12506.
  48. Livraghi-Butrico, A.; Grubb, B.R.; Wilkinson, K.J.; Volmer, A.S.; Burns, K.A.; Evans, C.M.; O’Neal, W.K.; Boucher, R.C. Contribution of mucus concentration and secreted mucins Muc5ac and Muc5b to the pathogenesis of muco-obstructive lung disease. Mucosal Immunol. 2017, 10, 395–407.
  49. Serafini, S.M.; Michaelson, E.D. Length and distribution of cilia in human and canine airways. Bull. Eur. Physiopathol. Respir. 1977, 13, 551–559.
  50. Murthy, P.K.L.; Sontake, V.; Tata, A.; Kobayashi, Y.; Macadlo, L.; Okuda, K.; Conchola, A.S.; Nakano, S.; Gregory, S.; Miller, L.A.; et al. Human distal lung maps and lineage hierarchies reveal a bipotent progenitor. Nature 2022, 604, 111–119.
  51. Smith, D.J.; Gaffney, E.A.; Blake, J.R. Modelling mucociliary clearance. Respir. Physiol. Neurobiol. 2008, 163, 178–188.
  52. Kesimer, M.; Kirkham, S.; Pickles, R.J.; Henderson, A.G.; Alexis, N.E.; Demaria, G.; Knight, D.; Thornton, D.J.; Sheehan, J.K. Tracheobronchial air-liquid interface cell culture: A model for innate mucosal defense of the upper airways? Am. J. Physiol. Lung Cell. Mol. Physiol. 2009, 296, L92–L100.
  53. Kesimer, M.; Ehre, C.; Burns, K.A.; Davis, C.W.; Sheehan, J.K.; Pickles, R.J. Molecular organization of the mucins and glycocalyx underlying mucus transport over mucosal surfaces of the airways. Mucosal Immunol. 2013, 6, 379–392.
  54. Buisine, M.-P.; Devisme, L.; Copin, M.-C.; Durand-Réville, M.; Gosselin, B.; Aubert, J.-P.; Porchet, N. Developmental Mucin Gene Expression in the Human Respiratory Tract. Am. J. Respir. Cell Mol. Biol. 1999, 20, 209–218.
  55. Kirkham, S.; Sheehan, J.K.; Knight, D.; Richardson, P.S.; Thornton, D.J. Heterogeneity of airways mucus: Variations in the amounts and glycoforms of the major oligomeric mucins MUC5AC and MUC5B. Biochem. J. 2002, 361, 537–546.
  56. Radicioni, G.; Ceppe, A.; Ford, A.A.; Alexis, N.E.; Barr, R.G.; Bleecker, E.R.; Christenson, S.A.; Cooper, C.B.; Han, M.K.; Hansel, N.N.; et al. Airway mucin MUC5AC and MUC5B concentrations and the initiation and progression of chronic obstructive pulmonary disease: An analysis of the SPIROMICS cohort. Lancet. Respir. Med. 2021, 9, 1241–1254.
  57. Kesimer, M.; Ford, A.A.; Ceppe, A.; Radicioni, G.; Cao, R.; Davis, C.W.; Doerschuk, C.M.; Alexis, N.E.; Anderson, W.H.; Henderson, A.G.; et al. Airway Mucin Concentration as a Marker of Chronic Bronchitis. N. Engl. J. Med. 2017, 377, 911–922.
  58. Henke, M.O.; John, G.; Germann, M.; Lindemann, H.; Rubin, B.K. MUC5AC and MUC5B mucins increase in cystic fibrosis airway secretions during pulmonary exacerbation. Am. J. Respir. Crit. Care Med. 2007, 175, 816–821.
  59. Ramsey, K.A.; Chen, A.C.H.; Radicioni, G.; Lourie, R.; Martin, M.; Broomfield, A.; Sheng, Y.H.; Hasnain, S.Z.; Radford-Smith, G.; Simms, L.A.; et al. Airway mucus hyperconcentration in non–cystic fibrosis bronchiectasis. Am. J. Respir. Crit. Care Med. 2020, 201, 661–670.
  60. Anderson, W.H.; Coakley, R.D.; Button, B.; Henderson, A.G.; Zeman, K.L.; Alexis, N.E.; Peden, D.B.; Lazarowski, E.R.; Davis, C.W.; Bailey, S.; et al. The Relationship of Mucus Concentration (Hydration) to Mucus Osmotic Pressure and Transport in Chronic Bronchitis. Am. J. Respir. Crit. Care Med. 2015, 192, 182–190.
  61. Welsh, K.G.; Rousseau, K.; Fisher, G.; Bonser, L.R.; Bradding, P.; Brightling, C.E.; Thornton, D.J.; Gaillard, E.A. MUC5AC and a Glycosylated Variant of MUC5B Alter Mucin Composition in Children with Acute Asthma. Chest 2017, 152, 771–779.
  62. Bonser, L.R.; Zlock, L.; Finkbeiner, W.; Erle, D.J. Epithelial tethering of MUC5AC-rich mucus impairs mucociliary transport in asthma. J. Clin. Investig. 2016, 126, 2367–2371.
  63. Roy, M.G.; Livraghi-Butrico, A.; Fletcher, A.A.; McElwee, M.M.; Evans, S.E.; Boerner, R.M.; Alexander, S.N.; Bellinghausen, L.K.; Song, A.S.; Petrova, Y.M.; et al. Muc5b is required for airway defence. Nature 2014, 505, 412–416.
  64. Borthwick, D.W.; West, J.D.; Keighren, M.A.; Flockhart, J.H.; Innes, B.A.; Dorin, J.R. Murine Submucosal Glands Are Clonally Derived and Show a Cystic Fibrosis Gene–Dependent Distribution Pattern. Am. J. Respir. Cell Mol. Biol. 1999, 20, 1181–1189.
  65. Morrison, C.B.; Edwards, C.E.; Shaffer, K.M.; Araba, K.C.; Wykoff, J.A.; Williams, D.R.; Asakura, T.; Dang, H.; Morton, L.C.; Gilmore, R.C.; et al. SARS-CoV-2 infection of airway cells causes intense viral and cell shedding, two spreading mechanisms affected by IL-13. Proc. Natl. Acad. Sci. USA 2022, 119, e2119680119.
  66. Carpenter, J.; Wang, Y.; Gupta, R.; Li, Y.; Haridass, P.; Subramani, D.B.; Reidel, B.; Morton, L.; Ridley, C.; O’Neal, W.K.; et al. Assembly and organization of the N-terminal region of mucin MUC5AC: Indications for structural and functional distinction from MUC5B. Proc. Natl. Acad. Sci. USA 2021, 118, e2104490118.
  67. Ehre, C.; Worthington, E.N.; Liesman, R.M.; Grubb, B.R.; Barbier, D.; O’Neal, W.K.; Sallenave, J.-M.; Pickles, R.J.; Boucher, R.C. Overexpressing mouse model demonstrates the protective role of Muc5ac in the lungs. Proc. Natl. Acad. Sci. USA 2012, 109, 16528–16533.
  68. Hasnain, S.; Wang, H.; Ghia, J.-E.; Haq, N.; Deng, Y.; Velcich, A.; Grencis, R.K.; Thornton, D.J.; Khan, W.I. Mucin Gene Deficiency in Mice Impairs Host Resistance to an Enteric Parasitic Infection. Gastroenterology 2010, 138, 1763–1771.e5.
  69. Kreda, S.M.; Davis, C.W.; Rose, M.C. CFTR, Mucins, and Mucus Obstruction in Cystic Fibrosis. Cold Spring Harb. Perspect. Med. 2012, 2, a009589.
  70. Zhu, Y.; Ehre, C.; Abdullah, L.H.; Sheehan, J.K.; Roy, M.; Evans, C.M.; Dickey, B.F.; Davis, C.W. Munc13-2−/− baseline secretion defect reveals source of oligomeric mucins in mouse airways. J. Physiol. 2008, 586, 1977–1992.
  71. Evans, C.M.; Williams, O.W.; Tuvim, M.J.; Nigam, R.; Mixides, G.P.; Blackburn, M.R.; DeMayo, F.J.; Burns, A.R.; Smith, C.; Reynolds, S.D.; et al. Mucin Is Produced by Clara Cells in the Proximal Airways of Antigen-Challenged Mice. Am. J. Respir. Cell Mol. Biol. 2004, 31, 382–394.
  72. Goldfarbmuren, K.C.; Jackson, N.D.; Sajuthi, S.P.; Dyjack, N.; Li, K.S.; Rios, C.L.; Plender, E.G.; Montgomery, M.T.; Everman, J.L.; Bratcher, P.E.; et al. Dissecting the cellular specificity of smoking effects and reconstructing lineages in the human airway epithelium. Nat. Commun. 2020, 11, 2485.
  73. Okuda, K.; Chen, G.; Subramani, D.B.; Wolf, M.; Gilmore, R.C.; Kato, T.; Radicioni, G.; Kesimer, M.; Chua, M.; Dang, H.; et al. Localization of Secretory Mucins MUC5AC and MUC5B in Normal/Healthy Human Airways. Am. J. Respir. Crit. Care Med. 2019, 199, 715–727.
  74. Chen, G.; Sun, L.; Kato, T.; Okuda, K.; Martino, M.B.; Abzhanova, A.; Lin, J.M.; Gilmore, R.C.; Batson, B.D.; O’Neal, Y.K.; et al. IL-1β dominates the promucin secretory cytokine profile in cystic fibrosis. J. Clin. Investig. 2019, 129, 4433–4450.
  75. Engelhardt, J.F.; Yankaskas, J.R.; Ernst, S.A.; Yang, Y.; Marino, C.R.; Boucher, R.C.; Cohn, J.A.; Wilson, J.M. Submucosal glands are the predominant site of CFTR expression in the human bronchus. Nat. Genet. 1992, 2, 240–248.
  76. Engelhardt, J.; Zepeda, M.; Cohn, J.A.; Yankaskas, J.R.; Wilson, J. Expression of the cystic fibrosis gene in adult human lung. J. Clin. Investig. 1994, 93, 737–749.
  77. Kreda, S.M.; Mall, M.; Mengos, A.; Rochelle, L.; Yankaskas, J.; Riordan, J.R.; Boucher, R.C. Characterization of wild-type and ∆F508 cystic fibrosis transmembrane regulator in human respiratory epithelia. Mol. Biol. Cell 2005, 16, 2154–2167.
  78. Sato, Y.; Mustafina, K.R.; Luo, Y.; Martini, C.; Thomas, D.Y.; Wiseman, P.W.; Hanrahan, J.W. Nonspecific binding of common anti-CFTR antibodies in ciliated cells of human airway epithelium. Sci. Rep. 2021, 11, 23256.
  79. Kälin, N.; Claaß, A.; Sommer, M.; Puchelle, E.; Tümmler, B. ∆F508 CFTR protein expression in tissues from patients with cystic fibrosis. J. Clin. Investig. 1999, 103, 1379–1389.
  80. Treutlein, B.; Brownfield, D.; Wu, A.; Neff, N.F.; Mantalas, G.L.; Espinoza, F.; Desai, T.J.; Krasnow, M.A.; Quake, S.R. Reconstructing lineage hierarchies of the distal lung epithelium using single-cell RNA-seq. Nature 2014, 509, 371–375.
  81. Wagner, D.E.; Weinreb, C.; Collins, Z.M.; Briggs, J.A.; Megason, S.G.; Klein, A.M. Single-cell mapping of gene expression landscapes and lineage in the zebrafish embryo. Science 2018, 360, 981–987.
  82. Nagendran, M.; Riordan, D.P.; Harbury, P.B.; Desai, T.J. Automated cell-type classification in intact tissues by single-cell molecular profiling. eLife 2018, 7, e30510.
  83. Montoro, D.T.; Haber, A.L.; Biton, M.; Vinarsky, V.; Lin, B.; Birket, S.E.; Yuan, F.; Chen, S.; Leung, H.M.; Villoria, J.; et al. A revised airway epithelial hierarchy includes CFTR-expressing ionocytes. Nature 2018, 560, 319–324.
  84. Plasschaert, L.W.; Žilionis, R.; Choo-Wing, R.; Savova, V.; Knehr, J.; Roma, G.; Klein, A.M.; Jaffe, A.B. A single-cell atlas of the airway epithelium reveals the CFTR-rich pulmonary ionocyte. Nature 2018, 560, 377–381.
  85. Okuda, K.; Dang, H.; Kobayashi, Y.; Carraro, G.; Nakano, S.; Chen, G.; Kato, T.; Asakura, T.; Gilmore, R.C.; Morton, L.C.; et al. Secretory Cells Dominate Airway CFTR Expression and Function in Human Airway Superficial Epithelia. Am. J. Respir. Crit. Care Med. 2021, 203, 1275–1289.
  86. Carraro, G.; Langerman, J.; Sabri, S.; Lorenzana, Z.; Purkayastha, A.; Zhang, G.; Konda, B.; Aros, C.J.; Calvert, B.A.; Szymaniak, A.; et al. Transcriptional analysis of cystic fibrosis airways at single-cell resolution reveals altered epithelial cell states and composition. Nat. Med. 2021, 27, 806–814.
  87. Tokita, E.; Tanabe, T.; Asano, K.; Suzaki, H.; Rubin, B.K. Club cell 10-kDa protein attenuates airway mucus hypersecretion and inflammation. Eur. Respir. J. 2014, 44, 1002–1010.
  88. Gamez, A.S.; Gras, D.; Petit, A.; Knabe, L.; Molinari, N.; Vachier, I.; Chanez, P.; Bourdin, A. Supplementing defect in club cell secretory protein attenuates airway inflammation in COPD. Chest 2015, 147, 1467–1476.
  89. Plopper, C.G.; Cranz, D.L.; Kemp, L.; Serabjit-Singh, C.J.; Philpot, R.M. Immunohistochemical Demonstration of Cytochrome P-450 Monooxygenase in Clara Cells throughout the Tracheobronchial Airways of the Rabbit. Exp. Lung Res. 1987, 13, 59–68.
  90. Mango, G.W.; Johnston, C.J.; Reynolds, S.D.; Finkelstein, J.N.; Plopper, C.G.; Stripp, B.R. Clara cell secretory protein deficiency increases oxidant stress response in conducting airways. Am. J. Physiol. Cell. Mol. Physiol. 1998, 275, L348–L356.
  91. Shamsuddin, A.K.M.; Quinton, P.M. Native Small Airways Secrete Bicarbonate. Am. J. Respir. Cell Mol. Biol. 2014, 50, 796–804.
  92. Shamsuddin, A.K.M.; Quinton, P.M. Concurrent absorption and secretion of airway surface liquids and bicarbonate secretion in human bronchioles. Am. J. Physiol. Lung Cell. Mol. Physiol. 2019, 316, L953–L960.
  93. Blouquit, S.; Regnier, A.; Dannhoffer, L.; Fermanian, C.; Naline, E.; Boucher, R.; Chinet, T. Ion and Fluid Transport Properties of Small Airways in Cystic Fibrosis. Am. J. Respir. Crit. Care Med. 2006, 174, 299–305.
  94. Li, X.; Tang, X.X.; Buonfiglio, L.G.V.; Comellas, A.P.; Thornell, I.M.; Ramachandran, S.; Karp, P.H.; Taft, P.J.; Sheets, K.; Abou Alaiwa, M.H.; et al. Electrolyte transport properties in distal small airways from cystic fibrosis pigs with implications for host defense. Am. J. Physiol. Lung Cell. Mol. Physiol. 2016, 310, L670–L679.
  95. Van Scott, M.R.; Hester, S.; Boucher, R.C. Ion transport by rabbit nonciliated bronchiolar epithelial cells (Clara cells) in culture. Proc. Natl. Acad. Sci. USA 1987, 84, 5496–5500.
  96. Kulaksiz, H.; Schmid, A.; Hönscheid, M.; Ramaswamy, A.; Cetin, Y. Clara cell impact in air-side activation of CFTR in small pulmonary airways. Proc. Natl. Acad. Sci. USA 2002, 99, 6796–6801.
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